MODULATION OF THE IMMUNE RESPONSE TO INFLUENZA VIRUS BY THE
CANNABINOID RECEPTORS 1 AND 2
By
Peer Willem Frederik Karmaus

A DISSERTATION
Submitted to
Michigan State University
In partial fulfillment of the requirements
For the degree of
DOCTOR OF PHILOSOPHY
Cell and Molecular Biology – Environmental Toxicology
2011

ABSTRACT
MODULATION OF THE IMMUNE RESPONSE TO INFLUENZA VIRUS BY THE
CANNABINOID RECEPTORS 1 AND 2
By
Peer Willem Frederik Karmaus
Cannabinoids are bioactive signaling molecules exerting several effects, including
modulation of immune function. One of the agents studied for its immunosuppressive
activity, Δ9-tetrahydrocannabinol (Δ9-THC), is thought to exert part of its activity on
immune cells via interaction with cannabinoid receptors 1 (CB1) and 2 (CB2). In
previous studies Δ9-THC caused an increase in steady state viral hemagglutinin RNA
levels, suggesting decreased viral clearance due to compromised immune function. In
addition, in CB1 and CB2 double knock out (CB1-/-CB2-/-) mice, a reduced viral burden
and greater magnitude in the immune response were observed, suggesting a role for CB1
and/or CB2 in regulating immune function. The identity of the functionally dysregulated
immune population(s) as a result of CB1 and CB2 deletion were unknown. In addition,
the effect of Δ9-THC treatment on the cellular immune response to influenza remained to
be elucidated. Overall, this dissertation project was designed to test the hypothesis that
Δ9-THC suppresses immune responses generated by influenza virus by perturbing an
endogenous signaling pathway relying in part on the presence of CB1 and/or CB2. In the
present studies, Δ9-THC suppressed cytotoxic T lymphocyte (CTL) function independent
of CB1 and CB2, suggesting an alternate mechanism for immune modulation by Δ9-THC
in these cells. Second, a kinetic study of influenza infection demonstrated greater immune
response magnitude in CB1-/-CB2-/- compared to WT mice. This was evidenced by

elevated steady-state mRNA levels for genes involved in the inflammatory response,
earlier T cell activation, greater percentages of cytokine-secreting effector cells, and
increased immunopathology in lung tissue. These events likely lead to lower viral burden
at the expense of greater tissue damage in CB1-/-CB2-/- mice compared to WT. These
differences observed in vivo were not reproduced by stimulation of T cells in vitro,
suggesting differences in antigen presenting cells (APC), which elicit effector cells.
Dendritic cells (DC) generated from bone marrow (bmDC) and alveolar macrophages
(AM) isolated from CB1-/-CB2-/- mice had greater maturation with and without Toll-like
receptor (TLR) stimulation compared to WT mice and bmDC elicited antigen-specific T
cells without the need for a maturation stimulus. Third, focusing on the peak day of the
inflammatory response induced by influenza virus at day 3, the immune response of APC
was characterized in WT and CB1-/-CB2-/- mice in the presence of Δ9-THC. Δ9-THC
suppressed influx of APC, including conventional DC (cDC), plasmacytoid DC (pDC),
and AM into the lung after influenza infection in WT only. While there was no change in
maturation of lung-isolated APC due to Δ9-THC treatment, only mature APC reach the
lung, thus Δ9-THC might have affected maturation of APC prior to migration to the lung.
Indeed, Δ9-THC suppressed DC maturation and ability to elicit antigen-specific T cells in
vitro. Taken together, Δ9-THC impairs the immune response to influenza, in part, by
suppressing APC function in a CB1 and/or CB2 dependent manner, while suppressing
CTL elicitation in a CB1 and CB2 independent manner. Also, the presence of CB1 and/or
CB2 on APC prevents exacerbated immune responses after influenza infection. These
studies suggest that CB1 and/or CB2 maintain immune homeostasis and prevent
excessive reactivity after immune stimulation.

Copyright by
PEER WILLEM FREDERIK KARMAUS
2011

ACKNOWLEDGEMENTS

First, I would like to acknowledge my mentor Dr. Norbert Kaminski for his continous
support and excellent mentorship during my years as a graduate student in his laboratory. My
achievements were only made possible as a result of his efforts.

Also I would like to thank my guidance committee for sharing their helpful opinions and
insights during discussions of my dissertation project. Specifically (in alphabetical order by last
name) I thank Dr. Patricia Ganey for jumpstarting and nurturing my interest in the interaction of
xenobiotics with the immune system; Dr. Jack Harkema for discussions of lung biology,
performing histopathological analyses, and allowing me to present my findings to a larger
audience during the Respiratory Research Initiative meetings; Dr. Barbara Kaplan for sharing her
expertise in cell and molecular immunology, her sense of humor, and 11:45 with me; Dr. Richard
Schwartz for sharing his keen sense of cellular signaling pathways and interactions on an
administrative level on several university committees.

Last but not least, I would like to acknowledge my labmates, my friends, my family, and
my girlfriend Agnes for their endless support and making every day more enjoyable.

v

TABLE OF CONTENTS

LIST OF TABLES.............................................................................................................. X
LIST OF FIGURES ...........................................................................................................XI
LIST OF ABBREVIATIONS ........................................................................................ XIV
INTRODUCTION ............................................................................................................... 1
I. Respiratory Tract and Lung Associated Immune Defenses against Pathogens ............... 1
A) Respiratory Tract Anatomy and Function .......................................................... 1
B) Differences in the Respiratory Tract between Humans and Rodents ................. 1
C) Lung Associated Lymphoid Organs ................................................................... 2
D) Mechanical Defenses .......................................................................................... 2
E) Respiratory Epithelial Cells ................................................................................ 3
F) Immune Functions of Type II and Clara Cells .................................................... 4
G) Expression of Pathogen Associated Molecular Pattern (PAMP) Receptors
in the Airways ..................................................................................................... 4
H) Responses Induced by Activation of PAMP Receptors ..................................... 6
I) Interplay between Epithelial and Immune Cells in the Lung ..............................9
II. Influenza Virus...............................................................................................................10
A) Human Health Impact ...................................................................................... 10
B) Classification .................................................................................................... 10
C) Virology – Influenza Virus Infection and Replication ..................................... 11
D) Mouse Model of Influenza: A/PR/8/34 ............................................................ 12
III. Immune Responses to Influenza Virus ........................................................................ 13
A) Chemotactic Responses during Influenza Virus Infection ............................... 14
B) Responses of Dendritic Cells to Influenza Virus.............................................. 14
C) Responses of Alveolar Macrophages to Influenza Virus ................................. 16
D) Responses of Other Antigen Presenting Cells to Influenza Virus.................... 16
E) Responses of CD4+ T helper cells to Influenza Virus ..................................... 17
F) Responses of CD8+ CTL to Influenza Virus .................................................... 18
G) Responses of NK cells to Influenza Virus........................................................ 19
H) Responses of Granulocytes to Influenza Virus ................................................ 19
I) Responses of B cells to Influenza Virus ............................................................20
J) Antigenic Drift and Shift ...................................................................................20
IV. Immunotoxicology of Cannabinoids ........................................................................... 21
vi

A) Cannabis Use .................................................................................................... 21
B) Cannabinoid Compounds ................................................................................. 22
C) Pharmocokinetics and Metabolism ................................................................... 25
D) Receptors for Cannabinoids ............................................................................. 27
1) Cannabinoid Receptors 1 and 2 ............................................................. 27
2) Alternate Cannabinoid Receptors.......................................................... 28
VII. Intracellular Signaling by Receptors for Cannabinoids .............................................30
A) Signaling by Gαi-protein coupled receptors: Cannabinoid Receptors
1 and 2 .............................................................................................................30
B) Signaling by Alternate Cannabinoid Receptors................................................ 33
VI. Effects of Cannabinoids on Immune Function ............................................................ 34
A) Effects on Antigen Presenting Cells ................................................................. 34
B) Effects on Granulocytes.................................................................................... 35
C) Effects on CD4+ T cells ................................................................................... 35
D) Effects on CD8+ T cells ................................................................................... 37
E) Effects on NK cells ........................................................................................... 37
F) Effects on B cells .............................................................................................. 38
G) Effects on Immune Outcome in Host Resistance Models ................................ 39
H) Phenotype of the Immune System as a Result of Loss or Lack of
CB1 and/or CB2 ...............................................................................................40
VI. Objectives .................................................................................................................... 44
MATERIALS AND METHODS ...................................................................................... 47
I.
II.
III.
IV.
V.
VI.
VII.
VIII.
IX.

Mice. ................................................................................................................ 47
Chemicals and Reagents .................................................................................. 47
T Cell Elicitation for Generation of Functional CTL ...................................... 48
Drug Treatment in vitro ................................................................................... 48
51Cr Release Assay ......................................................................................... 48
IFNγ T Cell Functional Analysis ..................................................................... 49
Proliferation Assay .......................................................................................... 49
T Cell Activation Assays ................................................................................. 49
Procedure for Immunofluorescent Staining of CD4+ and CD8+ Cells for
Flow Cytometry ...............................................................................................50
X.
Influenza instillation. .......................................................................................51
XI.
Necropsy and lung tissue collection. ...............................................................51
XII. BALF analysis and differential cell counts .....................................................51
XIII. Histological and immunohistochemical staining of lung tissue sections ........ 52
XIV. Isolation of alveolar macrophages from BALF ............................................... 53
XV. Flow Cytometry on lung isolated cells. ........................................................... 53
XVI. mRNA extraction and cDNA synthesis. .......................................................... 54
vii

XVII. Mouse immune panel – low density microarray. ............................................ 55
XVIII. Generation and stimulation of bone marrow-derived dendritic cells
(bmDC) in vitro. .............................................................................................. 55
XIX. bmDC co-culture with OT-1 cells. .................................................................. 56
XX. Isolation and activation of naïve CD4+ T cells. .............................................. 56
XXI. Δ9-THC treatment in vivo ............................................................................... 57
XXII. Quantitative Real-time PCR for type I IFN mRNA Statistical analyses ......... 57
XXIII. Statistical analyses ........................................................................................... 58
EXPERIMENTAL RESULTS .......................................................................................... 59
I. Δ9-THC Suppresses CTL Responses Independent of CB1 and CB2 ............................ 59
A) Kinetics of CTL Response ............................................................................... 59
B) Δ9-THC Suppressed CTL Responses during Elicitation but not during
the Effector Response ...................................................................................... 59
C) Concentration-dependent Suppression of CTL Effector Function .................. 62
D) Δ9-THC-Mediated Suppression of CTL Generation....................................... 65
E) Δ9-THC Does Not Affect CD8+ T cell Proliferation Induced by
P815 Co-culture ...............................................................................................71
F) Increase of CD69 Expression on CD8+ Cells with Δ9-THC Treatment .........74
G) Δ9-THC Synergizes with Io to Induce CD69 Expression ...............................81
II. In Vivo Influenza PR8 Immune Response Kinetics in WT and CB1-/-CB2-/- mice ....85
A) Enhanced leukocyte infiltration and injury in airways of CB1-/-CB2-/mice after influenza infection ..........................................................................85
B) Increased mRNA expression levels of pro-inflammatory immune
mediators in lungs from CB1-/-CB2-/- mice ...................................................91
C) T cells from CB1-/-CB2-/- mice undergo accelerated T cell activation..........96
D) Enhanced IL-17 and IFNγ production in NK, and T cells isolated from
lungs of CB1-/-CB2-/- mice in response to influenza ....................................100
E) Fewer Naïve CD4+ T cells in CB1-/-CB2-/- compared to WT mice .............104
F) Increased APC maturation and ability to induce proliferation and cytokine
production in T cells in CB1-/-CB2-/- mice compared to WT mice ..............109
III. In Vivo Influenza Study 3 dpi to Study the Effect of Δ9-THC on Immune
Endpoints of WT and CB1-/-CB2-/- Mice ...................................................................... 115
A) Δ9-THC Reduced Steady-state mRNA Expression of Genes Induced by
Influenza in the lungs of WT but not CB1-/-CB2-/- mice ................................... 115
B) Δ9-THC Suppressed Influenza-Induced Cytokine Production by Effector
Lymphocyte Populations. .................................................................................... 126
C) No Effect of Δ9-THC on Induction of the CD11b+Gr-1+ Population After
Influenza Infection ............................................................................................... 127
D) Reduced Presence of APC with Δ9-THC Treatment After Influenza Infection . 134
viii

E) Greater Maturation in APC from CB1-/-CB2-/- Compared to WT and
CB1-and/or CB2-Dependent Suppression of Maturation by Δ9-THC after
Influenza Infection. .............................................................................................. 134
F) Attenuation of bmDC Maturation by Δ9-THC In Vitro ...................................... 143
G) Impaired Elicitation of T cells after Δ9-THC Treatment of bmDC .................... 147
DISCUSSION .................................................................................................................. 152
I. Suppression of the CTL Response by Δ9-THC ........................................................... 152
A) Reduced CTL Response in CB1-/-CB2-/- Compared to WT mice ................ 152
B) Sensitivity of CTL to Δ9-THC Immune Modulation ..................................... 154
C) Alterations in T cell Activation by Δ9-THC .................................................. 155
D) Summary and Conclusion of the Effect of Δ9-THC on CTL......................... 157
II. Influence of Cannabinoid Receptors 1 and 2 in on the Magnitude of the Immune
Responses to Influenza .................................................................................................... 158
A) Greater Pathology in CB1-/-CB2-/- Compared to WT Mice after Influenza
Infection ......................................................................................................... 159
B) Altered T cell Responses in CB1-/-CB2-/- Compared to WT Mice after
Influenza In Vivo But not after T cell Stimulation In Vitro ........................... 160
C) Dysregulation of APC Responses in CB1-/-CB2-/- Mice Compared to
WT mice ........................................................................................................ 160
D) A Role For CB1 and/or CB2 in Regulating the Magnitude of Immune
Response to Influenza to Reduce Pathology ................................................. 162
III. Role of the Cannabinoid Receptors 1 and 2 in Δ9-THC-mediated Suppression of
Immune Responses to Influenza Virus Challenge ........................................................... 164
A) Contributions of CB1 and CB2 to Cannabinoid Modulation of APC ............ 165
B) Putative Mechanisms by Which Δ9-THC Might Alter APC Function .......... 166
C) Conclusive Remarks on the Role of CB1 and CB2 on APC .......................... 169
IV. Significance, Relevance, and Future Studies ............................................................ 169
REFERENCES ................................................................................................................ 178

ix

LIST OF TABLES
Table 1. Dissociation Constants (Ki) of Select Cannabinoids for CB1 and CB2 ............. 29

x

LIST OF FIGURES
Figure 1. Simplified TLR Signaling ...................................................................................... 8
Figure 2. Structures of Select Cannabinoids ......................................................................... 24
Figure 3. Simplified Gαi GPCR Signaling ............................................................................ 31
Figure 4. CTL Activity Peaks at Day 5 After Elicitation ...................................................... 60
Figure 5. Δ9-THC Suppresses CTL Activity During Elicitation But Not Effector Phase .... 61
Figure 6. Δ9-THC Suppresses CTL Activity in a Concentration-Dependent Manner .......... 63
Figure 7. Lower Viability of CD4+ and CD8+ Cells Following Δ9-THC Treatment .......... 66
Figure 8. Increased IFNγ Production in Live But Not Total Cells After Δ9-THC
treatment ................................................................................................................................ 68
Figure 9. No Effect of Δ9-THC on CTL Proliferation .......................................................... 72
Figure 10. Δ9-THC Increases CD69 Expression on CD8+ cells in a ConcentrationDependent Manner ................................................................................................................ 75
Figure 11. Δ9-THC Decreases CD25 Expression on CD8+ Cells from CB1-/-CB2-/mice in a Concentration-Dependent Manner ......................................................................... 78
Figure 12. Δ9-THC synergizes with Io to upregulate CD69 expression ............................... 82
Figure 13. More Severe Virus-Induced Necrotizing Bronchiolitis and Loss of Clara Cell
Secretory Protein in CB1-/-CB2-/- compared to WT Mice 3 Days Post-infection ............... 88
Figure 14. Greater Neutrophils and Macrophage Recruitment after Influenza Infection
in CB1-/-CB2-/- mice ............................................................................................................90
Figure 15. Relative transcript levels of genes associated with inflammation are increased
in lungs of CB1-/-CB2-/- mice compared to WT mice basally and after influenza
infection at 3 dpi ....................................................................................................................92
Figure 16. Peak H1 mRNA Levels 3 dpi in CB1-/-CB2-/- and WT Mice and Increases
in Steady-state CB1 Transcript Levels 7 dpi ........................................................................94
Figure 17.More rapid T cells activation after Influenza infection in CB1-/-CB2-/compared to WT mice ...........................................................................................................97
xi

Figure 18. Greater cytokine production by leukocytes from CB1-/-CB2-/- mice
compared to WT mice ...........................................................................................................101
Figure 19. Lower number of naïve CD4+ T cells and no difference in T cell stimulation
in vitro in CB1-/-CB2-/-mice compared to WT mice............................................................105
Figure 20. Increased expression of dendritic cell maturation markers in bmDC from
CB1-/-CB2-/- mice ................................................................................................................ 107
Figure 21. bmDC from CB1-/-CB2-/-elicit OT-1 responses without requirement for
maturation stimulus and more potently than WT .................................................................. 110
Figure 22. Alveolar Macrophages isolated from BALF are mature in CB1-/-CB2-/but not WT ............................................................................................................................. 112
Figure 23. Reduction of Gene Expression Associated with Influenza Infection as a
Result of Δ9-THC Treatment ................................................................................................ 116
Figure 24. Δ9-THC does not Alter BALF Cell Counts or Composition After Influenza
Infection ................................................................................................................................. 119
Figure 25. No Change in Percent Immune Effector Cell Composition in Lungs of Mice
Treated with Δ9-THC ............................................................................................................ 120
Figure 26. Decreased IFNγ Production after Δ9-THC Treatment in Lung Isolated
Immune Cells from WT but not CB1-/-CB2-/- mice ............................................................ 122
Figure 27. Decreased IL-17 Production after Δ9-THC Treatment in Lung-Isolated
Immune Cells from WT but not CB1-/-CB2-/- mice ............................................................ 124
Figure 28. No Effect by Δ9-THC Treatment on Presence of Influenza Infection-Induced Gr1+CD11b+ Lung-Isolated Cell Populations .......................................................................... 128
Figure 29. Δ9-THC Treatment does not Alter the Maturation Status of Gr-1+CD11c+
Cells ....................................................................................................................................... 130
Figure 30 Δ9-THC Reduces Recruitment of pDC, cDC, and AM into the Lungs of
A/PR/8/34-Infected Mice Dependent of CB1 and/or CB2 .................................................... 132
Figure 31. Δ9-THC does not Alter the Maturation Status of Lung-Isolated cDC after A/PR/8/34
Infection ................................................................................................................................. 135
Figure 32. Suppression of MHC II Expression in pDC by Δ9-THC after A/PR/8/34
Infection ................................................................................................................................. 137

xii

Figure 33. Δ9-THC does not Alter the Maturation Status of Lung-Isolated AM after
A/PR/8/34 Infection ............................................................................................................... 139
Figure 34. Δ9-THC does not Alter the Maturation Status of Lung-Isolated
Macrophages/Monocytes after A/PR/8/34 Infection ............................................................. 141
Figure 35. Δ9-THC Suppresses TLR-Stimulated bmDC Maturation Independent of
CB1and CB2 .......................................................................................................................... 144
Figure 36. Δ9-THC Impairs Antigen-Specific bmDC-Elicited T cell responses. ................. 148
Figure 37. Δ9-THC Does Not Alter Steady-state Whole Lung mRNA Levels of Type
I IFNs ..................................................................................................................................... 171
Figure 38. DC Cannabinoid Immune Modulation Model ..................................................... 177

xiii

LIST OF ABBREVIATIONS

-ssRNA

Negative single stranded RNA

Δ9-THC

Δ9-Tetrahydrocannabinol

11-OH

11-hydroxy

2-AG

2-arachidonoylglycerol

A/PR/8/34

Influenza virus strain A, Puerto Rico, Isolate 8, 1934

AEA

N-arachidonoylethanolamide or anandamide

AIDS

Acquired immunodeficiency syndrome

AM

Alveolar macrophage

ANOVA

Analysis of variance

APC

Antigen presenting cell

ASC

Apoptosis-associated speck-like protein containing a caspase recruitment
domain

ASK1

Apoptosis signal-regulating kinase 1

AT-II

Alveolar type II cells

BALF

Broncho-alveolar lavage fluid

BALT

Bronchus associated lymphoid tissue

BCS

Bovine calf serum

bmDC

Bone marrow-derived dendritic cell

C

Celsius

CASP-1

Caspase-1

CB1

Cannabinoid receptor 1

xiv

CB1-/-

Mice null for CB1

CB2

Cannabinoid receptor 2

CB2-/-

Mice null for CB2

CB1-/-CB2-/-

Mice null for both CB1 and CB2

CCL20

Chemokine (C-C motif) ligand 20 also known as Macrophage Inflammatory
Protein-3 alpha (MIP-3α)

CCSP

Clara cell secretory protein

CD

Cluster of differentiation

cDC

Conventional dendritic cell

cDNA

Complementary DNA

CFSE

Carboxyfluorescein succinimidyl ester

CO

Corn oil

CO2

Carbon dioxide

COOH

Carboxylic acid

COX-2

Cyclooxygenase-2

CTL

Cytotoxic T lymphocyte

CYP

Cytochrome P450 enzyme

DAG

Diacylglycerol

DC

Dendritic cell

DHBP

Dihydrobenzopyran

dsRNA

Double stranded RNA

FAAH

Fatty acid amide hydrolase

FBS

Fetal bovine Serum

xv

GFP

Green fluorescent protein

GMCSF

Granulocyte macrophage colony-stimulating factor

GMCSFR

Granulocyte macrophage colony-stimulating factor receptor

GPCR

G-protein coupled receptor

GRK

G protein-coupled receptor kinases

H

Hemagglutinin

h

Hours

HAART

Highly active antiretroviral therapy

HSV

Herpes simplex virus

IFN

Interferon

IFNα

Interferon alpha

IFNβ

Interferon beta

IFNγ

Interferon gamma

IL

Interleukin

IL-1

Interleukin 1

IL-6

Interleukin 6

IL-8

Interleukin 8

IL-12p40

Interleukin 12 beta subunit 40kD

IL-17

Interleukin 17

Ig

Immunoglobulin

IκB

Inhibitor of kappa B

Io

Ionomycin

IP3

Inositol trisphosphate

xvi

IRAK

Interleukin-1 receptor-associated kinase

IRF3

Interferon regulatory factor 3

IRF7

Interferon regulatory factor 7

JAK-STAT

Janus tyrosine kinase-signal transducer and activator of transcription

JNK3

C-Jun N-terminal kinase 3

Kv channel

Voltage-gated potassium channel

LD50

Lethal dose at which half of the tested animals die

LPI

Lysophosphatidylinositol

LPS

Lipopolysaccharide

M1

Matrix protein (influenza)

M2

Ion channel protein (influenza)

Mac/Mθ

Macrophage/monocyte

MAP kinase

Mitogen-activated protein kinase

MDA5

Melanoma differentiation associated gene-5

MDSC

Myeloid derived suppressor cells

MGL

Monoglyceride lipase

MHC

Major histocompatibility complex

MIP

Macrophage inflammatory protein

MKK4

Mitogen-activated protein kinase kinase 4

MyD88

Myeloid differentiation primary response gene (88)

N

Neuraminidase

NA

Naïve

NAD

Normal antibody diluent

xvii

NALT

Nose associated lymphoid tissue

NFκB

Nuclear factor kappa B

NFAT

Nuclear factor of activated T cells

NK cell

Natural killer cell

NLR

NOD-like receptors

NLRP3

NLR pyrin domain containing 3

NOD

Nucleotide-binding oligomerization domain-containing (NOD)

NP

nucleoprotein

OT-1

TCR receptor transgenic mice for specific for OVA257-264 (SIINFEKL)

PA

Polymerase subunit A (influenza)

PAMP

Pathogen-associated molecular pattern

PB1

Polymerase subunit B1 (influenza)

PB2

Polymerase subunit B2 (influenza)

pDC

Plasmacytoid dendritic cell

pfu

Plaque-forming units

PI

Phosphatidylinositol

PKR

Double-stranded RNA-dependent protein kinase

PLC

Phospholipase C

PMA

Phorbol myristate acetate

PPARγ

Peroxisome proliferator-activated receptor gamma

PR8

A/PR/8/34 (short form)

RANTES

Regulated on activation, normal T cell expressed and secreted

RIG-I

Retinoic acid-induced gene-I

xviii

ROS

Reactive oxygen species

RXRα

Retinoid X receptor alpha

SAL

saline

SAS

Statistical Analysis System

Sp

Surfactant protein

SPLC

Splenocytes

ssRNA

Single stranded RNA

TAK1

Transforming growth factor β–activated kinase 1

TAB

TAK1 binding protein

TBS

Tris buffered saline

TGFβ

Transforming growth factor beta

Th cell

T helper cell

TLR

Toll-like receptor

TNFα

Tumor necrosis factor alpha

TRAF

TNF receptor associated factor

TRIF

Toll/IL-1 receptor domain-containing adaptor inducing IFN-beta

TRPC1

Transient receptor potential channel 1

UC

Ulcerative colitis

VH

vehicle

vRNA

Viral RNA

WT

Wild type

xix

INTRODUCTION

I. Respiratory Tract and Lung Associated Immune Defenses against Pathogens
A) Respiratory Tract Anatomy and Function
The respiratory tract’s main function in healthy individuals is to facilitate gas exchange
of oxygen acquired from the atmosphere and carbon dioxide produced inside the body at the
alveolus. The main purpose of oxygen inside the cell is to serve as an electron acceptor in the
generation of energy in the form of adenosine triphosphate from carbon-containing energy
sources inside the mitochondria. The respiratory tract starts at the nose, continues through the
trachea, bronchi, lung lobes, bronchioles, and alveoli [1]. Furthermore, due to its large surface
area and contact with the external environment it is a site for antigen exposure and mucosal
immune reactions [2-4].

B) Differences in the Respiratory Tract between Humans and Rodents
There are differences in anatomy, respiration, and cellular composition of the lung in
rodents such as the mouse compared to human. Anatomically, aside from obvious size
differences, mice have four right (apical, cardiac, azygous, and diaphramatic) lung lobes and one
left lung lobe, while humans have three right (superior, middle, and inferior) and two (superior
and inferior) left lung lobes. Unlike humans, rodents do not have respiratory bronchioles, their
respiratory tract is linear, their noses differ in anatomy and cell distribution, and their airways
have a proportionally greater diameter [5-8]. Functionally, rodents are obligate nose breathers,
while humans are only obligate nose breathers as infants and later develop into naso-oral
breathers [8-10]. Furthermore, the lungs of mice contain more Clara cells than humans [11-13].

1

Aside from these few differences, the mouse is a suitable model to study immune responses
including those in the lung, due to the similarities to immune responses elicited to influenza for
example, as discussed below.

C) Lung Associated Lymphoid Organs
The lung consists of a large mucosal surface and due to inhalation presents a site of
chronic non-pathogenic and occasional pathogenic antigen exposure [3, 14]. While the nose,
mouth, and trachea have a natural flora of bacteria, the airways and alveoli are sterile in healthy
individuals [15-17]. The lung harbors resident immune cells, including alveolar macrophages
(AM) and dendritic cells (DC). In addition, the lung is highly perfused resulting in frequent
traffic of immune cells through lung tissue [14]. Lymphoid structures associated with the
respiratory tract include the cervical lymph nodes, hilar lymph nodes and mediastinal lymph
nodes, bronchus associated lymphoid tissue (BALT), and nose associated lymphoid tissue
(NALT). However, the cervical lymph nodes and NALT seem to be less important in the
generation of some immune responses such as those to influenza [18]. Other cells, including
epithelial cells, perform immune functions, too, as discussed in more detail below. BALT and
NALT consists of immune cell aggregates in the mucosa, including T cells, B cells, DC,
macrophages, and high endothelial venules, which allow exit of immune cells from the blood and
entry into the lymphatics. In addition, the formation of BALT can be induced by infection,
especially during chronic infections [14, 19-22]. The size of the BALT varies dramatically
among mouse strains and a particularly small BALT has been observed in the C57Bl/6 strain
compared to other strains [23].

2

D) Mechanical Defenses
The first lines of defense in the respiratory tract are the nose, which acts to filter inhaled
air and the larynx, which acts as a sphincter during expectoration and cough, reducing backflow
of inhaled substances [24]. The cough reflex acts as a mechanical defense to expel
mucosubstances potentially containing pathogens [25]. Despite these first defenses, inhaled
particles reach the respiratory epithelium. Due to constant exposure to environmental antigens,
the respiratory epithelium has several protective mechanisms against infection and various
means to regulate immune responses in the lung [26]. The physical barrier of the respiratory
epithelium presents another line of defense to insult. Also, the mucociliary apparatus constantly
moves inhaled particles that have deposited on the mucous layer of the respiratory tract via
sedimentation, impaction, Brownian diffusion, interception, and electrostatic precipitation out of
the airways [27, 28]. Mucus contains several antimicrobial factors including surfactant proteins
(Sp), defensins, cathelicidins, lactoferrin and lysozyme as well as additional antimicrobials
secreted from immune cells, such as immunoglobulin A (IgA) as discussed later [24, 29].

E) Respiratory Epithelial Cells
There are several types of cells with different functions in the respiratory epithelium in
humans and mice including: type I (AT-I) and type II alveolar cells (AT-II), ciliated epithelial
cells, mucous (goblet) cells, and Clara cells. While AT-I and AT-II cells are found in the alveoli,
the respiratory epithelium in the bronchi and bronchioles is lined with ciliated cells and goblet
cells. Clara cells are found in the bronchioles. The glylcoproteins that make up the mucus are
produced by goblet cells and the produced mucus is transported by the mucociliary apparatus
away from the airways towards the proximal end of the airways by the unidirectional beating of

3

cilia from ciliated cells. Clara cells are the main secretory cells in the airways and also possess
proliferative capacity [11, 30]. Furthermore they are involved in metabolism of xenobiotics in
the lung [31]. Type I epithelial cells are involved in the gas exchange and are terminally
differentiated. Most of the lung is lined by the type I cells due to their large size and their main
function is to oxygenate the blood, but are fewer in number than AT-II cells [32]. In addition,
they are able to transfer ions in and out of the respiratory lumen and thus contribute to lung
liquid homeostasis [33]. AT-II cells have secretory functions, metabolic capacity, and also are
able to differentiate into type I cells to allow for regeneration of the airway epithelium.
Pulmonary surfactant is among the secretions of AT-II cells and reduces surface tension of the
alveoli and thereby increases pulmonary compliance [31, 34, 35].

F) Immune Functions of Type II and Clara Cells
The release of surfactant protein A (SpA) and SpD is mediated by AT-II and Clara cells.
Both Sps have the ability to neutralize pathogens and facilitate phagocytosis and to reduce
cytokine production by inflammatory cells, thereby protecting lung integrity [36]. In addition, βdefensins and cathelicidins produced by the epithelial cells act as broad spectrum antimicrobials
which are thought to occur through electrostatic interactions with membranes of pathogenic
bacteria and subsequently inducing permeabilization. In part the release of these soluble
mediators is regulated by NaCl concentrations (high [NaCl] is inhibitive) in the lumen, but also
induced in response to presence of cytokines [37-39]. Lactoferrin is a protein involved in iron
binding, thereby limiting iron to bacteria, but it also acts directly by binding to the membranes of
microbes thereby preventing microbe colonization [40, 41]. Lysozyme is directly involved in the

4

lysis of pathogens, specifically demonstrated for bacteria [42, 43]. In addition the epithelial cells
contribute to cytokine production which is discussed in the next paragraphs.

G) Expression of Pathogen Associated Molecular Pattern (PAMP) Receptors in the
Airways
The ability of the lung epithelial cells and resident innate immune cells to secrete
cytokines in response to presence of pathogens is essential to forming immune responses in the
lung. Several PAMP receptors exist and are expressed in the respiratory epithelium or associated
cells including AM, DC, lung fibroblasts, airway epithelial cells, lung endothelial cells and lung
smooth muscle cells: toll-like receptors (TLR), nucleotide-binding oligomerization domaincontaining protein (NOD)-like receptors (NLR), retinoic acid inuduced gene I (RIG-I) like
receptors and other cytosolic DNA sensors [44]. Ten TLRs (TRL 1-10) have been identified in
the human genome, they respond to PAMPs, and natural or synthetic ligands for these receptors
have been identified [45, 46]. Type I epithelial cells in the alveoli express TLRs 1-6 and 9 [4749]. AM express TLR 1, 2, 4, and 6-8, plasmacytoid DC (pDC) in the lung express TLR 7 and 8
and conventional DC (cDC) TLRs 1-4. In addition, lung fibroblasts, endothelial cells, and
smooth muscle cells have been reported to express TLRs 2-4, 8 and 9 [44]. In addition to
recognition of foreign material, TLRs are able to recognize intracellular material, usually found
in the extracellular fluid after release from necrotic cell death. The intracellular material
includes, but is not limited to, heat-shock proteins (bind to TLR 2 and 4) and mRNA (binds to
TLR 3) [50-53]. Cytosolic DNA sensors include NLR, RIG-I, melanoma differentiation
associated gene-5 (MDA5), as well as double-stranded RNA-dependent protein kinase (PKR)
and others [54-56]. Among the NLRs, the most studied and best understood examples is NRL

5

pyrin domain containing 3 (NLRP3). Expression of NLRP3 was found in neutrophils and
macrophages and NLRP3 was highly expressed in monocytes and cDC but not expressed in
airway epithelial cells and only low levels in lymphocytes, eosinophils, and pDC [57, 58]. RIG-I
expression has been found in airway epithelial cells, cDC and fibroblasts but RIG-I is absent in
pDC [59-61]. MDA5 is involved in the cytosolic DNA sensing in macrophages and cDC but its
expression can be induced in other cell types by type I interferons (IFNs) [62-64]. PKR is
expressed in airway epithelial cells, immune cells and many other tissues and in addition to
sensing double stranded RNA (dsRNA) it incorporates other cellular stress signals [65, 66].
Collectively, cells of the respiratory epithelium and immune cells are contributing to the sensing
of pathogens by molecular patterns in the lungs.

H) Responses Induced by Activation of PAMP Receptors
After recognition of PAMPs, innate immune responses are initiated. For example, the
influenza virus pathogen directly activates TLR 7, NLRs, RIG-I, and MDA5, but other receptors
might be indirectly involved as a result of cell damage or as a result of viral replication [67-71].
While TLR 1, 2, 4-6, and 8-10 are expressed on the cellular surface, TLR 3, 7, and 9 are found in
the endosome [45, 46, 72]. TLR signaling shares conserved pathways within the receptor family.
Specifically, the adapter protein myeloid differentiation primary response gene (88) (MyD88)
activates interleukin-1 receptor-associated kinases (IRAK) 4, the subsequently activated kinase
IRAK1 then activates TNF receptor associated factor 6 (TRAF6), which results in the activation
of two transcription factors: nuclear factor kappa B (NFκB) by phosphorylation and thus
degradation of inhibitor of nuclear factor kappa-B (IκB) by IκB kinase after interaction with the
Transforming growth factor β (TGF-β)–activated kinase 1 (TAK1) / TAK1 binding protein

6

(TAB) complex. Also, the transcription factor activator protein 1 (AP-1) is activated as a result
of activation of mitogen-actived protein kinases (MAPK). These two transcription factors drive
the expression of a host of genes, including pro-inflammatory genes including interleukin (IL) 1,
IL-6, IL-12 p40, and tumor necrosis factor alpha (TNFα) [45, 46, 73, 74]. In addition, activation
of TRL3 and TLR4 results in MyD88-independent increase of the transcriptional activity of
interferon regulatory factor (IRF) 3 and IRF7, which are required primarily for the production of
type I IFNs IFN-β and IFN-α, respectively. Instead of MyD88, Toll/IL-1 receptor domaincontaining adaptor inducing IFN-beta (TRIF) serves as the adapter protein. TRIF either signals
through TRAF6, or TRAF3 [75, 76]. Activation through TRL7 and TLR9 triggers IRF7 activity
entirely dependent on MyD88 through IRAK4/IRAK1, leading to the production of IFNα [73].
Activation of the NLRs is best understood in NLRP3 and results in inflammasome formation,
containing the adapter protein apoptosis-associated speck-like protein containing a caspase
recruitment domain (ASC) together with caspase-1 (CASP-1), thereby activating IL-1 and IL-18
by proteolytic cleavage [77]. Complete NLRP3 activation requires TLR signaling as well as cell
damage signals which increase the transcriptional activation of inflammatory mediators into their
pro-form. In the case of IL-1, for example, the transcription of pro-IL-1 is induced by TLR
agonists, but the conversion and bioactivation of pro-IL-1 into IL-1 is performed by caspase 1
[78, 79]. The signals that directly activate NLRs include, but are not limited to, bacterial flagella,
anthrax lethal toxin, uric acid, asbestos, silica and saturated fatty acids [80-83]. In addition, the
NLRP3 inflammasome has been implicated in the recognition of influenza ssRNA and thus is
part of the host response to influenza and also has been found to play a role in the tissue healing
response [58, 84]. Also, as with the TLRs, there are endogenous ligands for the inflammasome
pathway, such as misfolded proteins [85, 86]. RIG-I, MDA5, and PKR activate IRF3 and thus

7

Figure 1. Simplified TLR Signaling.
For interpretation of the references to color in this and all other figures, the reader is
referred to the electronic version of this dissertation.

8

contribute to IFNβ production. Furthermore, PKR activates the NFκB pathway, upregulating
pro-inflammatory genes in response to dsRNA [87]. Overall, the activation of PAMP receptors
leads to the increase in production of pro-inflammatory cytokines and type I IFNs, which are
essential in the immune response to pathogenic infection of the respiratory tract including
influenza virus. The signaling described above is summarized in Figure 1.

I) Interplay between Epithelial and Immune Cells in the Lung
Interactions between the respiratory epithelium and resident, as well as recruited immune
cells in the lung are an important part of immune defense [88]. In addition to the cells of the
respiratory epithelium, resident immune cells in the lung include AM and DC. Intraepithelial DC
were found to be CD11b+CD11c+ and thus are phenotypically closer to cDC than pDC [14, 8991]. As a result, the main function of intraepithelial DC is antigen presentation rather than
proinflammatory cytokine release. Under homeostatic conditions, AM are involved in impairing
adaptive immunity by the release of nitric oxide, prostaglandins, IL-10 and transforming growth
factor-β (TGFβ) [92]. In the event of PAMP activation, proinflammatory immune responses are
initiated by epithelial cells, AM and DC in the lung and dictate the anteceding immune response.
As a result, soluble mediators comprised of cytokines, chemokines, lipids and reactive oxygen
species (ROS) cause an influx of immune cells to the airways first via the circulation and later
via the afferent lymphatics [26]. For example, migration of cDC to the airways depends on
airway epithelial cell produced Chemokine (C-C motif) ligand 20 (CCL20), which is enhanced
during influenza infection [93, 94]. In addition to attracting immune cell populations to the lung,
the respiratory epithelial cells and airway associated immune cells contribute to the licensing of
the immune cells attracted to the lung by the release of cytokines [95]. The plasticity of DC and

9

their ability to elicit adaptive immune responses according to the environment encountered in
tissues is well documented and is discussed in a separate section below [96]. Taken together,
epithelial and resident immune cells in the lung control immune homeostasis and initiate
proinflammatory responses in the lung to recruit immune cells involved in defense against
pathogens.

II. Influenza Virus

A) Human Health Impact
Influenza virus is a respiratory virus and infects ciliated (avian origin) and non-ciliated
(human origin) cells of the respiratory tract. Influenza causes on average about 40,000 deaths due
to infection and immunopathology or infection with a secondary pathogen and is believed to be
responsible for 200,000 hospitalizations per year in the US [97, 98]. The estimated total US
economic cost of an annual influenza virus season is close to $90 billion, which includes cost of
hospitalization, reduced productivity due to absence from the workplace and other costs
including those associated with deaths [99].

B) Classification
Influenza A, B and C viruses belong to the orthomyxoviridae family of RNA viruses.
The genome is composed of negative single stranded RNA ((-)ssRNA). Among the subtypes of
influenza viruses, the most common is influenza A virus, which is of greatest concern due to its
virulence and its potential to give rise to pandemics or epidemics. For the sake of brevity only

10

influenza A virus will be discussed. Based on antibody responses generated to the surface
proteins hemagglutinin (H) and neuraminidase (N), the influenza A virus has been subdivided
into several serotypes, for example H1N1 (human origin), the cause of the Spanish flu in 1918
and H5N1 (avian origin), responsible for the recent bird flu pandemic [100-102]. To identify
isolates, the following nomenclature is used: type of influenza (A, e.g.)/origin/isolate
number/year isolated. For example a common strain of virus used in laboratories and in the
experiments comprising this dissertation is A/PR/8/34, where PR stands for “Puerto Rico”, the
origin of this isolate.

C) Virology – Influenza Virus Infection and Replication
The genome of influenza A viruses consists of eight segmented RNA strands that encode
11 proteins: envelope glycoproteins H and N, matrix protein (M1), the ion channel protein (M2),
nucleoprotein (NP), three polymerase subunits (PB1, PB2, PA), and non-structural proteins NS1
and NS2 [103, 104]. Due to the virus’ small genome it lacks the machinery necessary for
autonomous replication and its propagation relies on a plethora of host factors [105]. To attach to
host cells the influenza virus recognizes the monosaccharide N-acetylneuraminic (sialic) acid on
the host cell surface via its hemagglutinin surface protein [106]. Entry of the influenza virus into
the host cell is usually mediated by clathrin-coated pits, but clathrin-independent pathways also
exist [107]. After endocytosis, viral uncoating occurs in the endosomal compartment due to a
reduction in pH that allows for a conformational change in hemagglutinin and the fusion of the
viral envelope with the endosomal compartment, thereby releasing the contents of the viral
capsid into the cytoplasm [108, 109]. After release from the capsid, the ribonucleoprotein
complex containing the viral genome is shuttled into the nucleus via nuclear localization signals

11

[110]. Inside the host cell nucleus, negative strand viral RNA (vRNA) is translated into positive
strand vRNA by the viral polymerases from the capsid for viral protein synthesis and
complimentary RNA, the intermediate of more negative sense genomic RNA. After nuclear
export, mRNA is translated into protein via membrane bound ribosomes in the endoplasmic
reticulum and modified in the Golgi apparatus. These post-translational modifications in the
Golgi direct assembly of viral protein into a virion at the cell membrane, where incorporation of
RNA segments into the virions occurs [101, 104]. After assembly at the plasma membrane, the
virion remains attached to the sialic acid residues on the membrane (as described before) and the
virions release depends on the sialidase activity of neuraminidase [111-113]. Due to the low
fidelity of the viral polymerases frequent mutations occur in the viral genome [114], which is the
underlying mechanism for antigenic drift (discussed later). In addition, in the case of dual
infection by two non-identical influenza viruses in the same host, the genomes of the viruses may
undergo a shuffling event, creating virus progeny with drastically altered RNA genome and
amino acid sequences compared to the parent virions. This is called antigenic shift (also
discussed in more detail later) [104, 115].

D) Mouse Model of Influenza: A/PR/8/34
Unlike ferrets, for example, mus musculus (laboratory mouse) cannot be infected with
human isolates of influenza virus. Furthermore, m. musculus is not a natural host species for
influenza virus. However, repeated passage of a human influenza virus isolates increases
virulence in mice, thereby adapting the virus to its new host. Compared to other species, the
mouse provides a relatively inexpensive and well characterized model of influenza infection and
the ensuing immune response. After intranasal instillation with influenza virus, mice develop

12

lesions of the upper and lower respiratory tract with similar histopathology to humans [116, 117].
Over many years, influenza mouse models have greatly contributed to the understanding of the
immune response to influenza [116]. One of the most commonly used experimental models to
study influenza infection employs the mouse-adapted A/PR/8/34 strain and murine host
challenge has been previously established and characterized in this laboratory [118-120].

III. Immune Responses to Influenza Virus
Influenza virus infection activates both innate and adaptive immune responses. The immune
response to influenza optimally remains within a response magnitude window, limiting excessive
immune responses, while at the same time inducing a sufficiently robust immune response to
clear the virus. The antigen-specific adaptive immune responses provide the host with protective
immunity against further infections from the same pathogen. By cell depletion studies it became
evident that several effector lymphocytes of the adaptive immune system are important in the
clearance of an influenza virus infection. Depletion of B cells, CD4+ T helper (Th) cells or
CD8+ cytotoxic T lymphocytes (CTL) delayed viral clearance and depletion of more than one
population resulted in almost complete virus-induced mortality with otherwise non-lethal
challenge [121]. The immune response to influenza remains localized to the site of infection
including the airways and lung associated lymph nodes, but is only found after viral clearance in
the spleen [122]. The innate immune responses were discussed in detail above and include
PAMP recognition by epithelial and resident immune cells in the lung. The recognition by
PAMPs induces innate immune responses and results in the release of chemokines and
proinflammatory cytokines and lung. Further discussion on innate responses will focus on the

13

contribution to inflammation, pathology, and the adaptive immune response. The responses of
immune cells important in the immune defense against influenza are discussed in more detail
below.

A) Chemotactic Responses during Influenza Virus Infection
Influenza virus induces the release of several chemokines at the site of infection: macrophage
inflammatory proteins (MIP) 1α, 1β, 2, and 3α, monocyte chemotactic protein 1, interferoninducible protein and Regulated on activation, normal T cell expressed and secreted (RANTES).
Cytokines and chemokines induced by IFN-γ and IL-8 appear to be important for the recruitment
of antigen presenting cells, neutrophils, T cells and NK cells to the lung after infection [93, 94,
123]. These chemokines are produced by the respiratory epithelium, monocytes/macrophages, T
cells, DC and neutrophils [124]. Thus recruited immune cells contribute to the amplification of
chemotactic signals generated initially by epithelial and resident immune cells.

B) Responses of DC to Influenza Virus
APC are involved in relaying stress information from TLR signaling or inflammasome
activation and elicit appropriate lymphocyte responses depending on the stress signal
encountered. Unlike lymphocytes their response to an immune stimulus does not depend on
clonal expansion. DC are the most effective APC and are thus most effective at eliciting
lymphocyte immune responses. As a result, they link the innate to the adaptive immune system
by relaying information from innate stimuli and elicting effector cell responses accordingly [96].
DC are found in the airways and even in the alveoli in the absence of an immune stimulus. After
an immune stimulus, such as influenza infection, DC migrate from the site of infection to the

14

draining lymph node and interact with and elicit effector immune cell populations [14].
Recruited DC populations display a mature phenotype based on the surface expression of T cell
co-stimulating clusters of differentiation (CD) 80, CD86, and major histocompatibility complex
II (MHC II), which is involved in antigen presentation to CD4+ T cells and constituitively
express MHC I as all other nucleated cells for antigen presentation to CTL [125]. After infection,
among APC, DC harbor the greatest amount of influenza virus as demonstrated by cellular
analysis of the lung immune cell populations after infection with a green fluorescent protein
(GFP) expressing virus [126]. Although these data do not differentiate between viral capture and
active viral infection of DC, DC are recognized for their superior ability to acquire and process
antigen [96]. Similar to effector lymphocyte populations, successful clearance of influenza virus
depends on several DC populations, however the distinct roles of these DC subpopulations are
poorly defined [127, 128]. There are two main types of DC, pDC and cDC. These cells are
identified by their surface expression of CD11c, CD11b, and Gr-1. cDC are CD11c+, CD11b+,
and Gr-1-, while pDC are CD11c+, CD11b-, Gr-1+ [127, 129-133]. Although cDC and pDC
arise from distinct immediate committed precursors, further upstream a common precursor is
shared [134, 135]. While the main function of cDC seems to be antigen capture, antigen
presentation, and the elicitation of effector immune cells, pDC produce great amounts of type I
IFNs and retain T cell elicitation functions, although are less apt at T cell elicitation than cDC
[136, 137]. Both cDC and pDC express the maturation markers CD80, CD86, and MHC II and
secrete an overlapping profile of cytokines, however the amount released varies depending on
the stimulus and can differ between cDC and pDC [128-130, 138, 139]. Due to their abilities to
present antigen and secrete cytokines, DC polarize effector lymphocyte responses, especially
CD4+ T helper (Th) cells. There are at least four different ways to polarize Th cells which is

15

dependent on the presence of certain types of cytokines: Th1 (induced by IL-12, primarily
secretes IFNγ), Th2 (induced by IL-4, primarily secretes IL-4, IL-5, IL-13), Th17 (induced by
TGF-β and IL-6, primarily secretes IL-17), Treg (induced by TGF-β and IL-10 or IL-2, primarily
secretes IL-10 and suppresses immune function) [140, 141]. Although mainly thought of as
producers of type I IFNs, pDC can also contribute to Th polarization [142]. Overall, DC are
central mediators of immune function and important for proper immune responses against
viruses and maintaining immune homeostasis in the absence of a pathogen.

C) Responses of AM to Influenza Virus
AM are resident immune cells in the lung and are identified as CD11c+CD11b-Gr-1- [133].
Alveolar macrophages are important as a first line of defense against viral infection and produce
type I IFNs in response to viral recognition [143]. The mechanism by which type I IFNs are
induced involved intracellular (PKR) and extracellular receptors (TLR) as discussed above.
Although AM might not have profound effects on the adaptive immunity, they can capture
antigen and are among the first cells to enter the draining lymph nodes after pathogen infection
of the lung [144, 145]. Furthermore, AM seem to play a role in maintaining tolerance in the lung
thereby suppressing inflammation in the absence of a pathogenic stimulus [146].

D) Responses of Other APC to Influenza Virus
In addition to DC and AM, the presence of pathogens in the lung recruits other immune cell
populations. These include be blood-derived macrophages and monocytes (CD11bloCD11clo)
and myeloid-derived suppressor cells (MDSC) (CD11b+Gr-1+) [133]. Blood-derived
macrophages are among other cells associated with inducing inflammatory conditions by release

16

of pro-inflammatory mediators including ROS and cytokines that can contribute to a robust
adaptive immune response, yet excessive function might result in pathology [147]. MDSC on the
other hand are known for their ability to suppress immune function. After influenza infection,
MDSC are thought to be induced to ameliorate the inflammation associated with the infection
and require cell-cell contact for their suppressive activity, which might be mediated by ROS
[148-150]. Furthermore, blood derived macrophages are involved in the response to influenza
virus. There are the classically activated macrophages (M1) and the alternatively activated
macrophages (M2). While M1 play a role in inflammation and mainly produce proinflammatory
cytokines as a result of PAMP activation in Th1 environment such as influenza infection, M2 are
often found in Th2 environments, such as helminth infections and some forms of asthma [151].
However, unlike DC, macrophages do not produce IL-12 and IL-23 in response to influenza
infection, which are the canonical cytokines driving the Th1 and Th17 lineages, respectively
[152].

E) Responses of CD4+ T helper cells to Influenza Virus
Naïve CD4+ T cells exhibit high plasticity to environmental cues, which allows them to
accept a phenotype according to the instructions received from APC, most importantly DC.
Unlike APC, T cells need to clonally expand and the milieu of cytokines encountered drives the
T cell clones towards a certain Th phenotype. T helper cells are elicited by DC via the interaction
with peptide antigens of usually 12-15 amino acids presented in the context of MHC II and the T
cell receptor (TCR) (signal 1) and by receiving a second signal from co-stimulatory molecules
such as CD80 or CD86 on DC via the T cell co-stimulatory receptor CD28 (signal 2). Th
contribute to the immune response to influenza by production of cytokines necessary for optimal

17

NK, CTL, and B cell responses but can also induce maturation in APC. In addition, they are able
to mediate direct lysis of virus infected target cells by Fas-Fas ligand interaction [121, 153].
While on the one hand, the cytokines produced by Th cells are necessary for appropriate
response of other immune cells as mentioned above, their non-discriminating action can
contribute to immunopathology [154]. During an influenza infection the main Th types found are
Th1 (producing mainly IFNγ) and Th17 (producing mainly IL-17) [155]. While IL-17 promotes
pro-inflammatory responses [156-158], IFNγ exerts potent antiviral activity [159, 160], with both
cytokines contributing to the immunopathology associated with influenza infection [161-163].
However, similar to APC, Th cells can contribute to a mechanism by which excessive
inflammation is reduced, such that in acute infection IL-10 is produced by Th cells, a cytokine
with potent immunosuppressive properties as well as important in the differentiation of B cells
[164].

F) Responses of CD8+ CTL to Influenza Virus
CTL are elicited by DC in an antigen-specific manner by interaction with 8-10mer peptides
presented in the context of MHC I, and like CD4+ T cells also depend on a co-stimulatory signal
from DC received through CD28 on the T cell’s surface. Differentiated CTL directly lyse target
cells presenting cognate peptides on MHC I (present on all nucleated cells) by release of a
cytolytic granules containing perforin and granzyme [154, 165]. Most of the CTL response is
directed towards a select few peptides of the proteasome-processed influenza proteome, which
are strong ligands of the MHC I binding pocket [166]. In addition to their lytic activity, CTL can
produce IFNγ and IL-17, the latter of which occurs at the expense of lytic activity [167]. As

18

observed with Th cells, CD8+ T cells also produce IL-10 to dampen excessive inflammation
during acute viral infections [164].

G) Responses of NK cells to Influenza Virus
NK cells are immediate responders in the immune response to influenza. They do not require
previous activation and elicitation in order to lyse target cells by binding of polymorphic and
non-polymorphic MHC I-like proteins via release of cytotoxic granules containing perforin and
granzyme. Furthermore, NK cells produce cytokines including IFNγ and IL-17. NK cells can
also be directly activated by TLRs or by interaction with DC [168-171]. In addition, NK cells
contribute to responses of other immune cells, such as CTL, by creating a cytokine milieu that
favors generation of these cells [172]. Also, NK cells have receptors for antibodies that allow
NK cells to bind to B cell-produced Ig in order to mediate their cell lysis [173].

H) Responses of Granulocytes to Influenza Virus
Among the granulocyte population, neutrophils have been predominantly associated with
playing a role in influenza infection. Early presence of neutrophils is required for optimal viral
clearance; however, excessive presence and function of neutrophils has been associated with
immunopathology [174, 175]. In addition to activation in response to the presence of cytokines,
TLR activation of neutrophils occurs during influenza infection [176]. Their functions include
the release of oxidants, proteases, and antimicrobial peptides, which contribute to host defense,
however may also induce adverse tissue damage in the host [177]. In addition, neutrophils are
involved in the removal of necrotic cells and produce chemokines that attract other immune cells
[178].

19

I) Responses of B cells to Influenza Virus
B cells ultimately provide long lasting protection against influenza virus by production of
antibodies that recognize surface antigens H and/or N [179-181]. B cells rely on stimulation by
CD4+ T cells or DC for their optimal function, thus the B cell response is delayed compared to
other leukocytes [182]. Initially B cells produce IgM, and after affinity maturation produce
IgG2a, IgG2b, and IgA to neutralize influenza virions [183, 184]. TLR receptors on B cells and
the products of TLR receptor ligation from other immune cells are important for achieving the
appropriate isotype switching of B cells in the immune response to influenza [185].

J) Antigenic Drift and Shift
Although prior influenza virus infections provide the host with protective immunity in the
event of an anteceding infection, this is only true if the repertoire of viral antigens does not differ
significantly between infections. Of the estimated 40,000 casualties due to influenza virus, there
are more deaths associated with virus infections after drastic changes in viral surface antigens.
These can result from minor antigenic differences (antigenic drift) as a result of error prone
replication or gene reassortment from concurrent infections of two non-identical viruses
(antigenic shift) in a carrier host and are both based on surface H and N [104]. The host immune
system is unable to recognize modified or completely new viral antigen due to lack of immune
memory, such that primary immune responses are generated. This is one of several ways that
viruses evade the immune system and creates the need for annual influenza vaccinations which
primes the host’s immune system with the projected viral antigens of the season allowing for an
accelerated and robust immune response [115, 186].

20

IV. Immunotoxicology of Cannabinoids
Immunotoxicology is defined as the study of xenobiotics and organisms that adversely
affect the function of the immune system. These toxic responses include suppression or
enhancement of immune function, loss of tolerance resulting in autoimmunity, and the initiation
of immune responses to neoantigens [187].

A) Cannabis Use
Cannabis or marijuana is the most commonly abused illicit drug with about 6% of the US
population above the age of 12 and more than 20% in the 18-20 year old group reporting
monthly use [188, 189]. Although human exposure to cannabinoids is primarily through smoking
or ingestion of parts of the marijuana plant, consumption of synthetically produced cannabinoid
compounds exist. Despite federal regulations, marijuana as well as other cannabinoids are used
for recreational and medical purposes [190]. Also, there are several states in the US, including
Michigan in 2008, that have decriminalized marijuana use for medical purposes [191]. Medical
uses include, but are not limited to, ameliorating pain associated with cancer, as an antiemetic in
chemotherapy, and as treatment for glaucoma, epilepsy and the autoimmune disorder multiple
sclerosis. Also marijuana use is common among acquired immunodeficiency syndrome (AIDS)
patients and was previously used as an appetite stimulant in patients with AIDS-associated
wasting syndrome before the emergence of highly active antiretroviral therapy (HAART) [192].
Due to the high prevalence of cannabis use in the US population and the potential of cannabis

21

use to contribute to adverse health outcomes, there is a public health interest in studying the
effects of cannabinoid compounds on various organ systems including the immune system.

B) Cannabinoid Compounds
The first cannabinoid compound identified for its psychoactive activity was Δ9tetrahydrocannabinol (Δ9-THC) in 1964 from Cannabis sativa or marijuana and is also the
plant’s main psychoactive component [193]. The family of plant-derived cannabinoids consists
of more than 60 structurally-related tricyclic phytochemicals. The production of plant-derived
terpenophenolic compounds occurs in the glandular trichomes predominantly of the female
marijuana plant [194]. These phytocannabinoids exibit varying binding affinity at their identified
target receptors, cannabinoid receptor 1 (CB1) and 2 (CB2). For example, while Δ9-THC is a
non-selective agonist of CB1 and CB2, cannabidiol (CBD), a phytocannabinoid with similar
structure to Δ9-THC, lacks binding activity at CB1 and CB2 [195]. Important for the binding
activity of phytocannabinoids is the dihydrobenzopyran (DHBP) structure as well as a hydroxyl
at the C-1 aromatic position and an alkyl group at the C-3 aromatic position. Furthermore, only
plant compounds in cis configuration act as ligands of CB1 and/or CB2 [196]. Besides CB1 and
CB2, other receptors have been proposed as targets for cannabinoids and receptors for
cannabinoids will be discussed in a separate section.
Endogenously-produced cannabinoids, also termed endocannabinoids, have been
identified and are synthesized within the cell from membrane lipids. Endocannabinoids were
identified based on their ability to bind to CB1 and/or CB2 [197]. The most studied and thus
best

characterized

endocannabinoids

are

2-arachindonoylglycerol

(2-AG)

and

N-

arachidonoylethanolamine or anandamide (AEA), which structurally resemble arachidonic acid.

22

The production of AEA from membrane lipids requires two enzymatic steps. First, the
membrane lipid phosphatidylethanolamine is cleaved by N-acyltransferase into N-arachidonoylphosphatidylethanolamine, which is then converted by the enzyme phospholipase D into AEA.
The enzyme N-acyltransferase is dependent on Ca2+ and cAMP for optimal activity. The
production of 2-AG is mediated by two possible pathways. The first pathway involves the
production of 1,2-diacylglycerol (DAG) from the membrane lipid phosphatidylinositol (PI) by
phospholipase C (PLC). DAG is then converted into 2-AG by DAG lipase. Alternately, PI is
converted into lyso-PI by phospholipase A1 and hydrolized into 2-AG by lyso-PLC [198]. To
elicit binding activity on CB1 and CB2, it is important that these endocannabinoids have a
monosubstituted amide with an alkyl group [196].
In addition, other non-cannabis-derived non-classical cannabinoids are emerging and
synthetic ligands for CB1 and CB2 have been developed. Many of the synthetic cannabinoids
were developed as pharmacological tools, such as receptor-specific agonists and antagonists or
as potential therapeutics [199]. One synthetic ligand developed for therapeutic purposes was the
CB1 antagonist SR141716, also known as Rimonabant. An important aspect of the antagonist’s
structure-activity relationship is the amide group, which, if substituted, results in loss of activity
[196]. The pharmaceutical company Sanofi-Aventis developed Rimonabant as an anti-obesity
and smoking cessation drug. Despite pre-clinical successes of Rimonabant, the FDA halted
further progress due to reports of alterations in mood, increases in episodes of depression, and a
2-fold increased risk of suicide in patients participating in the clinical trial. The studies suggested
that cannabinoids are inherently involved in the body’s reward system and caution should be

23

Figure 2. Structures of Select Cannabinoids.
For interpretation of the references to color in this and all other figures, the reader is referred
to the electronic version of this dissertation.

24

observed using cannabinoids as therapeutics [200-202]. In addition, consumption of synthetic
cannabinoids has become a recent trend among adolescents in industrialized countries including
the US. Disguised and sold as powdered bath salts or herbal mixes for incense, these synthetic
cannabinoids are not labeled for consumption. These compounds tend to be stable derivatives of
endogenous cannabinoids and unless identified do not fall under the substance abuse laws. Due
to their synthetic nature, their effects on biological systems are unpredictable and largely
unexplored at this point [203]. For comparative purposes, some of the chemical structures of the
cannabinoid compounds mentioned above are shown in Figure 2.

C) Pharmocokinetics and Metabolism
Depending on the route of administration, inhalation or oral consumption for example,
different pharmacokinetics are observed for cannabinoids. Cannabinoids are readily absorbed
after inhalation or oral consumption and rapidly enter the bloodstream. Bioavailability of Δ9THC is on average 30% after inhalation and is much lower and more variable after oral
consumption, found between 4-12% [204, 205]. Peak levels of Δ9-THC in the plasma reach
about 200 ng/mL after smoking of a marijuana cigarette but are lower after oral administration
[206]. Initially after consumption, Δ9-THC is rapidly distributed to vascularized tissues
including the brain. Due to Δ9-THC’s lipophilic nature it is widely distributed inside the body
and especially found in adipose tissue, resulting in a two compartment (bloodstream and tissues)
distribution. Also the spleen was identified as a long-term storage site for Δ9-THC [204, 206].
Metabolism of Δ9-THC occurs in a multiphasic manner and the parent compound can be
detected at low levels more than one week after dosing [204, 205]. After entering the blood
stream, Δ9-THC is subject to first pass metabolism by the liver, which results in rapid clearance

25

of Δ9-THC from the plasma. Depending on the route of exposure, major sites for Δ9-THC
metabolism might include the lung, especially AT-II and Clara cells after inhalation as well as
intestinal epithelial cells and microbiota after ingestion [204, 207].

Due to its lipophilic

properties and redistribution to fat-containing tissues, a second much slower phase of Δ9-THC
clearance is observed [206]. Δ9-THC and other phytocannabinoids are metabolized by the P450
enzymes CYP2C9 and CYP3A4. The most common metabolites are the partially active 11hydroxy-Δ9-THC, which is further hydroxylated into 8-alpha-11-dihydroxy-Δ9-THC, the
inactive 11-nor-9-carboxy-Δ9-THC or glucoronidated by UDP-glucuronosyltransferase into the
O-ester glucuronide of THC-COOH, the main metabolite found in urine and can also be exreted
from the feces [204, 205, 208-211]. While it is easier to determine the pharmacokinetics in pure
formulations of Δ9-THC, it has been reported that CBD interferes with the metabolism of Δ9THC and thereby maintaining its plasma levels and potentiating its physiological effects [212].
Thus metabolism of cannabinoid compounds after marijuana consumption is likely slower than
after administration of pure compounds.
The production of endocannabinoids is limited to the site of cellular synthesis and in the
immune system, the main sources of endocannabinoids are activated immune cells. Thus,
endocannabinoids are thought to signal in an autocrine and paracrine fashion. Different pathways
for the metabolism of endocannabinoids exist compared to those of phytocannabinoids. AEA is
readily metabolized by fatty acid amide hydrolyse (FAAH) into arachidonic acid and
ethanolamide and 2-AG is enzymatically cleaved by monoglyceride lipase (MGL) into
arachidonate and glycerol [198]. As a consequence, inhibitors and knockout mice of FAAH or
MGL have elevated endocannabinoid levels [213-215].

26

D) Receptors for Cannabinoids
1) Cannabinoid Receptors 1 and 2
Initially there were two identified targets for cannabinoids, the G-protein coupled
receptors (GPCR) CB1 and CB2. The two receptors share limited homology (48% conserved
amino acid identity) and have different expression patterns based on cell type [196, 216]. CB1 is
translated from a single exon, while the 5’ region of the mRNA transcript contains untranslated
sequence from a second exon. Also a rare splice variant of CB1 exists, which misses part of the
coding sequence of exon 1 [196, 217]. CB2 is also translated from a single exon and also
contains an exon in the 5’ untranslated region [196]. A number of residues in the extracellular
loops of CB1 and CB2 were found to be important for agonist binding from studies performed
using human sequences of the receptors. Specifically, by using site-directed mutational analysis,
lysine 192 and glycine 195 of helix three and valine 282 of helix five were found to be important
for ligand binding in CB1 and serine 119 of helix three and phenylalanine 197 of helix 5 in CB2
[196]. CB1 was first discovered in cells and tissues associated with the central nervous system,
while CB2 was discovered in splenocytes and then identified for its immunomodulatory activity
in macrophages [216, 218, 219]. Expression of CB1 and CB2 is observed throughout the body,
including almost every organ system. Highest expression of CB1 is observed in the central
nervous system including the brain, while only glial cells (microgial cells and astrocytes) of the
brain express CB2 [196, 220-222]. On the other hand, highest CB2 expression is observed in
cells of the immune system [196]. In the immune system the expression of CB1 and CB2 varies
between the different cell types. CB1 and CB2 steady state mRNA expression levels follow a
similar pattern, but CB1 expression is lower than CB2 in the immune system. Highest steady
state mRNA levels are observed in B cells followed by NK cells, monocytes, neutrophils, CD8+

27

T cells, and CD4+ T cells [223, 224]. In addition, the expression of CB1 and CB2 can change as
a result of cellular activation, resulting in increases in CB1 in T cells, for example [225, 226].

2) Alternate Cannabinoid Receptors
In addition to CB1 and CB2, there are several other receptors that have been proposed as
targets of cannabinoid ligands. Difficulties in studying cannabinoid pharmacology have occurred
due to off-target effects present in mixtures of putative ligands and antagonists for CB1 and/or
CB2. Ligands might produce off-target effects that are not blocked by available antagonists, or
the antagonists might produce off-target effects by themselves leading to data misinterpretations.
An important model to prove the existence of cannabinoid receptors other than CB1 and CB2 is
the CB1/CB2 double knockout (CB1-/-CB2-/-) mouse. Effects by cannabinoids in CB1-/-CB2-/mice and cells isolated from these mice can be attributed to alternate cannabinoid receptors [227231]. Among the alternate cannabinoid receptors are the nuclear receptor peroxisome
proliferator-activated receptor gamma (PPARγ), the Ca2+ ion channel transient receptor
potential channel 1 (TRPC1), and the orphan receptors G-protein coupled receptor 18 (GPR18),
GPR55 and GPR119 [232]. In this laboratory, the cytoplasmic Ca2+ influx in T cells was
induced by Δ9-THC [233] and this and other Ca2+ associated signaling have been demonstrated
to be independent of CB1 and CB2 but associated with TRPC1 [229, 231, 234]. Furthermore,
using compounds such as CBD, which does not bind CB1 or CB2, suppression of T cell
activation and cytokine production including IL-2 and IFNγ was observed [235]. The
suppression of nuclear factor of activated T cells (NFAT)-dependent production IL-2 by 2-AG in
T cells was found to be a consequence of a putative COX-2 metabolite of 2-AG, 15-deoxy-

28

Table 1. Dissociation Constants (Ki) of Select Cannabinoids for CB1 and
CB2.
Shown are dissociation Ki (nM) values of CB1/CB2 receptor ligands for the
displacement of a tritiated compound ([3H]CP55940, [3H]R-(+)-WIN55212, or
[3H]HU-243) from rat, mouse, or human CB1 and CB2 receptors in vitro. A low
dissociation constant value reflects dissociation of the tritiated compounds by
low concentrations of the indicated compound. This table is adapted
fromHowlett et al. 2002 and Pertwee et al 2010.

29

Δ12,14-prostaglandin J2-glycerol ester binding to the nuclear receptor PPARγ [236]. GPR18,
GPR55 and GPR119 are recent additions to alternate CB, thus the least is known about these
orphan receptors. Endogenous lipid ligands including endocannabinoids, but also non-classical
cannabinoids activate these receptors. For example, agonists for GPR55 include the lipid
lysophosphatidylinositol (LPI) and the CB1 inverse agonist AM251 and induce neutrophil
chemotaxis [237]. GPR18 is a target for Δ9-THC and endogenous lipid mediatiors causing
migration in a human endometrial cell line [238]. Binding affinities of select cannabinoids for
CB1 and CB2 are summarized in Table 1.

VII. Intracellular Signaling by Receptors for Cannabinoids
A) Signaling by Gαi- protein coupled receptors: Cannabinoid Receptors 1 and 2
Both CB1 and CB2 belong to the GPCR family and associate with Gαi/o [239]. The
following paragraph describes the signaling initiated by CB1 and CB2, what is common among
Gαi/o-protein coupled receptors, and what might play a role in CB1 and CB2 signaling. Due to
coupling to Gαi/o, ligation of these receptors inhibits adenylate cyclase activity, resulting in
lower cytosolic cAMP levels, for example after concurrent treatment with a pharmacologic agent
such as forskolin that will induce cAMP levels [240, 241]. The βγ subunits are important in
recruiting G protein-coupled receptor kinases (GRK) which recruit β-arrestin to the receptor and
initiate β -arrestin-mediated internalization of the GPCR [242]. After ligation of CB1, for
example, β-arrestin-mediated internalization leads to tolerance to psychotropic effects after
repeated exposures to cannabinoid ligands [243]. Furthermore, the GPCR subunits βγ couple

30

Figure 3. Simplified Gαi GPCR Signaling.
For interpretation of the references to color in this and all other figures, the reader is referred
to the electronic version of this dissertation.

31

CB1 and CB2 to the mitogen activated protein kinase (MAPK) and extracellular-signal-regulated
kinase (ERK) pathways [244]. Also activation of PI3K and PLCβ by Gβγ is possible, which
activates

downstream

effectors

AKT/NFκB

and

cause

the enzymatic

cleavage

of

phosphatidylinositol 4,5-bisphosphate into inositol 1,4,5-trisphosphate (IP3), thereby inducing
intracellular Ca2+ levels from the endoplasmic reticulum stores and releasing DAG, which
activates protein kinase C [245-247]. In addition to their role in targeting GPCRs for
endocytosis, β -arrestins can form complexes with other signaling proteins thereby activating
ERK1/2 and MAPK pathways. While adapter-protein 2 and clathrin mediate the endocytosis of
GPCRs after binding to β -arrestin, interaction of β -arrestin with members of the Src family of
kinases results in MAPK activation and/or internalization of the GPCR. Also, interaction of βarrestin with apoptosis signal-regulating kinase 1 (ASK1), c-Jun N-terminal kinase 3 (JNK3),
and mitogen-activated protein kinase kinase 4 (MKK4) leads to the activation of the JNK
pathway [248, 249]. While this has been demonstrated for other Gαi-coupled GPCRs, it is likely
that the same occurs for CB1 and/or CB2 although direct evidence is lacking [250]. Also,
regulator of G protein signalling (RGS) proteins exist and as their name suggests, RGS are
involved in the deactivation (or regulation) of GPCR signaling by facilitating the hydrolysis of
guanosine triphosphate to guanosine diphosphate. More recent findings have demonstrated that
in addition to regulating GPCR signaling at the receptor, the RGS proteins are able to act on
downstream targets of the GPCR cascade to regulate GPCR signaling [251]. The putative
signaling pathways activated by CB1/CB2 are summarized in Figure 3. Despite the information
on the downstream signaling of CB1 and CB2 it is currently unknown to what extent and how
each pathway leads to the suppression of immune responses associated with cannabinoid
treatment. To add further complexity to the signaling by CB1 and CB2, heterodimerization with

32

other GPCRs has been reported with CB1 and is common among the GPCR family and thus
likely to occur with CB2 as well [252, 253]. It is currently unknown what the heterodimerization
partners of CB1 and CB2 are and what putative changes in signaling result from
heterodimerization. There are reports that for some GPCRs, up to 80% of exist as homodimers
and as a result reduced activation of receptor-mediated downstream signaling occurs [253, 254].

B) Signaling by Alternate Cannabinoid Receptors
While GPR18, GPR55, GPR119 and TRPC1 are surface receptors, PPARγ is cytosolic
and requires its ligands to cross the cell membrane. Ligation of GPR55 results in activation of
Rho-family GTPases, also activation of MAPK/ERK pathways has been demonstrated. As a
result of Rho activation, intracellular Ca2+ release was observed, which led to activation and
nuclear translocation of NFAT [255]. GPR119 is a Gαs-coupled receptor and induces
intracellular cAMP levels upon ligation. In addition, Ca2+ levels are increased as a result of
downstream effects of cAMP on ATP-sensitive potassium channels and voltage-dependent
calcium channels [256]. Currently, no experiments have been performed to address the
intracellular signaling downstream of GPR18. TRPC1 participates in the store operated calcium
entry and thereby is involved in cell activation as a result of Ca2+ influx, for example occurring
during activation of T cells [257]. The receptor responds to depletion of intracellular Ca2+ stores
and allows for influx of extracellular Ca2+ to allow for sustained Ca2+ levels for optimal
activation [258]. Upon ligand binding, PPARγ heterodimerizes with retinoid X receptor alpha
(RXRα) and translocates to the nucleus to induce the transcription of its target genes. In addition,
PPARγ interacts with intracellular signaling directly, for example to reduce NFAT-mediated
induction of IL-2 expression in T cells [236]. Also cross-talk between PPARγ and Janus tyrosine

33

kinase-signal transducer and activator of transcription (JAK-STAT) signaling exists and ligation
of PPARγ results in the reduction of transcriptional activity of STAT5b [259].

VI. Effects of Cannabinoids on Immune Function
A) Effects on Antigen Presenting Cells
It has been demonstrated that Δ9-THC inhibits APC function including antigen
processing and presentation, and cytokine production [260]. The induction of TNFα by LPS and
IFN-γ was suppressed by Δ9-THC in human and murine macrophages. Notably, culture serum
concentrations of 5% abrogated the effect of Δ9-THC on induced TNFα production [261]. To the
same extent LPS-induced NO production by the RAW264.7 macrophage cell line and peritoneal
macrophages was decreased by Δ9-THC after treatment in vitro and was attributed to the
suppression of NFκB/Rel and CREB/ATF binding and reduction of intracellular cAMP levels
[262]. In addition, chemotaxis of macrophages to RANTES was reduced by Δ9-THC in a CB2dependent manner by the use of a specific antagonist [263]. Furthermore, depending on the
activation state of macrophages, different levels of induction of CB2 transcript were observed.
The highest induction was observed in macrophages with inflammatory functions, suggesting a
role for CB2 in a negative feedback loop to down-regulate inflammation in macrophages [264].
It has been observed that both CB1 and CB2 contribute to the modulation of DC function
by Δ9-THC with the use of CB1- and CB2–specific antagonists [265] and that CB1 specifically
diminishes K+ outward currents through Kv channels to suppress the ability of DC to elicit T
cell resonses [266]. Furthermore, suppression of DC function by Δ9-THC might, in part, be
independent of the G-protein coupled receptors as evidenced by Δ9-THC-mediated suppression

34

of maturation after in bmDC generated from CB1-/-CB2-/- mice. In support of this argument,
pertussis toxin, which blocks the Gαi subunit necessary for CB1 and CB2 signaling [267, 268],
did not completely abrogate the suppression of IL-12p40 production in bmDC [269]. Also
selective antagonists to either CB1 and CB2 were not able to block the suppression of IL-12p40
by Δ9-THC [270]. This suggests that although CB1 and CB2 are involved in Δ9-THC
modulation of DC, this might not be the exclusive mechanism. Δ9-THC has also been found to
induce suppressive MDSC populations, providing another putative mechanism by which Δ9THC suppresses immune function [271].

B) Effects on Granulocytes
Most of the studies of the effects of cannabinoids on granulocytes have been focused on
neutrophils. There are conflicting reports about the involvement of CB1 and CB2 and the modus
of the outcome of neutrophil function. It has been demonstrated that cannabinoids impair
neutrophil chemotaxis independent of CB1 and CB2 [272]. Others argue that CB2 plays an
important role in positively regulating the migration of neutrophils [273]. Furthermore,
neutrophil superoxide generation was suppressed by the non-selective CB1 and CB2 agonist CP
55,940 independent of CB1 and CB2, suggesting alternate targets [274]. In contrast, others have
shown a CB2-dependent stimulation of neutrophil function with endogenous and synthetic
cannabinoids [275].

C) Effects on CD4+ T cells
Due to the critical role of proliferation in CD4+ T cell responses and the requirement of
IL-2 for proliferation, which is rapidly and robustly induced after T cell activation, much of the

35

research on the immunomodulator action of cannabinoids have focused on proliferation, IL-2
production and their upstream signaling pathways. The first evidence for effects mediated by
cannabinoids on T lymphocytes came from in vivo sheep erythrocyte assays in rats, where Δ9THC treatment impaired the T cell dependent antibody formation against erythrocytes [276]. In
1977, Δ9-THC and its primary metabolite 11-OH-Δ9-THC were shown to suppress the
proliferation as measured by radioactive thymidine incorporation of phytohemagglutininactivated human T cells in vitro, which was confirmed by a second group [277, 278]. After
confirmation of the previous results it was also noted that no cytotoxicity was observed by Δ9THC on T cells with concentrations as high as 26 µM, but higher serum concentrations (20% of
culture media) reduced the suppressive effect of Δ9-THC on proliferation, suggesting
neutralization of Δ9-THC by serum proteins [278]. While not changing the percentage of CD4+
T cells in the spleen, Δ9-THC attenuated proliferation after pharmacological activation with
PMA/Io and in allogeneic mixed lymphocyte responses as well as IL-2 and IFNγ production
after PMA/Io activation [279]. In agreement with the Δ9-THC-mediated attenuation of
proliferation, Δ9-THC decreased the surface expression of IL-2 receptors with high affinity,
while increasing IL-2 receptors with low affinity [280]. The mechanism by which Δ9-THC
reduced IL-2 expression in T cells was explored in the T cell line EL-4.IL2 and it was reported
that Δ9-THC reduced the binding of AP-1 to the proximal promoter region of IL-2 [281]. Using
cannabinol, a weaker selective CB2 agonist than Δ9-THC, suppressed AP-1 binding by
decreasing nuclear presence of its protein components c-fos and c-jun. In addition, reduced
NFAT binding was observed as a result of cannabinol treatment leading to reduced production of
IL-2 [282, 283]. Moreover, the phosphorylation of c-fos and c-jun and the phosphorylation of
ERK1 of the MAPK/ERK pathway were reduced by cannabinol [283]. In light of the critical role

36

of Ca2+ in the activation of NFAT, the focus shifted to modulation of intracellular Ca2+ by
cannabinoids. Studies in this laboratory demonstrated that Ca2+ influx was temporally
dysregulated by Δ9-THC in T cells through the TRPC1 receptor [231, 233, 284]. More recently,
it has been demonstrated by this laboratory that Δ9-THC suppresses the induction of the coactivating molecule ICOS on CD4+ T cells after activation and it was suggested that this occurs
in part via the modulation of NFAT signaling [285]. Furthermore, it has been argued that with
certain stimulations, Δ9-THC shifts the polarization of the Th response to a Th2 phenotype
[286]. Moreover, there is evidence that the effects of cannabinoids on T cells are independent of
CB1 or CB2 which is in agreement with low expression levels for these receptors [231, 233, 234,
287]. In contrast, it has been demonstrated in the Jurkat T cell line that expression of CB1
increases after immune stimulation, suggesting that CB1 may act to suppress activity in activated
T cells [288].

D) Effects on CD8+ T cells
In CD8+ cytotoxic T lymphocytes (CTL), it was first noticed that proliferation was
suppressed in T cell subsets by Δ9-THC [289]. Cytolytic activity was attenuated in vitro by Δ9THC in CTL. The mechanism was proposed to be bimodal: 1. during differentiation and
proliferation of pre-cursor CTL and 2. during cytolytic activity, inhibiting target cell lysis [290].
Furthermore, in vivo treatment of Δ9-THC reduced CTL activity from mouse splenocytes in a
model of herpes simplex virus type-1 (HSV-1). Using microscopy, it was observed that
polarization towards, but not binding, to virus-infected target cells was inhibited [291].

E) Effects on NK cells

37

There is evidence that cannabinoids impair NK cell activity. Both Δ9-THC and the
primary metabolite 11-OH-Δ9-THC were able to suppress target cell lysis by NK cells [292].
Also, it was observed that while the lytic activity was suppressed by Δ9-THC the binding to
target cells was not affected [293]. The migration of NK cells was induced by the
endocannabinoid 2-AG in a CB2-dependent manner, while Δ9-THC failed to induce migration
[294]. The production of the proinflammatory cytokine TNF-α was suppressed by Δ9-THC in
NK cells activated with C. albicans [295].

F) Effects on B cells
Among the immune cells B cells express higher levels of CB2, suggesting that this receptor
mediates some of the effects associated with cannabinoid treatment. Cannabinoids have
differential effects on B cells. B cell growth is enhanced at low nanomolar concentrations of Δ9THC and synthetic cannabinoids and was abrogated by pertussis toxin, suggesting a GPCRdependent mechanism, which the authors attributed to CB2 expression in B cells [296].
Futhermore, CD40-induced proliferation was enhanced in a CB2-dependent manner in B cells by
the use of a CB2-specific antagonist. Also upregulation of CB2 occurred in B cells 24 hours after
CD40 ligand-induced proliferation [225]. Studies from this laboratory demonstrated that
endocannabinoids enhanced LPS-induced B cell proliferation at high cell densities, but
suppressed proliferation at low cell densities [297]. Moreover, the antibody forming cell
response to sheep red blood cell antigen was suppressed by Δ9-THC and the suppression was
abrogated by the addition of the cell permeable cAMP analog dibutyryl-cAMP, suggesting a
mechanism involving a Gαi-coupled GPCR, putatively CB2 [267]. Using the B cell line SRIH-B,
it was demonstrated that Δ9-THC enhanced IL-8, but suppressed MIP-1β production [298].

38

G) Effects on Immune Outcome in Host Resistance Models
Although the effects of Δ9-THC on isolated immune cells in vitro have been extensively
investigated, the effects of Δ9-THC on the immune system in vivo are more complex. Factors
contributing to this complexity include the type of immune stimulus, tissue location, as well as
the interplay between, and kinetics of, the cellular arms of immune response during an
immunological insult, none of which can be completely capitulated in vitro. To date, few studies
have focused on the effects of Δ9-THC on multiple immune cell populations using a pathogen
challenge model [119, 120, 286, 299-305].
The first studies of the effects of cannabinoids on host resistance were performed using
mouse models of infection with HSV-2 and the intracellular pathogen Listeria monocytogenes
[302]. Both pathogens induce strong Th1 immune responses in the host resulting in control
(HSV-2) or clearance (L. monocytogenes) of the pathogen by CTL when infected with lower
innoculums [306]. Δ9-THC decreased the host resistance to both pathogens as evidenced by
mice succumbing to lower innoculums of HSV-2 and L. monocytogenes compared to vehicle
(VH) treated controls. In a follow-up study using the HSV-2 model, Δ9-THC suppressed host
immune function to increase host mortality and viral titers after HSV-2 infection and Δ9-THC
treatment resulted in lower serum complement fixing and antibody neutralization titers compared
to VH [299]. Using the same model, it was subsequently determined that Δ9-THC reduced the
type I IFN response and previous studies in mice were confirmed in guinea pigs [300, 305].
In studies using a Legionella pneumophila model, Δ9-THC suppressed primary and
secondary immunity to the pathogen. L. pneumophila-induced serum levels of Th1 cytokines
IFNγ and IL-12 and splenic steady state mRNA levels of IL-12 receptor beta were attenuated by

39

Δ9-THC treatment. The effects by Δ9-THC were attributed in part to a shift from Th1 to Th2
immunity as increased serum levels of IL-4 and steady state mRNA levels of GATA-3, a key
Th2 transcription factor, were observed in the spleen. Furthermore, it was noted that both CB1
and CB2 contributed to the effects of Δ9-THC as determined by the use of receptor antagonists
and knockout mice [286, 303, 304].
Using a lung cancer model, Δ9-THC impaired immune responses to tumor implants,
which underwent accelerated growth as a result of Δ9-THC treatment. While the Th1 cytokine
IFNγ was suppressed by Δ9-THC, the immunosuppressive cytokines IL-10 and TGF-beta were
enhanced at the tumor site and in the spleens in mice receiving Δ9-THC treatment. In these
studies a CB2 antagonist was effective in blocking the suppressive effects of Δ9-THC, thus
leading the authors to conclude a CB2-dependent mechanism of suppression of anti-tumor
immunity [301].
Most recently, studies were performed to determine changes in host resistance due to Δ9THC treatment using a murine influenza virus host challenge model characterized in this
laboratory [118]. While Δ9-THC reduced the lung tissue pathology as well as the influx of
macrophages, CD4+, and CD8+ T cells into the airways, higher lung viral burden was observed
[119]. Using CB1-/-CB2-/- mice, it was observed that these mice had drastically lower lung viral
burden and greater protein levels of TNFα in the BALF. Furthermore, the Δ9-THC-mediated
reduction in lung tissue pathology was dependent on CB1 and/or CB2. While the immune
response to influenza was broadly characterized, it was unclear to which extent different immune
cell populations contributed to the observed endpoints.

H) Phenotype of the Immune System as a Result of Loss or Lack of CB1 and/or CB2

40

There are a multitude of studies examining the immune system of mice after genetic
ablation of CB1 and/or CB2 and humans with identified single nucleotide polymorphisms
(SNPs) in CB1 or CB2. Consistent among these studies is an enhanced immune response
phenotype with extinguished or reduced signaling through CB1 and/or CB2, either resulting in
exacerbation of the disease state in mouse models or increased propensity for developing
autoimmune disease in humans. Mice with a single deletion of CB1 or CB2 or CB1-/-CB2-/mice are healthy, fertile and of similar size and weight compared to WT mice. However,
increased mortality was observed in mice lacking CB1 [228, 307].
Much of the work of studying the phenotype of CB1-/- mice in the immune system has
focused on intestinal inflammation models [308, 309]. Intrarectal infusion of 2,4-dinitrobenzene
sulfonic acid and oral administration of dextrane sulfate sodium was used to induce colitis in WT
and CB1-/- mice, intestinal inflammation was exacerbated in CB1-/- compared to WT mice. The
authors attributed the exacerbation to the lack of signaling through CB1 myenteric neurons and
lack of CB1-mediated control of inflammation-induced irriation of intestinal smooth muscle cells
[309]. This study highlights the potential involvement of CB1 via the central nervous system
affecting the peripheral organ systems. Signaling at the nerve terminal of the input neuron is
believed to be reduced by postsynaptic release of endocannabinoids and ligation of CB1 on the
input neuron, thereby reducing the action potential of the input neuron [310]. Thus ablation of
CB1 could potentially lead to an overactive nervous system, which can lead to enhanced immune
function as is experienced with angiotensin II release after sympathetic nervous system
stimulation [311]. However no further characterization of the effects of the nervous system on
the immune system was performed. In addition, reduced endotoxin hypotension was observed in

41

CB1-/- or CB1-/-CB2-/- mice compared to WT mice, suggesting a stronger response to LPS in
the absence of CB1-/- [227].
While the lack ofCB2 alone or lack of both CB1 and CB2 in mice has an effect on the
percent composition of splenic immune populations [279, 312], CB2 deletion resulted in
abnormal formation of subsets of B and T cells compared to WT mice [313]. In particular, CB2/- mice showed a deficiency by percent composition of intestinal intraepithelial NKT cells and a
reduction in the percentages of marginal zone B cells and memory CD4+ T cells in the spleen.
Furthermore, peritoneal B1a B cells, a subset of B cells that do not contribute to memory
formation, was slightly reduced [313]. Additionally, deletion of CB2 resulted in accelerated agerelated loss of bone mass compared to WT, which was characterized by a phenotype similar to
osteoporosis in humans. The authors speculated that the loss of CB2 might have resulted in an
increase in proresorptive cytokines such as IL-1 and TNF-α, as these are reduced by CB2
ligation [314].
CB1-/-CB2-/- mice were generated by crossbreeding previously generated CB1-/- [307]
and CB2-/- [312] mice and the use of CB1-/-CB2-/- mice in experiments has been limited to a
few studies [120, 227-229, 231, 279, 315, 316]. In studies from this laboratory CB1-/-CB2-/mice were used mostly as a tool to demonstrate that cannabinoid-mediated suppression of T cell
responses was independent of CB1 and CB2. Only few studies have described immune related
endpoints in CB1-/-CB2-/- mice that offer insights into phenotypic differences compared to WT
mice, with three of the studies coming from this laboratory. First, Kaplan et al. showed a trend
towards increased release of IFNγ from pharmacologically stimulated CB1-/-CB2-/- compared to
WT splenocytes, albeit not evaluated statistically [229]. Karsak et al. described the involvement
of the endocannabinoid system in the amelioration of hypersensitivity responses to contact

42

sensitizers. In addition to greater ear swelling and more pronounced lesions in ear tissue,
predominantly chemokines and chemokine receptors were found to be upregulated in CB1-/CB2-/- mice compared to WT mice [315]. Springs et al. demonstrated that there was no
difference in the percent composition of immune cells in the spleens of WT and CB1-/-CB2-/mice. Moreover, although not deemed statistically significant, slightly greater lymphocyte
proliferation as a result of pharmacological stimulation as well as enhanced IL-2 production after
pharmacological activation was observed in splenocytes from CB1-/-CB2-/- mice compared to
WT mice in vitro. Also the response to sheep red blood cell antigen was enhanced in vivo in
CB1-/-CB2-/- compared to WT mice [279]. Most recently, a study from this laboratory by
Buchweitz et al. showed that the viral lung burden is reduced in CB1-/-CB2-/- compared to WT
mice after influenza infection. Furthermore, increases in BALF CD4+ T cell counts were
observed in CB1-/-CB2-/- compared to WT mice [120]. On the other hand, in an adjuvant-free
ovalbumin model of airway hypersensitivity, there were no overt differences in the airway
responses to ovalbumin. However, the presence of neutrophil populations was enhanced at 96 h
post challenge. Due to the absence of an inflammatory stimulus such as adjuvant in this model, it
suggested the necessity for an inflammatory stimulus to observe enhanced immune reactivity in
CB1-/-CB2-/- compared to WT mice [317]. Taken together, these studies demonstrate an
inherent role for CB1 and CB2 in the regulation of the magnitude of the immune response after
an inflammatory stimulus.
In humans, SNPs are often associated with a loss or lack of function of the target gene
product and have been observed in CB1 and CB2 [318, 319]. A population was identified with a
SNP in the CB1 transcript Cnr1, 1359 A/A to G/A, resulting in a silent mutation in CB1 at
position 454 threonine. The G/A population and a control (A/A) population were evaluated for

43

ulcerative colitits (UC) and Crohn’s disease. There was a significant association of the G/A
population with UC susceptibility, demonstrating a potential role for CB1 in UC. Due to the
apparent effect as a result of a silent mutation, the authors argued that CB1 splicing or RNA
stability could have been altered due to the SNP resulting in the observed phenotype [319]. In a
second study, a mutation in CB2 cDNA at position 188-189 from AA/AA to GG/GG resulted in
the substitution of glutamine into arginine at amino acid position 63 of the CB2 protein. As a
functional consequence, the inhibition of T cell proliferationin the CB2 GG/GG variant by the
cannabinoids 2-AG and N-arachidonlyglycine was reduced. Furthermore the GG/GG genotype
was significantly overrepresented in individuals diagnosed with autoimmune disease including
multiple sclerosis, rheumatoid arthritis, systemic lupus, and myasthenia gravis [318]. Overall,
these studies suggest that the cannabinoid system might be involved in preventing autoimmune
disease by modulation of immune function through the cannabinoid receptors.

VI. Objectives
The objective for the studies presented as part of this dissertation was to test the
hypothesis that Δ9-THC suppresses immune responses generated by influenza virus by
perturbing an endogenous signaling pathway relying in part on the presence of the GPCRs CB1
and/or CB2. While CB1 and/or CB2 were deemed important targets of endocannabinoids and
Δ9-THC in the impairment of APC-mediated immune responses, suppression of CTL responses
was independent of CB1 and CB2. First, the effect of Δ9-THC on the generation of CTL and its
CB1 and CB2 dependence was examined. These studies included the characterization of
functional consequences of Δ9-THC on CTL effectors as well as the involvement of early

44

signaling of T cells in an attempt to shed light on a putative mechanism by which Δ9-THC
perturbed the generation of CTL. Second, a kinetic study of influenza infection spanning
including 1, 3, 5, and 7 dpi in WT and CB1-/-CB2-/- mice was undertaken to: 1. determine the
peak time of the inflammatory immune response in WT and CB1-/-CB2-/- mice and 2.
investigate the cellular events leading to greatly reduced viral burden in CB1-/-CB2-/- compared
to WT mice [120]. Included in these studies were global analyses of steady state mRNA level
changes in the lung, analyses of lung tissue for pathology, and extensive ex vivo and in vitro
studies to identify immune cells potentially involved in the disparity of the viral burden in CB1/-CB2-/- compared to WT mice. Third, the effect of Δ9-THC on the generation of an immune
response to influenza infection was evaluated in WT and CB1-/-CB2-/- mice at 3 dpi, the peak
day of the inflammatory and highly APC-involved immune response. Similar approaches to the
previous studies were used, including evaluating steady state mRNA levels and extensive
immune cell response characterization using ex vivo analysis and in vitro models. The results of
these studies indicate that signaling through CB1 and/or CB2 is important in maintaining
immune homeostasis and in response to an immune stimulus, such as influenza virus, preventing
excessive immune reactivity. Specifically CB1 and/or CB2 are involved in regulating APC
function and loss of signaling through these receptors leads to increased inflammatory responses
and immunopathology after influenza infection. In particular DC, including cDC and pDC, are
regulated by signaling through these two receptors as their immune functions are exacerbated in
the absence of CB1 and CB2 signaling. Likewise, DC are very sensitive to Δ9-THC-mediated
immunosuppression via CB1 and/or CB2 as observed in the suppression of maturation and the
ability to elicit T cells in vitro. Moreover, T cell responses including CTL, are suppressed by Δ9-

45

THC independent of the expression of CB1 and CB2, suggesting alternate mechanisms for
immunomodulation by Δ9-THC for T cells.

46

MATERIALS AND METHODS

I.

Mice.
Mice were housed at Michigan State University (MSU) in pathogen-free animal housing

facilities at 21 to 24°C and 40 to 60% relative humidity with a 12 hour (h) light/dark cycle.
Water and Lab Chow (Nestlé Purina, St. Louis, MO) were provided ad libitum. All protocols
and procedures were performed in accordance with guidelines set forth by the MSU Animal Care
and Use Committee. CB1-/-CB2-/- mice on a C57Bl/6 genetic background were a kind gift of
Dr. Andreas Zimmer at the Universität Bonn, Germany [315] and were maintained at MSU and
C57Bl/6 (WT) mice were ordered from NCI. Mice were used for experiments between 8-12
weeks of age. CB1-/-CB2-/- mice were genotyped to ensure the targeted deletion of both
receptors. It is notable that primers from Applied Biosystems were not able to differentiate cnr2
gene expression between WT and CB1-/-CB2-/- mice, and therefore custom primers were used
[320]. T cell receptor transgenic mice with CD8+ T cells recognizing the peptide OVA257-264
(OT-1) were purchased from Jackson Laboratory (Bar Harbor, ME) and used at 8 weeks of age.

II.

Chemicals and Reagents
Δ9-THC was obtained from the National Institute on Drug Abuse (Bethesda, MD).

Ethanol was purchased from Decon Labs (King of Prussia, PA). Ionomycin (Io) and
dimethylsulfoxide (DMSO) were purchased from Sigma Chemical Co. (St. Louis, MO). RPMI
media was obtained from Gibco Invitrogen (Carlsbad, CA), and 51Cr as sodium chromate was
obtained from Perkin Elmer (Waltham, MA).

47

III.

T Cell Elicitation for Generation of Functional CTL
C57Bl/6 (WT) and CB1-/-CB2-/- mice were euthanized, the spleens isolated in a sterile

environment, and splenocytes (SPLC) enumerated using a Coulter Counter (Beckman Coulter,
Brea, CA). Cells from the mouse lymphoblast-like mastocytoma cell line P815 were irradiated
with 3000 rads to prevent proliferation, washed 3 times with RPMI and counted using a
hemacytometer. SPLC and irradiated P815 cells were combined at 1x10^6 and 1x10^5 cells,
respectively, in RPMI 1640 supplemented with 5% bovine calf serum (BCS) in a total volume of
200 µL in a round bottom 96 well plate. The plates were incubated in a humidified incubator
with 5% CO2 at 37°C for the indicated amounts of time.

IV.

Drug Treatment in vitro
At the time of co-culture of SPLC and irradiated P815, Δ9-THC (1, 5, 10 µM), vehicle

(VH, 0.1% ethanol) or RPMI (NA) was added. All Δ9-THC treatments had the same ethanol
content (0.1%) as vehicle control.

V.

51Cr Release Assay
After elicitation, cells were harvested and washed twice with RPMI 1640 media without

serum. P815 cells were washed once and 1x10^6 cells were incubated in the presence of
Na251CrO4 for 1 h in 10% BCS supplemented RPMI 1640 media in a volume of less than 50
µL. After incubation P815 cells were washed 3 times using RPMI 1640 media without serum.
51Cr-labelled P815 cells were adjusted to 1x10^5 cells/mL in 2% BCS RPMI media. Elicited
CTL were adjusted in 2% BCS complete RPMI media to ratios ranging from 50 (5x10^5) to 1
(1x10^4 cells): 1 (1x10^4) P815 cells, depending on the experimental design, in a volume of 200

48

µL. After co-culture, elicited CTL and P815 were added to a 96-well round bottom plate and
centrifuged at 200g for 1 min to force cellular interactions. Control wells for spontaneous release
(200 µL of P815 only) and total release (1% Triton-X 100 in 200 µL of P815 cells in RPMI)
were used to determine the range of experimental release. After 5 h of co-culture in a humidified
incubator with 5%CO2 at 37°C, cell lysis was assessed by aliquoting 100 µL of supernatant from
each well, which represents the experimental release. The cytolytic activity was calculated as
follows: % Release = (experimental release – spontaneous release) / (total release – spontaneous
release) x 100.

VI.

IFNγ T Cell Functional Analysis
CTL were elicited as described above. After 5 days, cells were harvested and co-cultured

with P815 at a ratio of 10:1 (see above) for 12 h in the presence of brefeldin A to prevent IFNγ
release and allow for detection by fluorescently labeled antibody. After co-culture, cells were
prepared for fluorescent antibody staining (described below).

VII.

Proliferation Assay

Prior to elicitation, SPLC were incubated with Cell Trace carboxyfluorescein
succinimidyl ester (CFSE) dye (Invitrogen, Carlsbad, CA) according to manufacturer’s
instructions. CTL were elicited as described above for generation of CTL. Dilution of dye
staining is indicative of proliferation and staining profiles from elicited samples were compared
to a 24 h SPLC only control, incubated without P815.

VIII. T Cell Activation Assays

49

CTL were elicited as described for generation of CTL; however, RPMI supplemented
with 2% BCS was used instead of 5%. For direct stimulation, 0.5 µM ionomycin (Io) dissolved
in 0.01% DMSO in RPMI was used as the Ca2+ ionophore to induce intracellular Ca2+ levels
and thereby increasing CD69 expression. After 6 h, CD69 surface staining was performed using
antibodies specific for CD69 and after 24 h for CD25 (staining described in detail below).

IX.

Procedure for Immunofluorescent Staining of CD4+ and CD8+ Cells for Flow
Cytometry
CTL were elicited as described above. Cells were washed in 96-well round bottom plates

with HBSS once and incubated with Near IR LIVE/DEAD (Invitrogen) dye to assess viability
according to manufacturer’s instructions. This dye stains cells that have lost membrane integrity.
Cells were washed twice with FACS buffer (1X HBSS, 1% bovine serum albumin (BSA), 0.1%
sodium azide). Subsequently, Fc receptors were blocked using Purified Rat Anti-Mouse
CD16/CD32 (BD Pharmingen, San Diego, CA). To stain surface proteins, cells were incubated
for 20 min with 0.25-0.5 µg of CD4 (clone RM4-5, Biolegend, San Diego, CA) and CD8 (536.7, Biolegend). For the T cell activation experiments, cells were additionally stained with CD69
(H1.2F3, Biolegend) and CD25 (PC61, Biolegend). Cells were then washed twice and fixed with
Cytofix (BD, Franklin Lakes, NJ). After re-stimulation (described above), on the day of flow
cytometric analysis, cells were washed once, then permeabilized with Cytoperm (Becton,
Dickinson and Company) by incubation for 20 min and stained with IFNγ (XMG1.2, Biolegend)
for 30 min. All experiments included single stain controls to compensate for fluorescence
overlap between detectors. Samples were individually analyzed for percent positive staining as

50

defined by a gate to perform statistical analysis and concatenated by treatment group for
graphing using FlowJo v.8.8.6. for Macintosh.

X.

Influenza instillation.
On the day of instillation, murine-adapted influenza virus A/PR8/34 (a kind gift of Dr.

Allen Harmsen, Montana State University) was diluted in 0.9% NaCl (saline) solution (SAL)
(Hospira, Lake Forest, IL) to 1 plaque forming unit (pfu) per µL. 25 µL were instilled per nostril
resulting in a total inoculum of 50 pfu per mouse. Mice (n=5 each genotype) received either
saline or A/PR8/34 under isofluorane anesthesia.

XI.

Necropsy and lung tissue collection.

Mice were anesthetized by intraperitoneal injection of 250 mg/kg pentobarbital (Fatal-Plus,
Vortech, Dearborn, MI) at 1, 3, 5, and/or 7 days post infection (dpi) and euthanized by
exsanguination of the abdominal aorta. Lungs were excised en bloc.

XII.

BALF analysis and differential cell counts.

Saline (2 x 0.9 mL) was used to lavage lungs and collect the broncho-alveolar lavage fluid
(BALF). Total cell counts were obtained by counting an aliquot of the total obtained BALF
using a hemacytometer. Differential cell counts were prepared by centrifuging BALF onto glass
slides using a Shandon Cytospin 3 centrifuge (Block Scientific, Nutley, NJ) at 600 rpm for 10
minutes. The slides were stained using the Diff-Quik kit (Dade Behring, Newark, DE) following
manufacturer’s instructions. The stained slides were counted for at least 200 discernable cell

51

entities including lymphocytes, eosinophils, macrophages and neutrophils, and percentages were
calculated. The total cell count was used to enumerate differential cell populations.

XIII. Histological and immunohistochemical staining of lung tissue sections.
After en bloc excision, left lung lobes were fixed and inflated under constant pressure (30
cm H2O) with 10% phosphate buffered formalin for one hour. After transverse cuts at airway
generation five, tissue sections were paraffin embedded and cut into 4 µM sections. Tissue
sections were either histochemically stained with hematoxylin and eosin (H&E) using a standard
procedure or were immunohistochemically stained with antibody against clara cell secretory
protein (CCSP). Glass slides (Fisher, Waltham, MA) were coated with 2% 3aminopropyltriethoxysilane and tissue sections were mounted onto slides and dried at 56°C
overnight. Xylene was used to deparaffinize the slides, which were hydrated with decreasing
concentrations of ethanol to distilled water. Subsequently, the pH was adjusted by soaking slides
in Tris Buffered Saline (TBS) pH 7.4 (Scytek Labs, Logan, UT) for 5 min. Following pH
adjustment for CCSP staining, standard avidin-biotin complexing for staining was performed at
room temperature using a DAKO Autostainer. All staining steps were performed using TBS with
Tween 20 (Scytek Labs) for rinses. In short, non-specific protein was blocked using Normal
Goat Serum (Vector Labs, Burlingame, CA) for 30 min followed by incubation with
Avidin/Biotin blocking system for 15 min each (Avidin D, Vector Labs / d-Biotin, St. Louis,
MO). The goat serum blocked slides were incubated with primary polyclonal rabbit anti-CCSP
antibody diluted 1:1,600 (Seven Hills, Cincinnati, OH) in Normal Antibody Diluent (Scytek).
Biotinylated goat anti-rabbit IgG (H + L) was diluted to 11.0µg/ml in NAD and incubated for 40
min followed by incubation with R.T.U. Alkaline Phosphatase ABC Reagent (Kirkegaard-Perry

52

Labs, Gaithersburg, MD) for 60 min. For the chromatic reaction Vector Substrate Kit I and
counter stain in Gill Hematoxylin (Thermo Fisher, Kalamazoo, MI) for 15 seconds was used.
Differentiation, dehydration, clearing and mounting was performed with synthetic mounting
media.

XIV. Isolation of alveolar macrophages from BALF.
BALF was obtained as described above from naïve WT and CB1-/-CB2-/- mice. Cells
were washed twice and counted with a Coulter Counter and incubated at 0.5x10^5 cells/mL in 2
mL in a 12 well dish. BALF cells were stimulated with and without (naïve, NA) LPS at 1 µg/mL
for 24 hours. To assess maturation, MHC I, MHC II, and CD86 staining was performed in
addition to CD11b and CD11c to gate on macrophage populations. Analysis of fluorescence
intensity of maturation markers was exclusively performed on the alveolar macrophage
populations (CD11bmidCD11chi), which constituted the majority of cells obtained from the
BALF.

XV.

Flow Cytometry on lung isolated cells.

From a second set of mice, lung tissue from left and right lung lobes was homogenized
using a cell dissociation sieve kit (CD-1, Sigma, St. Louis, MO), connective tissue was removed
and cells were immersed in RPMI media. For cytokine production analyses, cells were
restimulated in vitro with phorbolmyristate acetate (PMA) and Io (both from Sigma) at 40 nM
and 0.5 µM, respectively, for 5 h in the presence of Brefeldin A (Biolegend) in 2% BCS RPMI
media supplemented with 100 U/mL penicillin and 100 µg/mL streptomycin (all from GIBCO

53

Invitrogen, Carlsbad, CA). Cells isolated from lung or restimulated in vitro were washed 3 times
with FACS buffer (1 X Hank’s Buffered Saline Solution, 1% bovine serum albumin, 0.1%
sodium azide, pH adjusted to 7.6) and Fc receptors were blocked using purified rat anti-mouse
CD16/CD32 (BD Pharmingen, San Diego, CA). Surface markers were stained with specific
antibodies, using concentrations of 0.25-0.5 µg per 1x10^6 cells. FACS antibodies used were all
from Biolegend (San Diego, CA): CD4 (clone RM4-5), CD8 (53-6.7), CD69 (H1.2F3) and
NK1.1 (CD161, PK136). For lung APC populations the following antibodies (all from
Biolegend) were used: CD11b (M1/70), CD11c (N418), CD80 (16-10A1), CD86 (GL-1), Gr-1
(RB6-8C5), MHC I (H-2Kb/H-2Db, 28-8-6), and MHC II (I-A/I-E, M5/114.15.2). Cells were
fixed with Cytofix (BD, Franklin Lakes, NJ) before flow cytometric analysis. Intracellular
staining for IFNγ (XMG1.2) and IL-17 (TC11-18H10.1) was performed using 1 X Perm/Wash
(BD) for washes and incubation according to manufacturer’s instructions. Samples were
prepared in 96 well round-bottom plates and were analyzed using a BD Biosciences FACSCanto
II flow cytometer. Obtained data were analyzed individually and concatenated for graphical
representation with FlowJo v8.8.6 (Treestar Ashland, OR) for Mac.

XVI. mRNA extraction and cDNA synthesis.
Lung tissues from right lung lobes were minced with a dounce tissue grinder set (Sigma)
and RNA extracted from right lung tissue using TRIzol reagent (Sigma) according to
manufacturer’s instructions. Extracted mRNA was reverse transcribed into cDNA using 2.5 µg
total RNA in a 50 µL reaction volume with a High Capacity cDNA Archive Kit (Applied
Biosystems, Foster City, CA) according to manufacturer’s instructions.

54

XVII. Mouse immune panel – low density microarray.
Mouse low density microarray immune panels were obtained from Applied Biosystems
andcontained primers for 93 immune-related and three housekeeping genes using the TaqMan
PCR system. The panels were loaded with 50 ng of cDNA/well and centrifuged two times for 1
minute at 1100 rpm in a Sorvall Legend T (Thermo Fisher Scientific, Waltham, MA). 18S gene
expression was used to confirm consistent loading (all Ct values between 8-9) and was also used
to normalize gene expression changes. Data were analyzed with an Applied Biosystems 7900HT
Real Time PCR machine using the range of 2-5 cycles for 18S and 2-12 cycles for other genes to
set a background level in order to determine a threshold for the fluorescence emission of the
TaqMan probes. Fold expression levels were generated using ΔΔCt method with respect to WT
SAL control samples set to a fold change level of 1. Genes were normalized with blom
transformations and statistical analysis was performed with Statistical Analysis System (SAS)
9.1.3 (Cary, NC). Gene expression data were log-transformed, mean centered and normalized
using Cluster version 2.11 and visualized using Treeview version 1.60 [321].

XVIII. Generation and stimulation of bone marrow-derived dendritic cells (bmDC) in
vitro.
The isolation and generation of bmDC were performed as adopted from Lutz, et.al.,[322]
and Brandt, et.al.,[323]. In short, mice were euthanized, femurs and tibias were removed
aseptically, and bone marrow was flushed with sterile RPMI. Single cell suspensions were
obtained and connective tissue was removed with a cell strainer (BD Falcon, Franklin Lakes,
NJ). Cells were cultured at 2.5 x 10^5 cells/mL in 10% BCS RPMI supplemented with 20 ng/mL
GMCSF (Peprotech, Rocky Hill, NJ). After exchange of 50% of media with fresh media and

55

removal of supernatant cells on days 3, 6, and 8 after start of culture, bmDC were used on day 9
for treatments. Lipopolysaccharide (LPS) (salmonella typhimurium, Sigma),1 µg/mL, and single
stranded RNA (ssRNA) (Invivogen, San Diego, CA), 5 µg/mL,were used to stimulate bmDC for
24 hr in 2% FBS RPMI. Subsequently, cells were stained with LIVE/DEAD cell viability dye
(Invitrogen) according to manufacturer’s instructions and anti-mouse I-A/I-E (MHC II,
M5/114.15.2), CD11c (N418), CD11b (M1/70), CD80 (16-10A1), CD86 (GL-1), H2Kb/H2Db
(MHC I, 28-8-6) as described above.

XIX. bmDC co-culture with OT-1 cells.
bmDC, generated as described above, were seeded at 2 x 10^5 cells per well in a 96 well
round bottom plate and pulsed with OVA257-264 (SIINFEKL) (Anaspec, San Jose, CA) for 2 h
in 200 µL of RPMI media. Subsequently, bmDC were washed 3X with RPMI. Spleens from OT1 mice were aseptically removed and washed in RPMI. OT-1 cells were labeled with CellTrace
(Invitrogen) proliferation dye according to manufacturer’s instructions. OT-1 cells were added
into wells with SIINFEKL-pulsed and –unpulsed bmDC at a final concentration of 5 x 10^6
cells/mL and co-cultured for 4 days. After 4 days in culture, OT-1 cells were restimulated with
addition of SIINFEKL at 5 ug/mL for 5 h in the presence of Brefeldin A in 2% BCS RPMI.
Staining was done with LIVE/DEAD cell viability dye and antibodies to CD8, IFNγ as described
above.

XX.

Isolation and activation of naïve CD4+ T cells.

Naïve CD4+ T cells were isolated from splenocyte preparations using a naïve CD4+ T
cell isolation kit according to manufacturer’s instructions (Miltenyi, Auburn, CA). To

56

immobilize αCD3 (145-2C11, BD Biosciences, San Jose, CA), flat bottom 96-well plates were
coated with 80 µL of αCD3 at 5 µg/mL overnight and washed 3 times with sterile PBS before
addition of cells. After purification, 0.5 x 10^6 naïve CD4+ T cells were added per well in a total
volume of 200 µL of RPMI supplemented with 5 % BCS and 1 % penicillin/streptomycin
(Sigma). To elicit Th0, αCD28 (37.51, BD Biosciences) was added at a final concentration of 1
ug/mL. For T helper 17 (Th17)-polarizing conditions, additional supplementation with TGF-beta
(5 ng/mL) (R&D Systems, Minneapolis, MN), IL-6 (20 ng/mL) (Jena Bioscience,Jena,
Germany), and IL-2 (40 U/mL) (RocheApplied Science, Indianapolis, IN) as well as αIFNγ (10
µg/mL) (BD Pharmingen, San Diego, CA) and αIL-4 (10 µg/mL) (BD Pharmingen) was
necessary [324]. Four days after elicitation, cell cultures were restimulated with PMA/Io and
stained as described above.

XXI. Δ9-THC treatment in vivo
The National Institute on Drug Abuse (NIDA, Bethesda, MD) provided Δ9-THC for
experimental use. For in vivo experiments, mice received corn oil (CO) or Δ9-THC by oral
gavage at 0.1 ml/g body weight for five consecutive days (-2 to 2 dpi) surrounding the
instillation of A/PR/8/34. Δ9-THC was dissolved in corn oil (CO) and administered at a final
dose of 75 mg/kg mouse/day.

XXII. Quantitative Real-time PCR for type I IFN, CB1, CB2, and H1 mRNA
cDNA was obtained as described above and was analyzed for IFNα2 and IFNβ1
expression by specific Taqman primer probe sets (Applied Biosystems) according to
manufacturer’s instructions. Each target gene (IFNa, IFNβ, CB1, CB2, and H1) was multiplexed

57

with an 18S loading control. Baseline levels for the loading control and the target genes were set
as described above and fold change levels were calculated as previously described. Data were
analyzed and graphed using GraphPad Prism (La Jolla, CA) version 4.03.

XXIII. Statistical analyses
Statistical analysis was performed with SAS and GraphPad Prism 4.03 (La Jolla, CA)
software. To determine statistical significance of the effect of treatment (A/PR/8/34 or SAL)
with respect to time, One Way ANOVAs with comparison to WT 1 dpi SAL were performed.
Factorial ANOVA with a Tukey’s post-hoc analysis was chosen for the statistical test with a 2 by
2 design including interactions for comparison between each genotype at each dpi.

The

independent variables were genotype (WT or CB1-/-CB2-/-) and treatment (A/PR/8/34 or SAL)
and the dependent variable was the obtained experimental value. For studies using Δ9-THC and
CO, a 2x2x2 (genotype, immune stimulation, drug treatment) design was used. A p-value of 0.05
or less was deemed statistically significant. In scenarios where data were non-parametric, such as
data represented as percentages, Friedman’s test was used as a substitute for ANOVA. To
determine a statistical significance of a Δ9-THC treatment effect, T-tests or non-parametric
Mann-Whitney U tests were performed.

58

EXPERIMENTAL RESULTS

I.

Δ9-THC Suppresses CTL Responses Independent of CB1 and CB2	
  

A) Kinetics of CTL Response
In order to determine the kinetics of the P815-induced CTL response, CTL were elicited
for 3, 5, 7, and 9 days after co-culture with irradiated P815 cells. Splenocytes from WT and
CB1-/-CB2-/- mice were used as the source of naïve CD8+ T cells. Elicited CTL were assayed
for CTL activity by 51Cr release assay using 51Cr-labelled P815 target cells. A ratio of 50:1
CTL:P815 cells was used to maximize release, thereby increasing sensitivity on non-peak days.
51Cr release is a result of CTL-dependent lysis of P815 cells and the peak day of CTL activity
occurred at day 5 post elicitation (Figure 4). At the peak day of CTL elicitation, CB1-/-CB2-/splenocytes had a statistically significant reduction in CTL activity, when compared to WT (p ≤
0.01). The same trend was observed for non-peak days, although it was not deemed statistically
significant. Subsequent experiments determining CTL effector activity under various conditions
were performed at day 5 post elicitation.

B) Δ9-THC Suppressed CTL Responses during Elicitation but not during the Effector
Response
To determine the sensitivity of CTL function to Δ9-THC treatment in the P815 allogeneic
model, Δ9-THC and/or vehicle (0.1% ethanol) were added directly to culture during either the
elicitation or effector phase of the CTL response. A concentration of 10 µM Δ9-THC was used
to determine sensitivity of CTL to immune modulation, a concentration that historically impaired
T cell immune responses in the absence of direct cytotoxicity [279]. Δ9-THC suppressed CTL

59

Figure 4. CTL Activity Peaks at Day 5 After Elicitation. To determine the peak day of
CTL activity, SPLC from WT and CB1-/-CB2-/- mice were co-cultured with irradiated P815
cells to elicit an allogeneic CTL response. At varying days after elicitation, CTL were
harvested and assayed for target (P815) lysing activity by reincubation at a ratio of 50:1
(Effector CTL : P815 Targets) with 51Cr labeled P815 cells. Percent release was calculated as
described in Methods (n=4). The experiment was performed once, while more replicate
experiments were performed for WT only. ## p ≤ 0.01 indicates difference between WT and
CB1-/-CB2-/-.

60

Figure 5. Δ9-THC Suppresses CTL Activity During Elicitation But Not Effector Phase.
SPLC were treated with Δ9-THC and/or VH (0.1% ethanol) and stimulated with irradiated
P815 cells for 5 days. After harvest, CTL were reincubated at a 10:1 ratio with 51Cr labeled
P815 cells in the presence of Δ9-THC and appropriate controls (NA – naïve: no treatment;
VH – 0.1 % ethanol) (n=4). Data shown are representative of two repeat experiments. * p ≤
0.05 indicates difference as compared to VH.

61

activity only when added prior to, but not after, the elicitation phase (Figure 5). Likewise,
repeated addition of Δ9-THC during elicitation and effector phases did not further reduce CTL
activity below single addition of the drug during the elicitation phase (Figure 5).

C) Concentration-dependent Suppression of CTL Effector Function
Next, we assessed whether CTL responses are suppressed by Δ9-THC in a concentrationdependent manner. At E:T ratios ranging from 10:1 to 1:1, Δ9-THC suppressed CTL responses
in a concentration-dependent manner with statistically significant suppression at concentrations
as low as 1 µM (Figure 6 A).

Also, lower E:T ratios resulted in lower CTL activity,

demonstrating that the elicited cell populations are responsible for the cytolytic activity and that
Δ9-THC suppressed CTL activity independent of the E:T ratio (Figure 6 A). Regression analysis
was performed on CTL activity results and a statistically significant difference was observed
from zero regression values for E:T ratios 10:1 (r2=0.8650, p<0.0001), 5:1 (r2=0.7246,
p=0.0004), 2.5:1 (r2=0.7461, p=0.0003), 1:1 (r2=0.7692, p=0.0002). Next, we assessed whether
the effect by Δ9-THC was dependent on the presence of CB1 and/or CB2. Δ9-THC suppressed
CTL activity in a concentration-dependent manner in CTL elicited from CB1-/-CB2-/splenocytes with significantly reduced CTL activity at concentrations of 5 µM or more as in WT
(Figure 6 B).

Regression analysis revealed a statistically significant difference from zero

regression of E:T ratio of 10:1 in WT (r2=0.6893, p=0.0008) and showed the same trend in CB1/-CB2-/- albeit not significant (r2=0.2905, p=0.0706) . Consistent with previous observations,
CTL activity of CB1-/-CB2-/- splenocytes was lower in magnitude compared to WT, although
differences in genotypes were not deemed statistically significant.

62

Figure 6. Δ9-THC Suppresses CTL Activity in a Concentration-Dependent
Manner. SPLC from WT were treated with 1, 5 and 10 µM of Δ9-THC and/or VH
(control) and co-cultured for 5 days with irradiated P815 cells. After harvest, CTL
were reincubated with 51Cr labeled P815 cells at indicated ratios of 10:1 to 1:1 (A)
(n=4). In a second experiment SPLC from WT and CB1-/-CB2-/- were elicited and
restimulated with 51Cr labeled P815 cells at a ratio of 10:1 (B) (n=4). Data shown
are representative of two repeat experiments. * p ≤ 0.05, ** p ≤ 0.01 indicate
differences as compared to VH.

63

Figure 6. Δ9-THC Suppresses CTL Activity in a Concentration-Dependent Manner.

64

D) Δ9-THC-Mediated Suppression of CTL Generation
Although MHC I mismatch is a strong stimulus for driving the expansion of
differentiated CD8+ T cell effectors, the overall number of viable T cells within control cultures
(NA or VH treatment groups) on day 5 of co-culture with P815 allogenic target cells was
modest. This demonstrated that T cell proliferation and clonal expansion in response to MHC Irestricted alloantigens in this assay system induced a relatively small number of T cells from the
spleen. To assess the effects of Δ9-THC on its ability to impair alloantigen-induced expansion
of T cell effectors, the number of viable CD4+ and CD8+ T cells was enumerated on Day 5 of
the elicitation phase of the CTL response. Viability was determined using LIVE/DEAD dye
(Invitrogen), which permeates dead cells and thus low staining is observed in viable cell
populations. The percentage of viable CD4+ (Figure 7 A, B) and CD8+ (Figure 7 C, D) was
markedly reduced by Δ9-THC treatment 5 days after elicitation with concentrations as low as 1
µM in both WT and CB1-/-CB2-/- samples. No significant differences were found in the percent
of viable CD8+ cells between CB1-/-CB2-/- and WT splenocytes (Figure 7 C, D); however, a
greater number of viable CD4+ T cells was observed in CB1-/-CB2-/- compared to WT
splenocytes (Figure 7 A, B).
IFNγ production by CD8+ T cells is often used as a surrogate for CTL activity. After
gating for single live cells (Figure 8 A), CD4+ cells did not produce significant amounts of IFNγ
as P815 cells are MHC I-mismatched to C57Bl/6 cells and do not express significant amounts of
MHC II (Figure 8 B). However, significant IFNγ production was detected by CD8+ cells, which
was enhanced in CTL elicited from WT compared to CB1-/-CB2-/- (Figure 8 C-E).

In the

presence of Δ9-THC, enhanced IFNγ production was observed in CD8+ T cells derived from

65

Figure 7. Lower Viability of CD4+ and CD8+ Cells Following Δ9-THC Treatment.
CTL were elicited from SPLC of WT and CB1-/-CB2-/- mice in the presence or absence of
VH (0.1 % ethanol) or Δ9-THC (1, 5, and 10 µM). After 5 days in culture, CD4 and CD8
surface staining was performed and viability was assessed using LIVE/DEAD staining.
Viability was determined within CD4+ (A, B) and CD8+ (C, D) cells after singlet and
lymphocyte size gating (n = 4). Data shown are representative of four repeat experiments. * p
≤ 0.05, ** p ≤ 0.01 indicate differences as compared to VH, ## p ≤ 0.01 as compared to WT.

66

Figure 7. Lower Viability of CD4+ and CD8+ Cells Following Δ9-THC Treatment.
67

Figure 8. Increased IFNγ Production in Live But Not Total Cells After Δ9-THC
treatment.
CTL of WT and CB1-/-CB2-/- were elicited as before with and without VH (0.1%
Ethanol) or Δ9-THC (1, 5, 10 µM) treatment for 5 days. Cells were restimulated for 12 h
in the presence of Brefeldin A and stained for CD4+, CD8+, LIVE/DEAD and IFNγ.
Gating scheme is shown (A) for populations of CD4+ (B) and CD8+ (C) cells and dot
plots indicate concatenated samples (n=4). Bar graph for CD8+ cells are shown within live
populations % (D), MFI (E), and in % of total populations (F) (n=4). Data shown are
representative of three repeat experiments. ** p ≤ 0.01 indicate differences as compared
to VH, ## p ≤ 0.01 as compared to WT.

68

Figure 8. Increased IFNγ Production in Live But Not Total Cells After Δ9-THC treatment.

69

Figure 8 cont’d.

70

/-CB2-/- mice, although not deemed statistically significant (Figure 8 C-E).
Also, IFNγ production was reduced in samples from CB1-/-CB2-/- compared to WT mice
as previously observed in 51Cr release assays (Figure 8 C-E). Analysis of CD8+ IFNγ-secreting
cells, as a percentage of viable cells, takes into account the reduced viability as a result of Δ9THC addition (see Figure 7). However, while there was an enhancement of IFNγ-secreting cells
with Δ9-THC treatment, there were also fewer viable cells after Δ9-THC treatment, thus overall
the percentage of CD8+ cells secreting IFNγ within the total population did not change (Figure 8
F).

E) Δ9-THC Does Not Affect CD8+ T cell Proliferation Induced by P815 Co-culture
Prior to becoming functional CTL, proliferation occurs in T cells to expand the effector
pool. Thus, we focused on proliferation as a potential endpoint leading to decreased viability and
decreased CTL response.

Whole splenocytes were labeled with CFSE dye to determine

proliferation by dye dilution.

No statistically significant differences in proliferation were

observed between VH and Δ9-THC-treated groups in CD8+ or in CD4+ cells from WT or CB1/-CB2-/- mice (Figure 9 A-D). In comparison to WT, CD4+ cells obtained from CB1-/-CB2-/spleens underwent much greater proliferation. CD4+ T cells were not directly stimulated in this
model, suggesting that greater proliferation in CB1-/-CB2-/- CD4+ cells might either be due to
higher basal proliferation in CB1-/-CB2-/- cells or a result of the cytokines produced by CD8+
cells. In contrast, notably more CD8+ cells were elicited in samples from WT compared to CB1/-CB2-/- spleens (Figure 9 C, D).

71

Figure 9. No Effect of Δ9-THC on CTL Proliferation.
SPLC were isolated from WT and CB1-/-CB2-/- mice and labeled with Cell Trace CFSE
according to manufacturer’s instructions. Four days after co-culture with irradiated P815 in
the presence or absence of VH (0.1 % ethanol) and Δ9-THC (1, 5, 10 µM), surface staining
with CD4 and CD8 were performed and CFSE fluorescence was assessed by FACS. Shown
are concatenated samples (n=4) of CFSE staining as a result of proliferation in CD4+ (A, B)
and CD8+ cells (C, D). Data shown are representative of two repeat experiments. * p ≤ 0.05
indicate differences as compared to VH, ## p ≤ 0.01 as compared to WT.

72

Figure 9. No Effect of Δ9-THC on CTL Proliferation.
73

F) Increase of CD69 Expression on CD8+ Cells with Δ9-THC Treatment
In an effort to determine events leading to the Δ9-THC-mediated reduction in cell
viability and CTL activity seen at day 5, without any apparent effects on proliferation, we
focused on T cell activation. Two markers of cellular activation found on both CD4+ and CD8+
T cells are CD69 and CD25. Whereas CD69 is thought to be one of the earliest markers of
activation and is rapidly upregulated, CD25 expression increases gradually on activated T cells
and is observed later than CD69 on activated T cells [325, 326].

We focused on CD69

expression at 6 h and CD25 expression at 12 h after co-culture with irradiated P815 cells. Cells
were gated on singlets, by size, and then on CD4 or CD8 (Figure 10 A). In CD4+ cells, CD69
expression was unaffected by co-culture with P815 cells; however, cells from CB1-/-CB2-/mice showed higher basal expression compared to WT mice (Figure 10 B, C). In addition, no
effect with Δ9-THC treatment was observed on CD69 expression on CD4+ cells (Figure 10 B,
C). In contrast, the expression of CD69 on CD8+ was increased by co-culture with P815 cells
and CD8+ T cells from CB1-/-CB2-/- expressed higher levels of CD69 than WT CD8+ T cells
(Figure 10 D, E). CD69 expression was increased in a concentration-dependent manner by Δ9THC in CD8+ T cells from both WT and CB1-/-CB2-/- spleens, which was significant at
concentrations as low as 5 µM in CB1-/-CB2-/- -derived cells (Figure 10 D, E).
CD25 expression on CD4+ T cells was increased as a result of stimulation with P815
cells and increased levels of CD25 expression were observed in naïve unstimulated samples from
CB1-/-CB2-/- compared to WT mice (Figure 11 A, B). There was a trend towards a Δ9-THCdependent increase in CD25 expression in both WT and CB1-/-CB2-/- CD4+ cell populations,
but this trend was not statistically significant (Figure 11 B). In CD8+ cells, the expression of
CD25

74

Figure 10. Δ9-THC Increases CD69 Expression on CD8+ cells in a ConcentrationDependent Manner
SPLC from of WT and CB1-/-CB2-/- mice were incubated with irradiated P815 cells for 6 h
to induce CD69 surface expression, in the presence or absence of VH (0.1 % ethanol) or Δ9THC (1,5, and 10 µM). Cells were gated on singlets and lymphocyte populations by size and
then on CD4 and CD8 (A). CD4+ (B, C) and CD8+ (D, E) cells positive as defined by the box
gate for CD69 in % are shown (n=4). Data shown are representative of three repeat
experiments. Difference due to stimulation by P815 is indicated by ++ p ≤ 0.01, between
genotypes is indicated by ##, p ≤ 0.01 due to Δ9-THC by *, p ≤ 0.05** p ≤ 0.01.

75

Figure 10. Δ9-THC Increases CD69 Expression on CD8+ cells in a Concentration-Dependent Manner
76

Figure 10. cont’d.

77

Figure 11. Δ9-THC Decreases CD25 Expression on CD8+ Cells from CB1-/-CB2-/- mice
in a Concentration-Dependent Manner
SPLC from of WT and CB1-/-CB2-/- mice were co-cultured with irradiated P815 cells for 12
h to induce CD25 levels, in the presence or absence of VH (0.1 % ethanol) or Δ9-THC (1,5,
and 10 µM). Cells were gated on singlets and lymphocyte populations by size and cells
positive for CD25 in % are shown (n=4). Data shown are representative of two repeat
experiments. Difference due to stimulation by P815 is indicated by ++ p ≤ 0.01, between
genotypes is indicated by ##, p ≤ 0.01 due to Δ9-THC by *, p ≤ 0.05.

78

Figure 11. Δ9-THC Decreases CD25 Expression on CD8+ Cells from CB1-/-CB2-/- mice in a Concentration-Dependent
Manner
79

Figure 11 cont’d

80

was much lower than in CD4+ cells (Figure 11 A-D). Expression of CD25 on CD8+ cells of
both genotypes was increased by co-culture with P815 cells (Figure 11 C, D). In CD8+ cells
from CB1-/-CB2-/- mice, a concentration-dependent decrease with Δ9-THC treatment was
observed, while in WT mice Δ9-THC had no effect (Figure 11 C, D).

G) Δ9-THC Synergizes with Io to Induce CD69 Expression
In light of the rapid Δ9-THC-mediated increase in CD69 expression of CD8+ T cells and
its inverse correlation with Δ9-THC-mediated suppression of CTL activity, we focused on
intracellular signals leading to CD69 protein expression. Previously, it was determined that
independent of CB1 and CB2, Δ9-THC partially mediated its effects through the TRPC1
channel, which causes an increase in intracellular Ca2+ levels [284].

Using the calcium

ionophore, Io, we tested whether a rise in intracellular Ca2+ might contribute to the observed
increase in CD69 expression. In addition, the effect of Δ9-THC on Io-induced CD69 expression
was evaluated. Again, cells were gated as described above (Figure 10 A). In CD4+ cells, Δ9THC increased expression of CD69 moderately in the absence of an activating stimulus when
isolated from CB1-/-CB2-/-, but not WT mice (Figure 12 A, C). Δ9-THC increased CD69
expression in CD8+ splenocytes from both WT and CB1-/-CB2-/- mice in the absence of an
activating stimulus (Figure 12 A, E).

After stimulation with Io, CD69 expression was

dramatically increased in CD4+ and CD8+ cells from both genotypes (Figure 12 B, D, F). While
CD69 expression was similar between WT and CB1-/-CB2-/- CD8+ cells (Figure 12 B, F), a
marked reduction in CD69 expression was observed in CB1-/-CB2-/- CD4+ cells, when
compared to WT CD4+ cells (Figure 12 B, D). Most importantly, Δ9-THC synergized with Io to

81

Figure 12. Δ9-THC synergizes with Io to upregulate CD69 expression.
SPLC were incubated in the presence of Io (0.5 µM) or vehicle (0.01 % DMSO) in the
presence or absence of VH (0.1 % ethanol) or Δ9-THC (1,5, and 10 µM). For NA-0 h staining
was performed after isolation of a single cell suspension from the spleen otherwise 6 h after
co-culture, CD69 staining was performed. Cells were gated on singlet, lymphocytes and
within CD4 (A-D) or CD8 (A, B, E, F) positive populations. Data shown in histograms are
concatenated (n=4) and are representative of two repeat experiments. Difference due to
stimulation by Io is indicated by ++ p ≤ 0.01, between genotypes is indicated by ##, p ≤ 0.01
due to Δ9-THC by **, p ≤ 0.01.

82

Figure 12. Δ9-THC synergizes with Io to upregulate CD69 expression.

83

Figure 12 cont’d

84

further increase CD69 surface expression in CD4+ and CD8+ cells derived from both genotypes
(Figure 12 B, D, F).

II. In Vivo Influenza PR8 Immune Response Kinetics in WT and CB1-/-CB2-/- mice

A) Enhanced leukocyte infiltration and injury in airways of CB1-/-CB2-/- mice after
influenza infection
Principal histopathology in the lungs of WT mice instilled with influenza virus was a
necrotizing bronchiolitis of large-diameter axial and pre-terminal bronchioles (Figure 13 A).
These airways had conspicuous necrosis and exfoliation of surface epithelium associated with a
mixed inflammatory cell infiltration. Inflammatory cell influx along with edema was also present
in peribronchiolar and perivascular tissues (Figure 13 A). CB1-/-CB2-/- mice instilled with
influenza virus developed a similar, but more severe, necrotizing bronchiolitis. No lung lesions
were present in WT and CB1-/-CB2-/- SAL-control mice (Figure 13 A). Not only were the
epithelial and inflammatory responses in large-diameter bronchioles more marked, the virusinduced epithelial necrosis, exfoliation and accompanying inflammation extended to more distal,
small-diameter, terminal bronchioles with some extension into adjacent alveolar ducts (Figure 13
A). This difference in severity and extent of viral bronchiolitis between CB1-/-CB2-/- and WT
mice was also immunohistochemically demonstrated by a loss of anti-inflammatory CCSP in
bronchiolar epithelium lining the terminal bronchioles of virus-treated CB1-/-CB2-/- mice
(Figure 13 B). Influenza infection induced airway neutrophil and monocyte infiltration most
strongly at 3 dpi as enumerated in the BALF (Figure 14). The magnitude of the cell infiltrate
was enhanced in CB1-/-CB2-/- compared to WT mice 3 dpi, the peak day of inflammatory cell

85

Figure 13. More Severe Virus-Induced Necrotizing Bronchiolitis and Loss of Clara Cell
Secretory Protein in CB1-/-CB2-/- compared to WT Mice 3 Days Post-infection.
Light photomicrographs of axial airways (AA; generation 5) and terminal bronchioles (TB;
generation 5) from the left lung lobe of wild type and CB1-/-CB2-/- mice intranasally instilled
with saline vehicle alone or PR8 influenza virus in saline at 3 dpi. The insets are low
magnification of tissue sections of the left lung lobe taken perpendicular to the AA at generation
5 (see methods section for microdissection details). All tissues were stained with hematoxylin
and eosin (A) or immunohistochemically stained for Clara Cell Secretory Protein (red
chromagen) located in bronchiolar epithelium (arrows) at 3 dpi (B). Virus-induced necrotizing
bronchiolitis is evident in the AA of WT and CB1-/-CB2-/- mice with greater necrosis and
exfoliation (arrows) of the surface epithelium (e) and interstitial cellular inflammation and edema
(asterisks) in the CB1-/-CB2-/- mouse compared to the WT mouse. Similar necrotizing
bronchiolitis is evident in the terminal bronchiole (TB) of CB1-/-CB2-/- mouse, but not in the
virus-treated WT mouse. Compared to saline-instilled control mice there was an apparent loss of
CCSP staining in the bronchiolar epithelium (e) lining the large-diameter AA and smaller
diameter preterminal and terminal bronchioles of virus-instilled CB1-/-CB2-/- mice. In virustreated WT mice, there was a loss of CCSP only in large-diameter AA, but CCSP was still
present in preterminal and terminal bronchioles.

86

Figure 13 Continued. More Severe Virus-Induced Necrotizing Bronchiolitis and
Loss of Clara Cell Secretory Protein in CB1-/-CB2-/- compared to WT Mice 3 Days
Post-infection (continued). All CCSP stained tissue sections were counter-stained with
hematoxylin. AD, alveolar duct; ap, alveolar parenchyma; p, pleura; a, alveolus; bv,
blood vessel; asterisk, necrotic cellular debris and inflammatory cells in airway
lumen.Light microscopic examinations of histologic tissue sections were conducted
using an Olympus BX41 clinical microscope with UPlanApochromat 10x, 20x, and 40x
and Achromat 60x brightfield objectives (www.olympusamerica.com). A 5.0 Megapixel
Qlympus Q-Color5 camera system was used for all light photomicroscopy. This
included a High Resolution FireWire digital CCD color camera, designed for
publication and documentation, with a 2/3” Bayer color sensor and 2580 x 1944 pixel
resolution. The camera was connected to a Dell Precision 380 computer and QCapture
Suite software for Windows allowing for real-time viewing and full computer control.
For interpretation of the references to color in this and all other figures, the reader is
referred to the electronic version of this dissertation.
of the camera.

87

WT

ß

CB1-/-CB2-/-

Figure 13. More Severe Virus-Induced Necrotizing Bronchiolitis and Loss of Clara Cell
Secretory Protein in CB1-/-CB2-/- compared to WT Mice 3 Days Post-infection
88

Figure 13 cont’d.

WT

CB1-/-CB2-/-

89

Figure 13 cont’d.

90

Figure 14. Greater Neutrophils and Macrophage Recruitment after Influenza Infection
in CB1-/-CB2-/- mice.
Lungs of C57Bl/6 and CB1-/-CB2-/- mice (n=5) were lavaged with SAL to collect BALF.
Total cell number was determined by counting of cells in BALF and DiffQuick staining was
performed to distinguish between immune cell populations, which were enumerated by based
on the percentage found on the slide using light microscopy. * (p ≤ 0.05) ** (p ≤ 0.01) marks
a significant change induced by influenza within a timepoint, while # (p ≤ 0.05) ## (p ≤ 0.01)
marks a significant difference between genotypes.

90

influx into the airways. At 7dpi, the immune cell infiltrates in the airways were lower in CB1-/CB2-/- compared to WT mice.

B) Increased mRNA expression levels of pro-inflammatory immune mediators in
lungs from CB1-/-CB2-/-mice
Based on the enhanced immune cell infiltration and increased tissue injury in influenzainfected lungs of CB1-/-CB2-/- mice 3 dpi, gene expression changes were assessed using a PCRbased immune panel with total lung RNA from 1 and 3 dpi (Figure 15). Differential gene
expression was observed in the lungs of uninfected (SAL-instilled) CB1-/-CB2-/- compared to
WT mice (Figure 15 A). Specifically, increased expression of transcripts for MHCII subunit H2Eb1, cytokines Il1a, IL1b, chemokine Ccl3, chemokine receptor Ccr4, and enzyme prostaglandin
synthase 2 Ptgs2 were observed. In addition, decreases in chemokine Ccl19 and transferrin
receptor 1 Tfrc were evident (Figure 15 A).
In influenza-infected mice, most influenza-associated gene expression changes were not
evident prior to 3 dpi (data not shown). For the purpose of this study, the focus was placed on
genes that were differentially regulated between CB1-/-CB2-/- compared with WT mice.
Exacerbated expression of cytokines, Csf2 (GMCSF), Il13, Il17, and Tnf, negative regulator of
cytokines, Socs2, chemokine receptor Ccr7, surface marker Cd38, and the previously mentioned
H2-Eb1 and Ptgs2 was observed (Figure 15 B). Collectively, gene transcript levels indicating
increased inflammation and maturation of antigen presenting cells and the presence of cellular
responses involving cytokine secretion in CB1-/-CB2-/- mice defined the overall profile.

91

Figure 15. Relative transcript levels of genes associated with inflammation are
increased in lungs of CB1-/-CB2-/- mice compared to WT mice basally and after
influenza infection at 3 dpi.
RNA from lung samples (n=4) was converted to cDNA and analyzed using an Applied
Biosystem’s Mouse Immune Panel. Genes were included based on 1.5 fold up- or
downregulation and fold changes in gene expression levels were analyzed using
factorial ANOVAs for significance. Genes with significantly different expression
between CB1-/-CB2-/-

and WT mice were visualized in a heat map with red

indicating an increase and blue a decrease in expression in a log2 relative expression
scale as shown on the right side of the figure. The p-value of the factorial ANOVA is
shown on the left side of each gene. Arrangement of the heat map is as follows: A –
differentially regulated genes basally, without influenza infection between CB1-/CB2-/- and WT mice, B – gene expression differentially expressed in CB1-/-CB2-/mice after influenza infection. For interpretation of the references to color in this and
all other figures, the reader is referred to the electronic version of this dissertation.

92

Figure 15. Relative transcript levels of genes associated with inflammation are increased in lungs of CB1-/-CB2-/mice compared to WT mice basally and after influenza infection at 3 dpi.

93

Figure 16. Peak H1 mRNA Levels 3 dpi in CB1-/-CB2-/- and WT Mice and
Increases in Steady-state CB1 Transcript Levels 7 dpi
RNA was isolated from whole lungs (n=5) and assessed for steady state levels of H1
RNA (A), CB1 (B) and CB2 (C) mRNA using 18S as a housekeeping gene. All fold
changes are relative to WT SAL 1 dpi. Factorial ANOVA was used to determine
significance for H1 RNA levels and One-Way ANOVA was used for CB1 and CB2
levels. ** p = 0.01 compared to WT SAL.

94

Figure 16. Peak H1 mRNA Levels 3 dpi in CB1-/-CB2-/- and WT Mice and Increases in Steady-state CB1
Transcript Levels 7 dpi
95

C) No difference in the H1 RNA Levels between WT and CB1-/-CB2-/- mice, But
Increase in CB1 Transcript in WT Mice after Influenza Infection.
RNA was isolated and converted to cDNA as described before and H1 levels were assessed
in lung RNA isolated from WT and CB1-/-CB2-/- mice CB1 and CB2 were assessed in WT
samples only. The greatest levels of H1 RNA were observed at 3 dpi in both, WT and CB1-/CB2-/- mice. In contrast to previous observations in this laboratory [120], there was no
significant difference between the H1 RNA levels between WT and CB1-/-CB2-/- mice (Figure
16 A). While CB2 steady-state mRNA levels remained unchanged after PR8 infection (Figure 16
C), CB1 steady-state mRNA levels increased, reaching significance at 7 dpi compared to SAL
(Figure 16B).

D) T cells from CB1-/-CB2-/- mice undergo accelerated T cell activation
To investigate the effect of gene expression changes on the T cell response in CB1-/CB2-/- mice after influenza infection, CD69 expression was monitored on 1-7 dpi as a marker of
T cell activation in whole lung preparations. In contrast to BALF, in which myeloid cells
predominated (shown in Figure 14), whole lung preparations harbored cells of the lymphoid
lineage. Induction of surface CD69 was observed 3 dpi in both CD4+ and CD8+ T cells, with
greater staining intensity observed in CD4+ T cells (Figure 17). CD69 expression was greater in
CD4+ T cells from CB1-/-CB2-/- compared to WT mice as early as 3 dpi (Figure 17 A). The
same trend was observed in CD8+ T cells, yet to a lesser degree than in CD4+ T cells (Figure 17
B). Increased CD69 expression was observed in SAL-instilled CB1-/-CB2-/- compared to WT
mice. At 7 dpi, WT and CB1-/-CB2-/- mice displayed similar levels of surface CD69 in both

96

Figure 17. More rapid T cells activation after Influenza infection in CB1-/-CB2-/compared to WT mice.
At 1, 3, 5, and 7 dpi cells were isolated from lung tissue (n=5), Fc receptors were blocked on
the surface and samples stained for surface markers CD4, CD8 and CD69. FlowJo software
was used to analyze samples and create histograms shown. Gating was performed on single
cell populations, size and expression of CD4 and CD8 (bottom left). Within CD4+ (A) and
CD8+ (B) populations histograms depict the fluorescence intensity of CD69 staining. Due to
bimodal expression of CD69, percent positively gated cells, as show in the first panel (top
right, A), were enumerated (CD4: C, CD8: D). Friedman’s tests for non-Gaussian data were
performed and significance was indicated in figures as: # p = 0.05 and ## p = 0.01 in
comparison of WT to CB1-/-CB2-/-.

97

Figure 17. More rapid T cells activation after Influenza infection in CB1-/-CB2-/- compared to WT mice.
98

Figure 17 cont’d.

99

CD4+ and CD8+ T cells with CD69 expression starting to wane in cells from CB1-/-CB2-/- mice
(Figure 17 A, B).

D) Enhanced IL-17 and IFNγ production in NK, and T cells isolated from lungs of
CB1-/-CB2-/- mice in response to influenza
The observation of elevated steady state mRNA levels of IFNγ and IL-17, and more rapid T
cell activation as assessed by CD69 in CB1-/-CB2-/-, when compared to WT mice, suggested a
regulatory function by cannabinoid receptors in antiviral T cell immunity. In light of the
increased transcriptional expression of the T cell-produced cytokines, IFNγ and IL-17, in the
lung (Figure 15) and their pivotal roles in lung inflammation, identification of the cellular
source(s) of IFNγ and IL-17 was important in determining the cell types affected by CB1 and
CB2 deletion.
The gating scheme for the analysis of cytokine secretion from whole lung homogenates is
shown in Figure 18 A. Minor basal IFNγ production, mainly from CD4+ and CD8+ cells, was
observed in SAL control samples in WT (Figure 18 B). In CB1-/-CB2-/- mice, percentages of
IFNγ producing cells in the absence of influenza challenge were markedly enhanced compared
to WT mice, especially in CD4+ cells (Figure 18 B). The percentage of IFNγ-producting cells
was modestly decreased 3 and 5 dpi in WT but not CB1-/-CB2-/- CD4+ cells, while increases in
IFNγ-producing cells were observed in CD8+ and NK1.1+ populations (Figure 18 B).Overall,
the percentage of IFNγ producing cells was greater in all three (CD4+, CD8+, NK1.1+)
delineated effector cell types isolated from CB1-/-CB2-/-mice compared to WT mice.
On the other hand, basal IL-17 production was limited predominantly to NK1.1+ cells in
WT mice (Figure 18 B). In contrast, both CD4+ and NK1.1+ cells from uninfected CB1-/-CB2-/-

100

Figure 18. Greater cytokine production by leukocytes from CB1-/-CB2-/- mice
compared to WT mice.
Lungs from mice were mechanically disrupted and passed through a sieve (n=5). After
removal of connective tissue, cells were counted and restimulated with PMA/Io for 5 h
in the presence of Brefeldin A. Cells were blocked with Fc receptors and stained with
CD4, CD8, and NK1.1 in addition to cytokine staining with IFNγ and IL-17. Flow
cytometry samples were gated as depicted in A. Cytokine secretors were identified as
percent of positive cells within surface delineation and unstimulated samples were
used as controls (B). Immune cell populations were enumerated for percent cytokine
expression in FlowJo (type of cytokine indicated on the left) within effector cell
populations (surface marker indicated on the right) in FlowJo and bar graphs were
generated using GraphPad Prizm (C). Friedman’s tests for non-Gaussian data were
performed and significance is indicated in figures as: # p = 0.05 and ## p = 0.01 in
comparison of WT mice to CB1-/-CB2-/- mice.

101

Figure 18. Greater cytokine production by leukocytes from CB1-/-CB2-/- mice
compared to WT mice.

105

Figure 18 cont’d.

106

mice produced significant amounts of IL-17 (Figure 18 B). At 3 dpi, the IL-17 producing cell
types did not change, but the percentage of detectable IL-17-producing cells increased, especially
in CD4+, and to a lesser extent in CD8+ cells, with greater percentages of IL-17 positive cells
detected in CB1-/-CB2-/- mice compared to WT mice (Figure 18 B). By 5 dpi, the percentage of
cells staining positive for IL-17 was greatly reduced in NK and CD4+ cells in WT and CB1-/CB2-/- mice and in CD8+ cells of CB1-/-CB2-/- but not WT mice. In summary, IL-17 and IFNγ
hypercytokinemia in effector cells isolated from the lungs was evident in the absence of
influenza infection and even more pronounced after infection in CB1-/-CB2-/- compared to WT
mice.

E) Fewer Naïve CD4+ T cells in CB1-/-CB2-/- compared to WT mice
Since IFNγ- and IL-17-secreting CD4+ T cells were found in uninfected, SAL-instilled
lungs of CB1-/-CB2-/- mice, we focused on the percent composition of naïve T cells
(CD62L+CCR7+). After gating for CD4+ cells from splenocytes (Figure 19 A), CD62L-CCR7and a CD62L-CCR7+ non-NA CD4+ T cells populations were identified and found in greater
percentages in CB1-/-CB2-/- mice (Figure 19 B, C). To assess whether deletion of CB1 and/or
CB2 had a direct effect on the ability of T cells to secrete cytokines, naïve CD62L+ CD4+ T
cells were isolated from splenocytes and stimulated with immobilized αCD3 and soluble αCD28
under Th17 polarizing conditions to induce IL-17 secretion. Under these conditions, which favor
IL-17 production, IL-17 was detected from both WT and CB1-/-CB2-/- non-naïve T cells
(CD62L-) and naïve T cells (CD62L+) (Figure 19 D). No difference in the production of IL-17
was observed in CD4+ T cells from CB1-/-CB2-/- compared to WT (Figure 19 E-G). This

104

Figure 19. Lower number of NA CD4+ T cells and no difference in T cell stimulation in
vitro in CB1-/-CB2-/-mice compared to WT mice.
Splenocytes (n=3) were obtained from WT and CB1-/-CB2-/- mice and analyzed by flow
cytometry with gating shown in A. Non-NA T cells (CCR7-CD62L- or CD62L-), were gated
for in B and analyzed statistically by Friedman’s test in C. NA (CD62L+) (NA) and non-naïve
(non-NA) CD4+ cells were isolated from splenocytes and driven towards a Th17 phenotype
in the presence of TGF-β and IL-6 as described in methods. Cells were gated as shown in D
and then analyzed for cytokine production in E. Results for Friedman’s tests for non-Gaussian
data were of experimental data in Th17 polarizing conditions from E are shown in F and G.
The experiment is representative of two repeat in vitro studies. * p≤0.05, ** p≤0.01
comparing WT and CB1-/-CB2-/-.

105

Figure 19. Lower number of NA CD4+ T cells and no difference in T cell stimulation in vitro in CB1-/-CB2-/-mice
compared to WT mice.
106

Figure 20. Increased expression of DC maturation markers in bmDC from CB1-/-CB2-/mice.
bmDC were generated as described in the methods. In short, bone marrow was isolated from
femurs and tibias and cultured in the presence of GMCSF (20 ng/mL) for 9 days, then
immature bmDC were washed and stimulated with TLR agonists (n=3). A, bmDCgated for
singlets, LIVE (LIVE/DEAD dye), and high CD11b and CD11c expression. B, surface
expression of maturation markers MHC I, MHC II, CD80, CD86 were analyzed after
stimulation for 24 h with LPS (1 µg/mL) and ssRNA (5 µg/mL). C, due to the Gaussian
distribution of the fluorescent staining, mean fluorescence intensity (MFI) was used to
compare samples using statistics. The results shown are representative from four identical
experiments.

107

Figure 20. Increased expression of DC maturation markers in bmDC from CB1-/-CB2-/- mice.
108

suggests that hypercytokinemia experienced in CB1-/-CB2-/-mice in vivo might be a result of
interaction with APCs since T cells intrinsically did not produce different level of cytokines.

F) Increased APC maturation and ability to induce proliferation and cytokine
production in T cells in CB1-/-CB2-/-mice compared to WT mice
To determine whether APC responses are in fact deregulated in CB1-/-CB2-/-mice, as
suggested by a lower percentage of NA T cells, an in vitro model was used to investigate DC,
which link innate to adaptive immune responses based on their role in T cell elicitation.
Immature bone marrow-derived dendritic cells (bmDC) were generated in vitro from femurs and
tibias of CB1-/-CB2-/- and WT mice and stimulated with LPS (1 µg/mL) or ssRNA (5 µg/mL)
for 24 h to induce a mature phenotype. Flow cytometric staining for CD11b and CD11c was
performed to confirm conventional DC (cDC) phenotype (CD11b+CD11c+) [133] (Figure 20
A). Staining for MHCI, MHCII, CD80, CD86 was performed to evaluate the maturation status
(Figure 20 B). Increased staining for MHCI, MHCII and CD86 was found in bmDC of CB1-/CB2-/- compared to WT mice without TLR treatment (Figure 20 B). LPS upregulated surface
expression of MHCII and CD86, while ssRNA increased CD86 expression in both genotypes. In
addition, ssRNA treatment elevated MHCI in bmDC from CB1-/-CB2-/-mice only. In contrast,
CD80 expression did not change with TLR treatment (Figure 20 C). Overall, bmDC generated in
vitro from CB1-/-CB2-/- but not WT mice displayed a pre-activated phenotype. Upon TLR
activation, maturation markers were induced with greater magnitude in CB1-/-CB2-/-compared
to WT mice.
To determine whether the observed phenotypic changes on bmDC exerted an effect on T
cell elicitation and function, bmDC pulsed with SIINFEKL were cultured with OT-1 splenocytes

109

Figure 21. bmDC from CB1-/-CB2-/-elicit OT-1 responses without requirement for
maturation stimulus and more potently than WT.
OT-1 cells were elicited as described in the methods section. In short, bmDC were matured
with different TLR agonists for 24 h and pulsed with SIINFEKL peptide for 2 h, washed, and
incubated with OT-1 cells for 4 days, then restimulated for 5 h to induce cytokine secretion
(n=3). A, gating scheme for OT-1 cells; singlet, size, and LIVE gating was applied before
visualizing data. B, proliferation was assessed by Cell Trace dye dilution and a gate depicting
proliferated populations was drawn according to WT-NA population shown in tinted grey.
The legend for the other experimental groups is below the figure and the TLR agonist used is
shown beside proliferation graph. C, proliferation and concurrent IFNγ secretion were
visualized using dot plots. LPS (1µg/mL) and ssRNA (5µg/mL) were used to stimulate bmDC
maturation as previously (Figure 6). Proliferation and IFNγ secretion gates were drawn
according to a WT-NA group without SIINFEKL peptide, which showed no proliferation or
IFNγ secretion. D, data from proliferation and cytokine plots are summarized using
percentages obtained from shown gates. Statistical significance is indicated as # - p ≤ 0.05
comparing WT to CB1-/-CB2-/- using Friedman’s test for non-parametric data. The results
shown are representative from three identical experiments.

110

Figure 21. bmDC from CB1-/-CB2-/-elicit OT-1 responses without requirement for maturation stimulus and more potently
than WT.

111

Figure 22. AM isolated from BALF are mature in CB1-/-CB2-/- but not WT.
AM were isolated by flushing the lungs with saline, washed and plated in the presence or
absence of LPS (1 µg/mL) (n=3). Maturation was assessed by staining for MHC I, MHC II,
and CD86 on the surface of AM. A, AM were gates as CD11b+CD11c hi cells. B, histograms
depicting changes in fluorescence intensity of MHC I, MHC II, and CD86 in AM obtained
from lavage fluid of WT and CB1-/-CB2-/- mice. C, summary of MFI data, statistical
significance is indicated as ** p ≤ 0.01 WT-NA vs. WT-LPS, # p ≤ 0.05 and ## p ≤ 0.01 WT
vs CB1-/-CB2-/- within groups. The results shown are representative from two identical
experiments.

112

Figure 22. AM isolated from BALF are mature in CB1-/-CB2-/- but not WT.

113

and evaluated after gating on live cells (Figure 21 A). OT-1 splenocytes were also labeled with a
dye for assessment of proliferation by dye dilution and stained for IFNγ to assess function after
restimulation. No proliferation or IFNγ was detected in OT-1 splenocytes incubated with NA
bmDC from WT mice (Figure 21 B, C). However, OT-1 splenocytes added to bmDC from CB1/-CB2-/- mice were induced to proliferate and secreted IFNγ in the absence of a maturation
stimulus such as LPS or ssRNA (Figure 21 B, C). LPS and ssRNA treatment induced maturation
in bmDC from WT mice sufficiently to detect proliferation and IFNγ secretion in co-cultured
OT-1 splenocytes; however, still far less than after co-culture with bmDC derived from CB1-/CB2-/- mice with LPS or ssRNA treatment as evidenced by the percentage of proliferating cells
or percentage of proliferating and IFNγ+ cells (Figure 21 D). Overall, the contribution of bmDC
to elicit T cell responses is consistent with the overall phenotypic difference observed between
WT and CB1-/-CB2-/-mice observed after influenza infection in vivo.
AM isolated from BALF of NA WT and CB1-/-CB2-/- mice were used to extend the
findings with bmDC. As LPS was much more potent and the amount of cells available in the
BALF are limited, we reduced our study design to a single TLR agonist, especially since TLRinitiated signaling by ssRNA and LPS converges at MyD88 [327]. AM, as identified by CD11bCD11c+ expression [133], were obtained from BALF and stimulated overnight with LPS (Figure
21). As previously seen in the bmDC experiments, AM isolated from CB1-/-CB2-/- mice
exhibited a pre-activated phenotype compared to macrophages from WT mice based on the
expression of MHCII and CD86 (Figure 22 B, C). LPS stimulation again increased maturation
markers, especially MHC II (Figure 22 B, C) and AM from CB1-/-CB2-/- mice presented a more
mature phenotype than WT after TLR stimulation. Overall, the increased maturation of APC

114

from CB1-/-CB2-/- mice suggests that dysregulation due to extinguished CB1 and CB2 signaling
predominantly affects the functional regulation of APC such as DC and AM.

III. In Vivo Influenza Study 3 dpi to Study the Effect of Δ9-THC on Immune Endpoints of
WT and CB1-/-CB2-/- Mice

A) Δ9-THC Reduced Steady-state mRNA Expression of Genes Induced by Influenza in
the lungs of WT but not CB1-/-CB2-/- mice
Previously, effects of simultaneous deletion of CB1 and CB2 were assessed using the
A/PR/8/34 influenza model and 3 dpi was determined to be the peak time of the inflammatory
response. To further investigate the effects of Δ9-THC on the anti-influenza immune response in
the lungs of WT and CB1-/-CB2-/- mice, a 96-gene immune panel was employed to assess gene
expression changes. 18S rRNA was used as the loading control and all samples were normalized
to 18S rRNA gene expression. In uninfected mice, the expression of mRNA of very few genes
was altered by Δ9-THC (Figure 23 A). In contrast, PR8 induced broad steady-state mRNA
expression level changes, many of which were suppressed by Δ9-THC in WT but not CB1-/CB2-/- mice, suggesting CB1 and/or CB2-dependent modulation (Figure 23 B). In addition,
mRNA levels for only a few genes were upregulated in CB1-/-CB2-/- mice as a result of Δ9THC treatment, suggesting CB1 and CB2-independent mechanisms of Δ9-THC immune
modulation.

115

Figure 23. Reduction of Gene Expression Associated with Influenza Infection as a Result
of Δ9-THC Treatment.
Mice (n=4) were treated with corn oil vehicle (CO) or Δ9-THC (75 mg/kg) for 5 consecutive
days surrounding the intranasal instillation of PR8 or SAL. Lung RNA was isolated 3 dpi and
converted into cDNA and gene expression levels were analyzed using a Taqman low density
gene array. Fold change values were normalized with blom transformation then log
transformed, mean and median centered across genes with Cluster and visualized using
Treeview. Shown are genes significantly different between CO and Δ9-THC samples, with a
p-value of p ≤ 0.1 in SAL (A) and PR8 instilled (B) groups 3 dpi. Data were analyzed using
factorial ANOVA in SAS 9.1.3 and p-values are indicated in the left most column and
represent statistical differences as a result of Δ9-THC treatment. For interpretation of the
references to color in this and all other figures, the reader is referred to the electronic version
of this dissertation.

116

Figure 23. Reduction of Gene Expression Associated with Influenza Infection as a Result of Δ9-THC Treatment.

117

Figure 23 cont’d.

118

Figure 24. Δ9-THC does not Alter BALF Cell Counts or Composition After Influenza
Infection.
BALF was isolated by flushing of lungs of mice 3 dpi twice with 0.9 mL 0.1% saline (n=5).
Total cell numbers were counted using a hemacytometer and after Diff-Quick staining of cells
centrifuged onto slides and differential cell counts were performed. Shown are the percent of
differentially stained cells out of the total. Data were analyzed using an ANOVA: +++
(p≤0.001) difference between NA (SAL) and PR8; ### (p≤0.001), difference between WT
and CB1-/-CB2-/-.

119

Figure 25. No Change in Percent Immune Effector Cell Composition in Lungs of Mice
Treated with Δ9-THC.
Lungs were mechanically disrupted 3 dpi and single cell suspensions were obtained. Cells
were stained with fluorescently labeled antibodies for surface markers CD4, CD8, NK1.1 and
analyzed by flow cytometry. Cells were gated on single cells, size, CD4+, CD8+, or CD4CD8-NK1.1+ (A). The percent of cells in the lung is shown for each treatment group (n=5)
(B). Data were analyzed using Friedman’s test for non-parametric data: +++ (p≤0.001)
difference between NA (SAL) and PR8; # (p≤0.05), difference between WT and CB1-/-CB2/-.

120

Figure 25. No Change in Percent Immune Effector Cell Composition in Lungs of Mice Treated with Δ9-THC.

121

Figure 26. Decreased IFNγ Production after Δ9-THC Treatment in Lung-Isolated
Immune Cells from WT but not CB1-/-CB2-/- mice.
After mechanical disruption of lung tissue, single cell suspensions were restimulated in vitro
with PMA/Io (40 nM/ 0.5 µM) to induce cytokine secretion and in the presence of Brefeldin
A in 2% BCS RPMI for 5 hours to inhibit cytokine release. After restimulation, cells were
stained for CD4, CD8, and NK1.1 surface expression. On the day of flow cytometric analysis,
cells were stained for intracellular IFNγ and analyzed for fluorescence intensity (n=5). Data
were analyzed using Friedman’s test for non-parametric data: * difference between CO and
Δ9-THC ++ (p≤0.01) difference between NA (SAL) and PR8; ## (p≤0.01), ### (p≤0.001)
difference between WT and CB1-/-CB2-/-. Shown are samples concatenated (n=5) within
treatment groups.

122

Figure 26. Decreased IFNγ Production after Δ9-THC Treatment in Lung-Isolated Immune Cells from WT but not CB1-/CB2-/- mice.
123

Figure 27. Decreased IL-17 Production after Δ9-THC Treatment in Lung-Isolated
Immune Cells from WT but not CB1-/-CB2-/- mice.
Lungs were mechanically disrupted and restimulated in vitro as described before. Markers
CD4, CD8, and NK1.1 were stained on the surface of isolated cells and intracellular staining
for IL-17 and flow cytometry was performed (n=5). Non-parametric percentage data were
analyzed using Friedman’s test: * difference between CO and Δ9-THC, ++ (p≤0.01)
difference between NA (SAL) and PR8; # (p≤0.05), ## (p≤0.01), ### (p≤0.001) difference
between WT and CB1-/-CB2-/-. Displayed are concatenated samples (n=5) of each treatment
group.

124

Figure 27. Decreased IL-17 Production after Δ9-THC Treatment in Lung-Isolated Immune Cells from WT but not CB1-/CB2-/- mice.
125

B) Δ9-THC

Suppressed

Influenza-Induced

Cytokine

Production

by

Effector

Lymphocyte Populations.
Recruitment of neither leukocytes into the BALF (Figure 24) nor effector cells in whole lung
isolates was affected by Δ9-THC (Figure 25). However, greater numbers of neutrophils and
monocyte/macrophages were found in the BALF and lower numbers of CD8+ cells were found
in the lungs of CB1-/-CB2-/- compared to WT mice. As previously reported, these endpoints are
consistent with enhanced inflammatory responses observed in CB1-/-CB2-/-mice. In light of the
absence of altered cellular recruitment to the lungs by Δ9-THC, effector function was assessed in
lymphocyte populations by measuring cytokine secretion in CD4+, CD8+, and NK1.1+ cells. As
previously determined, percentages of cells staining positive for cytokine secretion were
increased in uninfected lungs of CB1-/-CB2-/- compared to WT mice (Figure 26, 27). Influenza
infection significantly induced the percentage of IFNγ-producing cells in WT CD4+ samples and
there was a trend towards induction of the IFNγ positive percentage in CD8+ and NK1.1+ cells.
Δ9-THC suppressed the percentage of IFNγ-producing populations in WT but not CB1-/-CB2-/CD4+ cells, while CD8+ and NK1.1+ cells showed a similar trend, yet were not deemed
statistically significant. The percentage of the IL-17-producing population was also induced by
influenza infection in WT mice, especially in NK1.1+ cells. The percentage of IL-17-producing
cells was suppressed by Δ9-THC significantly in NK1.1+ cells (Figure 27). Taking into account
the percentage of cells positive for cytokine production, CD8+ T cells had almost negligible
contributions to IFNγ and IL-17 production when compared to CD4+ and NK1.1+ cells (Figure
25, 26). Consistent with the histopathology as observed by H&E staining in previous
experiments [120], Δ9-THC enhanced the presence of IFNγ-producing populations in CD4+ and
NK1.1+ cells and percentages of IL-17-producing cells in CD8+ and NK1.1+ cells from CB1-/-

126

CB2-/- mice, which was significantly different from Δ9-THC-treated WT samples (Figure 26,
27). Collectively, Δ9-THC suppressed the percentages of functional effector CD4+ and NK1.1+
cells 3 dpi in a CB1 and/or CB2-dependent manner.

C) No Effect of Δ9-THC on Induction of the CD11b+Gr-1+ Population After Influenza
Infection
One potential mechanism for a reduction in cytokine production by Δ9-THC in effector
lymphocyte populations is the induction of immune suppressor populations, among which are the
myeloid-derived suppressor cells (MDSC). MDSC are identified as CD11b+Gr-1+ and can be
induced during infection and inflammation [150]. A slightly higher basal presence of MDSC was
observed in CB1-/-CB2-/- compared to WT mice (Figure 28). Influenza infection drastically
induced MDSC populations in WT and CB1-/-CB2-/-, but no significant effect of Δ9-THC on
the induction of MDSC was observed. Albeit not statistically significant, higher percentages of
MDSC were observed in CB1-/-CB2-/- compared to WT mice after influenza infection,
consistent with previous reports of greater inflammatory responses in these mice. Thus these
MDSC seem to be attracted to inflammatory environments and may be one of several
mechanisms to ameliorate inflammation due to influenza infection. On MDSC, the expression of
MHC I, MHC II and CD86 was greatly induced after influenza infection (Figure 29). Δ9-THC
administration had no effect on the expression levels of MHC I and CD86 after influenza
infection, but there were trends showing a decrease in MHC II in WT and an increase in MHC II
in CB1-/-CB2-/- mice. Overall, there is no direct evidence for the contribution of MDSC to the
Δ9-THC-mediated suppression of cytokine production in lymphocytes, but these cells might play
a role in suppressing influenza-induced inflammation [148].

127

Figure 28. No Effect by Δ9-THC Treatment on Presence of Influenza Infection-Induced
Gr-1+CD11b+ Lung-Isolated Cell Populations. Cells were isolated from lungs as described
before. Cell surfaces were stained with fluorescently labeled antibodies to Gr-1 and CD11b
and analyzed using a flow cytometer (n=5). Percentage data were analyzed using Friedman’s
test: +++ (p≤0.001) difference between NA (SAL) and PR8; ## (p≤0.01) difference between
WT and CB1-/-CB2-/-. Shown are concatenated samples (n=5) within each treatment group.

128

Figure	
  28.	
  No	
  Effect	
  by	
  Δ9-­‐THC	
  Treatment	
  on	
  Presence	
  of	
  Influenza	
  
Infection-­‐Induced	
  Gr-­‐1+CD11b+	
  Lung-­‐Isolated	
  Cell	
  Populations.	
  

	
  

129

Figure 29. Δ9-THC Treatment does not Alter the Maturation Status of Gr-1+CD11c+
cells. Lung-isolated cells were gated on the Gr-1+CD11b+ population shown in Figure 6.
Within the gated Gr-1+CD11b+ population, expression of maturation markers MHC I, MHC
II, CD86 was assessed using fluorescence intensity after flow cytometric analysis (n=5). The
data as shown was concatenated within each group. Data were analyzed using ANOVA: +++
(p≤0.001) difference between NA (SAL) and PR8.

130

ß	
  
Figure	
  29.	
  Δ9-­‐THC	
  Treatment	
  does	
  not	
  Alter	
  the	
  Maturation	
  Status	
  of	
  Gr-­‐1+CD11c+	
  cells.	
  
131

Figure 30. Δ9-THC Reduces Recruitment of pDC, cDC, and AM into the Lungs of
PR8-Infected Mice Dependent of CB1 and/or CB2. Mice (n=5) were intranasally
instilled with saline (NA) or 50 pfu of PR8 virus and 3 dpi lungs were excised en bloc and
homogenized by mechanical disruption. Single cell suspensions obtained from the lung
were stained for CD11b, CD11c, Gr-1, MHC I, MHC II, and CD86. CD11b, CD11c, and
Gr-1 were used as markers to identify separate cell populations as shown in the gating
scheme (A), which were then enumerated by percentage (B). Friedman’s test was used to
perform statistics on non-parametric percentage data: ** (p≤0.01), * (p≤0.05) difference
between CO and Δ9-THC, ++ (p≤0.01), +++ (p≤0.001) difference between NA (SAL) and
PR8; ## (p≤0.01) difference between WT and CB1-/-CB2-/-.

132

	
  
Figure	
  30.	
  Δ9-­‐THC	
  Reduces	
  Recruitment	
  of	
  pDC,	
  cDC,	
  and	
  AM	
  into	
  the	
  Lungs	
  of	
  PR8-­‐Infected	
  Mice	
  Dependent	
  of	
  CB1	
  
and/or	
  CB2.	
  
133

D) Reduced Presence of APC with Δ9-THC Treatment After Influenza Infection
To become functional effectors, lymphocytes depend on the presence of, and interaction with,
APC. In the lung, several populations of immune cell populations including APC are present and
can be identified according to their surface expression of CD11b, CD11c, and Gr-1 [131, 328]
(Figure 30 A). Granulocytes are CD11b+CD11c-, macrophages are CD11bloCD11clo, cDC are
CD11b+CD11c+, while AM are CD11c+CD11b- as well as Gr-1-. Similar to AM, pDC as also
express CD11c and are identified as CD11b-CD11c+Gr-1+. The presence of pDC, cDC, and
granulocytes was significantly increased in both WT and CB1-/-CB2-/- mice as a result of
influenza infection, while there was no change in the macrophage/monocyte (Mac/MФ)
population (Figure 30 B). Furthermore, greater percentages of AM were found in lungs of CB1-/CB2-/- compared to WT mice after influenza infection. The increase of pDC, cDC, and AM after
infection was attenuated in Δ9-THC-treated WT but not CB1-/-CB2-/-mice, suggesting CB1and/or CB2- dependence of the Δ9-THC effect (Figure 29 B). In summary, a reduction in
lymphocyte effector function by Δ9-THC might be attributed to reduced numbers of APC,
specifically, AM, cDC and pDC present in the lungs in Δ9-THC-treated WT mice with influenza.

A) Greater Maturation in APC from CB1-/-CB2-/- Compared to WT and CB1-and/or
CB2-Dependent Suppression of Maturation by Δ9-THC after Influenza Infection.
Maturation markers MHC I, MHC II and CD86 were assessed in APC subpopulations: cDC
(Figure 31), pDC (Figure 32), AM (Figure 33) and Mac/MФ (Figure 34). No changes in
expression of maturation markers as a result of influenza infection were observed in cDC or AM
(Figure 31 and 33); however, in pDC and Mac/MФ surface expression of MHC II and CD86 was

134

Figure 31. Δ9-THC does not Alter the Maturation Status of Lung-Isolated cDC after
PR8 Infection. Within CD11b+CD11c+ population (gates shown in Figure 30), maturation
markers MHC I, MHC II, CD86 were determined by flow cytometry (n=5) (A). The data as
shown was concatenated within each treatment group. Bar graphs show mean fluorescence
intensities (MFI) for indicated maturation markers (B) (n=5). Analysis was performed using
ANOVA (B): # (p≤0.05), difference between WT and CB1-/-CB2-/-.

135

	
  
Figure	
  31.	
  Δ9-­‐THC	
  does	
  not	
  Alter	
  the	
  Maturation	
  Status	
  of	
  Lung-­‐Isolated	
  cDC	
  after	
  PR8	
  Infection.	
  
136

Figure 32. Suppression of MHC II Expression in pDC by Δ9-THC after PR8 Infection.
Within the CD11b-CD11c+ population (gates shown in Figure 30) the Gr-1+ pDC population
(Figure 8 A) was assessed for expression of maturation markers MHC I, MHC II, and CD86
by flow cytometry (A) (concatenated, n=5). Bar graphs are indicating MFI (B) (n=5). Samples
were analyzed using ANOVA (B): * (p≤0.01), difference between Δ9-THC and VH (CO);
+++ (p≤0.001) difference between SAL and PR8; #, (p≤0.05), ### (p≤0.001), difference
between WT and CB1-/-CB2-/-.

137

Figure 32. Suppression of MHC II Expression in pDC by Δ9-THC after PR8 Infection.
138

Figure 33. Δ9-THC does not Alter the Maturation Status of Lung-Isolated AM after
PR8 Infection.
Within CD11b-CD11c+ population (gates shown in Figure 30) the Gr-1- AM population
(Figure 8 A) was gated and expression of maturation markers MHC I, MHC II, and CD86
was assessed by flow cytometry (concatenated, n=5). Bar graphs indicate MFI (C) (n=5).
Samples were analyzed using ANOVA.

139

Figure 33. Δ9-THC does not Alter the Maturation Status of Lung-Isolated AM after PR8 Infection.
140

Figure 34. Δ9-THC does not Alter the Maturation Status of Lung-Isolated
Macrophages/Monocytes after PR8 Infection.
Cells were gated on the CD11b lo CD11c lo population for macrophages and/or monocytes as
shown in Figure 30. Expression of maturation markers MHC I, MHC II, and CD86 was
assessed by FACS (shown are concatenated samples, n=5 per group) (A). Bar graphs were
generated using MFI output of FACS data from (n=5) (B). Samples were analyzed using
ANOVA: +++ (p≤0.001) difference between SAL and PR8; #, (p≤0.05), ### (p≤0.001),
difference between WT and CB1-/-CB2-/-.

141

Figure 34. Δ9-THC does not Alter the Maturation Status of Lung-Isolated Macrophages/Monocytes after PR8 Infection.
142

increased as a result of influenza infection (Figure 32 A, B and Figure 34 A, B). More robust
expression levels of MHC II were observed in cDC, pDC and Mac/MФ from CB1-/-CB2-/mice compared to WT mice after influenza infection (Figure 31, 32, and 34). Δ9-THC
suppressed MHCII surface expression in pDC in a CB1- and/or CB2- dependent manner (Figure
32) although similar trends were observed in MHCII and CD86 in AM (Figure 33) and MHCII in
Mac/MФ (Figure 34) albeit not statistically significant. Taken together, these results suggest that
to a great extent the suppression of lymphocyte effector function by Δ9-THC occurs by reducing
the migration of APC into the lungs, rather than suppressing the maturation status on APC.
Furthermore, greater maturation was observed in inflammatory APC such as pDC and Mac/MФ
in CB1-/-CB2-/- compared to WT mice.

B) Attenuation of bmDC Maturation by Δ9-THC In Vitro
An alternate explanation for the reduced presence of cDC, pDC and AM in the lungs as a result
of Δ9-THC treatment without changes in maturation, could be that migration is dependent on the
maturation status. Indeed, only mature DC are able to migrate, thus it is plausible that the site of
infection is enriched for mature DC as immature DC are not able to migrate to the site of
infection [329-331]. However, Δ9-THC might elicit its effects on the maturation of DC, thereby
reducing their migration to the site of infection. An in vitro bmDC model was used to study the
effects of Δ9-THC on DC maturation induced by TLR ligation. These bmDC were identified as
CD11b+CD11c+ and thus similar to the cDC phenotype (Figure 35 A). LPS (1µg/mL), a TLR4
ligand [332] significantly induced maturation in WT and CB1-/-CB2-/- bmDC, as evidenced by
MHC II, CD80 and CD86 expression (Figure 35 B, C). In contrast, R848 (5µg/mL), a TLR 7/8
agonist [333] did not induce bmDC maturation to the same extent as LPS, which was especially

143

Figure 35. Δ9-THC Suppresses TLR-Stimulated bmDC Maturation Independent
of CB1 and CB2.
Bone marrow was used to generate bmDC in the presence of GMCSF (20 ng/mL) for 9
days. bmDC were washed and incubated for 24 hours in the presence or absence (NA)
of LPS (1µg/mL) or R848 (5µg/mL) and co-treated with VH (0.1% ethanol) or Δ9THC (10 µM). After incubation, staining for CD11b, CD11c (not shown), MHC I,
MHC II, and CD86 were performed as previously described. Cells obtained after
culture were CD11b+CD11c+ (A), indicating cDC phenotype. Graphs shown are
concatenated samples (B, n=3). ANOVA tests were performed on samples from 2 A
and B (C), ** (p≤0.01), difference between Δ9-THC and VH (CO); ++ (p≤0.01), +++
(p≤0.001) difference between SAL and PR8; ## (p≤0.01), difference between WT and
CB1-/-CB2-/-. The experiment is representative of three identical repeat experiments.

144

Figure 35. Δ9-THC Suppresses TLR-Stimulated bmDC Maturation Independent
of CB1 and CB2.

145

Figure 35 cont’d.

146

noticeable for CD86 expression (Figure 35 B, C). As previously observed, greater maturation
levels as evidenced by MHCI and MHCII expression were observed in unstimulated bmDC
generated from CB1-/-CB2-/- when compared to WT. Δ9-THC (10 µM) reduced the LPS- but
not R848-induced maturation, as evidenced by lower MHC II in CB1-/-CB2-/- and CD86
expression in WT and CB1-/-CB2-/-. Collectively, bmDC maturation was suppressed by Δ9THC independent of CB1 and/or CB2.

C) Impaired Elicitation of T cells after Δ9-THC Treatment of bmDC
To determine the functional consequences of Δ9-THC-mediated suppression of TLRstimulated maturation of bmDC, a T cell co-culture model was used. Cells from OT-1 mice with
a transgenic T cell receptor in the CD8+ T cell compartment specific to cognate epitope
OVA257-264 (SIINFEKL) were incubated with bmDC pulsed with SIINFEKL. The ability of
OT-1 CD8+ T cells to proliferate and secrete IFNγ is a direct result of the interaction with
bmDC, and is thus a functional assay for the ability of bmDC to elicit T cells. Four days after coculture, OT-1 cells were restimulated with SIINFEKL and after staining with fluorescently
labeled antibodies were gated as described in methods to exclude any cells other than CD8+ T
cells (Figure 36 A). No proliferation was observed in T cell co-cultures with NA WT bmDC in
contrast to bmDC from CB1-/-CB2-/-, which readily elicited T cells without the need for prior
stimulation as previously reported (Figure 36 B). While LPS- or R848-treated bmDC induced
antigen-specific T cell proliferation in WT, Δ9-THC (10 µM) potently reduced proliferation. Due
to the high response magnitude in naive bmDC from CB1-/-CB2-/- mice, TLR stimulation had
little effect on T cell elicitation and Δ9-THC treatment of CB1-/-CB2-/- bmDC did not alter their
ability to elicit T cells (Figure 36 B). In addition to proliferation, it is important to determine T

4
137

Figure 36. Δ9-THC Impairs Antigen-Specific bmDC-Elicited T cell responses.
bmDC, NA and TLR treated, with VH (0.1% ethanol) or Δ9-THC were pulsed with the OT-1
TCR-specific peptide SIINFEKL for 1 hour and washed three times prior to incubation with
Cell Trace-labeled OT-1 splenocytes for 4 days. Cells were restimulated with SIINFEKL and
stained for CD8 and IFNγ and gated as depicted in the scheme to obtain dot plots (A). OT-1
cells were co-cultured with bmDC and then gated as described in 3 A. Dot plots with Cell
Trace loss indicating proliferation on the X-axis and CD8 staining on the Y-axis are shown.
Gate shown indicates proliferation (loss of Cell Trace fluorescence compared to control) and
CD8 staining (B) or IFNγ staining (C). Shown are concatenated samples of each group (n=3).
Kruskal-Wallis tests were performed on samples from 3 B-E, * (p≤0.05), difference between
Δ9-THC to VH (CO); ++ (p≤0.01), ## (p≤0.01), ### (p≤0.001) difference between WT and
CB1-/-CB2-/-. The experimental data is representative of two identical repeat experiments.

148

Figure 36. Δ9-THC Impairs Antigen-Specific bmDC-Elicited T cell responses.
149

Figure 36 cont’d.

150

secretion of cytokines. Concurrent analysis of IFNγ secretion and proliferation was performed
after elicitation. Similar to the proliferation results, without TLR stimulation no cytokine release
was observed in OT-1 cells elicited by co-culture with SIINFEKL-pulsed WT bmDC, but
cytokine release was induced with TLR stimulation and significantly suppressed with Δ9-THC
(10 µM) treatment (Figure 36 C). CB1-/-CB2-/- bmDC elicited OT-1 T cell responses of high
magnitude without the need for prior stimulation and were resistant to Δ9-THC suppression
(Figure 36 C). Overall, these results suggest that CB1 and/or CB2 are important in the
suppression of DC-mediated T cells responses by Δ9-THC. Furthermore, in light of maturationdependent migration, the impairment of DC maturation by Δ9-THC might consequentially
attenuate DC migration, thereby reducing effector cell responses in the lung after influenza
infection.

151

DISCUSSION

I.

Suppression of the CTL Response by Δ9-THC

The observation that the influx of CD8+ T cell populations after influenza infection in vivo
was greatly impaired in response to Δ9-THC treatment warranted further studies to characterize
the effect of Δ9-THC on CD8+ T cells at the cellular level [119]. A model using P815 cells as an
allogeneic stimulus to elicit CTL and cells isolated from CB1-/-CB2-/- and WT mice was
employed to investigate the effect of Δ9-THC on the generation of of CTL in vitro. Collectively,
these studies provide evidence that Δ9-THC-mediated suppression of CD8+ T cell function
occurs independent of CB1 and CB2. Moreover, Δ9-THC alters early signaling events that
involve putative changes in intracellular Ca2+ levels, eventually resulting in lower numbers of
CTL effectors. Finally, this study introduces an inherent role for CB1 and/or CB2 in the
elicitation of CTL, because elicitation and function of CTL was reduced in CB1-/-CB2-/- mice in
vitro.

A) Reduced CTL Response in CB1-/-CB2-/- Compared to WT mice
In contrast to other reports describing enhanced immune responses in CB1-/-CB2-/- mice
[120, 315], in this in vitro model of alloantigen-induced CTL elicitation and effector function,
CTL activity in cells from CB1-/-CB2-/- mice was reduced when compared to WT mice.
Although this was consistent with our previous study in which we observed a reduction in the T
cell-dependent humoral immune response to sheep erythrocytes in vitro in CB1-/-/CB2-/- as
compared to WT mice [279], there are several studies demonstrating enhanced immune function
in CB1-/-CB2-/- mice [120, 315]. In a previous study from this laboratory [120], it was

152

demonstrated that CB1-/-CB2-/- mice, in addition to responding with a greater magnitude to
immune stimuli, have altered immune homeostasis compared to WT mice as evidenced by
greater numbers of CD4+ T cells and higher levels of TNF-α in the BALF in the absence of
influenza challenge. In addition, there appeared to be mechanisms in CB1-/-CB2-/- mice that
might compensate for the loss of CB1 and/or CB2, as evidenced by greater levels of the
endocannabinoids, 2-arachidonoylglycerol and anandamide [315], and increased transcripts of
cnr2 using primers specific for the undeleted portion of the gene [334]. Although CB1-/-CB2-/mice seem to be particularly vulnerable to proinflammatory insult, there were no drastic
differences observed in an adjuvant-free airway hypersensitivity model between CB1-/-CB2-/and WT mice [317]. Reasons for reduced function of CTL from CB1-/-CB2-/- mice compared to
WT mice, although not altogether clear, might be due to the present studies being conducted in
vitro, and/or the nature in which the immune response was initiated, in this case by direct
stimulation of CD8+ T cells with an alloantigen.
Greater proliferation was detected in CD4+, but lower proliferation in CD8+ cells, from
CB1-/-CB2-/- compared to WT mice. This lower proliferation in CD8+ cells might contribute to
the reduced CTL activity in CB1-/-CB2-/- compared to WT mice.

Furthermore, higher

percentages of CD4+CD25+ were observed in unstimulated splenocytes of CB1-/-CB2-/compared to WT mice. These CD4+CD25+ cells are thought to be regulatory T cells (Treg)
[335], which are involved in the suppression of the immune responses. It has been demonstrated
that an increase in CD4+CD25+ cells leads to reduced CTL responses; thereby Tregs could
potentially contribute to the reduced activity in vitro in CTL generated from CB1-/-CB2-/- mice.
[336]. This in vitro model, due to direct stimulation of CTL, lacks a proinflammatory stimulus
and does not produce IL-6, which has been implicated in overriding Treg responses, thus Treg

153

responses might be maintained during the in vitro culture [337]. As our focus was on the Δ9THC-mediated suppression of CTL responses, we did not further characterize these
CD4+CD25+ cells for expression of Foxp3 or other cellular markers of the Treg lineage [338].
Additional studies are required to determine the role of Tregs in CB1-/-/CB2-/-mice.

B) Sensitivity of CTL to Δ9-THC Immune Modulation
Addition of Δ9-THC during the elicitation and effector response showed that only naïve
CD8+ cells, but not differentiated CTL, were sensitive to Δ9-THC-mediated suppression of CTL
function as assessed by 51Cr release assay, suggesting Δ9-THC-mediated immune modulation
during elicitation. Arguably, the concentrations of ∆9-THC at which altered CTL activity was
observed in this investigation are higher than detected in the serum of recreational marihuana
smokers. However, it is important to emphasize two points. The first is that in addition to ∆9THC, marihuana smoke contains over 60 other structurally-related cannabinoids, a number of
which are also well established immune modulators and the majority of which remain to be
evaluated for immunomodulatory activity [339, 340]. Second, and equally important, the goal of
this study was to relate the decrease in viral clearance observed after ∆9-THC treatment of
animal with influenza-infected airways [119] and the possibility that this may be due to
cannabinoid-mediated suppression in CTL activity. In the case of inhaled marihuana smoke, the
cannabinoid concentrations attained in the lung are significantly higher than systemic serum ∆9THC concentrations and may be more closely related to those used in the current study [341,
342].
prior

The timeframe of sensitivity to Δ9-THC might reflect a high level of T cell plasticity
to

differentiation

and

reduced

plasticity

after

differentiation

[343].

Other

immunomodulatory compounds such as 2,3,7,8-tetrachlorodibenzo-p-dioxin and glucocorticoids

154

exert their greatest effect on early stages of immune cell differentiation and tend to have a lower
efficacy if added after the onset of the differentiation program [344, 345]. Specifically,
cyclosporin A is very efficacious in suppressing in vitro culture of allogeneic cells only when
added prior to elicitation [346]. Overall, this suggests a critical window of sensitivity for
immunomodulatory compounds, including Δ9-THC, for immune suppression, specifically during
the transition of naïve CD8+ T cells to CTL during elicitation.
Using IFNγ as a surrogate for CTL activity, it was demonstrated that CD8+ cells are the
sole producers of IFNγ after allogeneic stimulation, suggesting that only CTL directly contribute
to the 51Cr release from labeled target cells. Although it appeared that Δ9-THC increased IFNγ
expression in CTL, the cells were gated on the live cell populations. Upon examination of the
results from the total population, there was no effect of Δ9-THC on IFNγ, suggesting alternative
interpretations of the data. Taken together, our results suggest that while there seemed to be a
population of CTL whose activity cannot be suppressed by Δ9-THC, which become the IFNγsecreting cells, the remaining cells are sensitive to Δ9-THC and were not capable of surviving
the five-day culture period. Thus, the suppression by Δ9-THC does not appear to reduce the
amount of IFNγ each cell produces, but rather impaired the differentiation of a select pool of
naïve cells from becoming effector CTL.

C) Alterations in T cell Activation by Δ9-THC
No effects of Δ9-THC treatment on the proliferation of CD4+ or CD8+ cells WT or CB1/-CB2-/- mice were observed. To link the functional outcome of Δ9-THC treatment to earlier T
cell signaling, the expression of activation markers, CD25, forming the high affinity IL-2
receptor, and CD69, the earliest known lymphocyte activation marker [325] were assessed.

155

While CD25 expression was increased as a result of P815 co-culture, only CD8+ cells from
CB1-/-CB2-/- mice had reduced CD25 expression as a result of Δ9-THC treatment. CD69
expression was also increased as a result of co-culture with P815 cells in both WT and CB1-/CB2-/- CD8+ cells and Δ9-THC treatment further enhanced the P815-induced rise in CD69
expression in both genotypes. The ability of Δ9-THC to increase CD69 expression on the surface
of CD8+ T cells from WT and CB1-/-CB2-/- after P815 cell stimulation provides further
evidence for CB1 and/or CB2-independent mechanism(s) of Δ9-THC immune modulation.
Notably, expression of CD69 was inversely correlated with CTL activity. For example, the
highest CD69 expression and lowest CTL activity were observed in cells treated with the highest
concentration of Δ9-THC (10 µM) from CB1-/-CB2-/- mice.
CD69 expression is controlled by several transcription factors, including AP-1 in the
proximal promoter [347] as well as EGR, ATF/CREB [348], and NFκB [325]. However, the
pathways regulating the expression of CD69 are different depending on the cell type [349]. It is
not entirely clear what cellular signals contribute to CD69 induction in CD8+ T cells, but it is
known that Ca2+ influx into T cells increases CD69 expression [350] and, at least in thymocytes,
extracellular Ca2+ was found to be critical for CD69 induction [351]. In these studies Δ9-THC
synergized with Io to increase CD69 expression suggesting that Ca2+ plays an important role in
the Δ9-THC-mediated increase of CD69 expression. Previously, this laboratory reported that
Δ9-THC strongly induced intracellular Ca2+ in T cells, which was mediated by TRPC1 [233,
284] independent of the requirement for T cell activation. These studies, taken together with
previous results, suggest that modulation of an activation stimulus by concurrent induction of
Ca2+ through Δ9-THC-mediated opening of TRPC1 channels contributes directly to the
suppression of CTL activity. In addition, there are reports indicating that Δ9-THC has a high

156

binding affinity for GPR55 [352] and causes increases in intracellular Ca2+ levels via GPR55
[353], but there are also conflicting reports demonstrating that Δ9-THC possesses a low affinity
for GPR55 [354]. The role of GPR55 in the present study is unknown; however, due to the
expression of GPR55 in the spleen [352] and its ability to modulate Ca2+ currents, it is tempting
to speculate that GPR55 contributes to alterations in Ca2+ signaling induced by Δ9-THC. Ca2+
is a ubiquitous intracellular signaling molecule and, in addition to activation, Ca2+ is central to
the induction of anergy and cell death, which are seemingly separate processes, yet involve
shared mechanisms and endpoints [355, 356]. Thus, due to the role of Ca2+ in the generation of
CTL, it seems plausible that Δ9-THC-mediated alterations in Ca2+-signaling are involved in the
suppression of CTL responses. An alternate explanation for the synergism of Δ9-THC with Ioinduced CD69 levels might involve MAPK activation of AP-1. For example, it has been
demonstrated that the βγ subunits of GPCRs as well as β -arrestin are responsible for MAPK
activation [244, 248, 249]. Although plausible, this scenario is speculative but has been observed
for Gαi-coupled receptors.

D) Summary and Conclusion of the Effect of Δ9-THC on CTL
Collectively, the reduced cytolytic activity, decreased number of CTL effectors, and the
increase in CD69 expression, although temporally distinct, might be interrelated, and a
consequence of early Δ9-THC-induced changes in Ca2+ signaling. Concordantly, immune cells
appear to initially undergo greater activation (CD69), but become unresponsive to secondary
stimulation; in essence, anergic.

The critical role of calcium in leukocyte activation and

subsequent differentiation and effector function is well established. Δ9-THC treatment prior to
cellular activation significantly elevates intracellular Ca2+ levels, which in turn interferes with

157

either entry or execution of the T cell differentiation program.

The specific downstream

signaling pathways affected through the Δ9-THC-mediated elevation of intracellular Ca2+ levels
affecting T cell function remain to be elucidated but likely involve p38 MAPK, JNK, and ERK.
Indeed, prior studies by this laboratory demonstrated decreased ERK phosphorylation by
cannabinol, another cannabinoid acting independently of CB1 and CB2 [357]. Thus, the
immediate perturbation of intracellular Ca2+ levels by cannabinoids, including Δ9-THC, seems
to be the initial event triggering a cascade of events, eventually resulting in immune suppression.
Signaling by cannabinoids seems to alter the regulation of Ca2+ that is central to the initiation of
a T cell response to impair immune function.

II.

Influence of Cannabinoid Receptors 1 and 2 in on the Magnitude of the Immune
Responses to Influenza

Although the experiments performed on CTL in vitro provided insights into the suppression
of CTL function by Δ9-THC, the cellular mechanisms contributing to lower viral burden in
CB1-/-CB2-/- compared to WT mice remained to be elucidated. Cannabinoid receptors and their
endogenous ligands have been investigated extensively for their immunomodulatory activity;
however, only a few studies have investigated the outcome of concomitant targeted deletion of
both CB1 and CB2 on immune competence [120, 279, 315]. Based on a sizable literature
demonstrating that plant-derived, synthetic and endogenous cannabinoids typically attenuate
immune responses [196, 358], it has been long hypothesized that one role of CB1 and CB2 might
be to contribute to immune homeostasis by preventing overstimulation of the immune system.

158

Our findings indicate that a principle function of cannabinoid receptors is to modulate APC
function and a lack thereof is critically involved in the described hyper-responsive immune
phenotype observed in CB1-/-CB2-/- mice. To assess the role of CB1 and CB2 in the
immunopathology induced by influenza, the majority of this investigation was focused on events
at 3 dpi, the peak day of the inflammatory response in this A/PR/8/34 influenza model . The
inflammatory response is to a great extent responsible for the immunopathology observed as a
result of influenza infection.

A) Greater Pathology in CB1-/-CB2-/- Compared to WT Mice after Influenza Infection
After influenza infection, immunopathology was evident in lungs of CB1-/-CB2-/- mice to a
greater degree than in lungs of WT mice as observed by morphologic differences in H&E
staining and the loss of CCSP. In addition, the influx of macrophages and neutrophils into the
airways was more pronounced with a concomitant increase in necrosis and exfoliation of surface
epithelium surface in CB1-/-CB2-/- compared to WT mice. Since the viral burden was not
different between WT and CB1-/-CB2-/- mice, the damage to the respiratory epithelium likely
resulted as a consequence of an exacerbated immune response. The inflammatory mediators of
macrophages and neutrophils are likely responsible for damage to the airways [359, 360]. Within
the airways, CCSP is thought to perform a protective function, especially involving the reduction
of neutrophilia due to viral infections, such as respiratory syncytial virus [361]. Thus, the
increase in neutrophils in the BALF of CB1-/-CB2-/- mice is consistent with the observed
decrease in CCSP and increased airway injury in CB1-/-CB2-/- compared to WT mice. In WT
mice CB2 levels did not change after PR8 infection, however CB1 levels were increased
suggesting a potential role for CB1 in the amelerioration of immune responses in the lung.

159

B) Altered T cell Responses in CB1-/-CB2-/- Compared to WT Mice after Influenza In
Vivo But not after T cell Stimulation In Vitro
Several unique gene expression changes were evident in CB1-/-CB2-/- compared to WT
mice after infection at 3 dpi. In contrast to basal differences in transcription, several lymphocytespecific cytokines (IL-13 and IL-17) and indices of lymphocyte activity (CD38, found on
cytokine releasing T cells, especially Th17 [362, 363]) were upregulated after influenza
infection. The presence of cytokine producing lymphocytes was confirmed by flow cytometry
and greater

percentages of IFNγ- and IL-17-producing CD4+, CD8+, and NK1.1+ were

observed in CB1-/-CB2-/- mice compared to WT mice after influenza infection. These findings
are in accordance with accelerated activation as evidenced by CD69 staining on the surface of
CD4+ and CD8+ cells. Taken together, exacerbated lymphocyte responses were observed in
CB1-/-CB2-/- compared to WT mice.
To investigate whether inherent differences in the function of WT and CB1-/-CB2-/- are
present in the lymphocyte compartment, the ability of naïve (CD62L+) and non-naïve (CD62L-)
CD4+ T cells to polarize toward a Th17 phenotype after αCD3/αCD28 stimulation was
examined. The levels of IL-17 produced from CB1-/-CB2-/- CD4+ T cells were not significantly
different from WT. Consistent with in vitro studies of mixed lymphocyte responses and PMA/Io
stimulation [279], cytokine production was similar after in vitro stimulation of naïve CD4+ T
cells from CB1-/-CB2-/- and WT mice, suggesting no inherent difference in T cell function
between the two genotypes.

C) Dysregulation of APC Responses in CB1-/-CB2-/- Mice Compared to WT mice

160

During T cell isolation consistently lower numbers of naïve T cells were isolated from
CB1-/-CB2-/- mice, and this phenomenon was confirmed by lower percentages of naïve
(CD62L+ and CCR7+CD62L+) CD4+ T cells in spleens of CB1-/-CB2-/- compared to WT
mice. Accordingly, CCL19 mRNA levels, the chemokine that binds to CCR7, were reduced in
CB1-/-CB2-/- mice. In an IκB transgenic mouse model, enhancement of NFκB signaling reduced
the naïve T cell compartments as well, while at the same time, greater IL-17 production was
observed [364]. In the present study, a similar scenario was observed, in which increased
inflammation resulted in the generation of more Th17 cells at the expense of the naïve T cell
compartment. Collectively, with no observable difference of in vitro cytokine production from
naïve CD4+ T cells, evidence for greater numbers of non-naïve T cells suggested involvement of
APC in eliciting T cell responses rather than inherent differences in the T cell compartment due
to loss of CB1 and CB2. Thus, the environment encountered and the ability of APC to elicit
stronger lymphocyte responses provided a plausible explanation for the enhanced cytokine
production by lymphocytes in vivo and reduction in the naive lymphocyte compartment in vivo
observed in CB1-/-CB2-/- mice.
Indeed, profound differences in APC maturation of CB1-/-CB2-/- compared to WT mice
were observed. Basal and TLR-inducible maturation of bmDC and AM was more robust in CB1/-CB2-/- mice as assessed by expression of surface markers CD86, MHC I, and MHC II.
Consistent with this was the observation of increased mRNA levels of H2Eb1, one of the MHC
II subunits, in the lung. In CB1-/-CB2-/- mice, increased expression of CCL3, IL-1, and
cyclooxygenase-2 (Ptgs2) genes were observed, which are often produced by APC and
characteristic of an inflammatory response [365]. Concordantly, but less well understood and
characterized, Tfrc, an iron transporting receptor has been reported to be downregulated in

161

macrophages during chronic inflammation as well as in response to inflammatory stimuli [366,
367]. Taken together, a very consistent profile of increased inflammation and maturation of APC
exists in lungs of CB1-/-CB2-/- compared to WT mice.
The consequence of APC maturation was reflected in the marked differences in the
ability of bmDC pulsed with SIINFEKL to stimulate proliferation in OT-1 cells and their ability
to become functional effectors as assessed by IFNγ secretion. While this model did not
investigate the role of CB1 and CB2 in antigen processing and presentation by DC, it has been
previously demonstrated that cannabinoids including Δ9-THC, can suppress APC function [368,
369]. Furthermore, it has been previously reported that Δ9-THC-mediated suppression of costimulatory activity by macrophages was dependent on CB2 [312, 370, 371]. Most recently, CB1
was also shown to play a role in regulation of co-stimulation by DC [266]. Only after TLR
stimulation were measurable OT-1 responses elicited by bmDC from WT, while the ability of
bmDC from CB1-/-CB2-/- surpassed that of WT with the same TLR stimulation. These in vitro
studies demonstrate, that at the cellular level, altered function of APC, especially DC, is critical
in the observed phenotypic differences between CB1-/-CB2-/- and WT mice.

D) A Role For CB1 and/or CB2 in Regulating the Magnitude of Immune Response to
Influenza to Reduce Pathology
While the phenotypic changes as a result of CB1 and CB2 deletion in the immune system
described above appear to have improved resistance to virus infection as evidenced by a reduced
lung viral burden after influenza challenge [120], our results indicate that this occurs with a
concomitant increase in immunopathology. The drastic loss of CCSP staining and greater
inflammation as evidenced by H&E staining in addition to cytokine production by NK and T

162

cells and lung mRNA transcript levels for proinflammatory mediators suggest that while CB1-/CB2-/- mice have lower viral burden in the lung [120], the lung tissue damage and airway injury
was more severe when compared to WT mice. Excessive immune activation can result in
lethality from influenza virus infection [372, 373], although in the present study, a sublethal
inoculum of influenza virus was administered to avoid lethality to the murine host [118]. Taken
together, the loss of a mechanism to negatively regulate immune function, as shown in CB1-/CB2-/- mice, resulted in increased immunopathology to a pathogenic stimulus, in this case
influenza.
Our results suggest that CB1 and/or CB2, although reducing the magnitude of the
influenza-induced immune response, provide protection against excessive airway injury in the
lung from exacerbated immunopathology. In humans, single nucleotide polymorphisms (SNPs),
which can result in a loss of function, have been identified in CB1 and were associated with
ulcerative colitis [319]. Similarly a SNP in CB2 was associated with an increased overall risk of
autoimmune disease [318]. In autoimmune diseases such as Crohn’s disease and experimental
autoimmune encephalomyelitis, a murine multiple sclerosis model, increased maturation of DC
was observed [374, 375]. Furthermore, it has been demonstrated that DC can induce and
contribute to maintaining autoimmunity [376]. Taken together with the present studies, CB1
and/or CB2 might be considered host factors that attenuate immune responses, in this case to
influenza, in order to reduce tissue damage. In humans, who, unlike laboratory animals, are
routinely exposed to pathogenic bacteria, viruses, fungi and their PAMPs, DCs confer immunity
by acting as accessory cells in the elicitation of effector cell responses (e.g., T cells). However,
exacerbated and/or unwanted DC responses can occur through a reduction or absence of
signaling through CB1 and/or CB2 for example as a result of a loss of function SNP.

163

Consequently, in the absence of regulation by the cannabinoid system, DC-elicited effector
responses might contribute to the pathologies described above including autoimmune disease.
Overall, these findings provide evidence that the enhanced immune phenotype in CB1-/-CB2-/mice after influenza infection influences a cascade of events that exist, in part, due to altered
basal immune function and functional exacerbation of select immune populations after immune
stimulation. As demonstrated with NA cells in vitro, our results suggest that the lymphocyte
compartment is not inherently responsible for exacerbated cytokine production, but is dependent
on dysregulation of APC function as a result of extinguished cannabinoid receptor signaling.
Collectively, these results indicate that the cannabinoid receptors play a fundamental role in
attenuating inflammation and immune responses orchestrated by APC.

III.

Role of the Cannabinoid Receptors 1 and 2 in Δ9-THC-mediated Suppression of
Immune Responses to Influenza Virus Challenge

The studies characterizing and comparing the kinetics of the immune response to influenza in
WT and CB1-/-CB2-/- mice demonstrated a critical involvement of CB1 and/or CB2 expressed
on APC in reducing the magnitude of immune response. The investigation was then extended to
characterize which APC populations in vivo were affected by the loss of CB1 and CB2 in the
anti-influenza response and to determine the effect of Δ9-THC on the maturation and recruitment
of APC into the lung. Since the discovery of cannabinoids and their receptors CB1 and CB2,
their effects on the immune system in health and disease have remained elusive. In part, it has
been difficult to determine which immune cell types are regulated by CB1 and CB2 signaling

164

and the ensuing consequences on cell function. In this study, an influenza model was used to
stimulate the immune system and the inflammatory response 3 dpi was investigated after cotreatment with the non-selective CB1 and CB2 agonist, Δ9-THC, in WT and CB1-/-CB2-/- mice.
Lower inflammatory responses were observed in WT as compared to CB1-/-CB2-/- mice with
Δ9-THC treatment as evidenced by reduced cytokine secretion by CD4+ T cells and NK cells,
lower percentages of APC populations in the lung, and decreased expression of genes associated
with the immune response to influenza as previously demonstrated. Furthermore, Δ9-THC
reduced T cell responses through modulation of DC function in WT but not in CB1-/-CB2-/mice. Thus, although other targets for Δ9-THC exist, CB1 and/or CB2 expressed on cDC are
important in mediating the immunomodulatory effects of Δ9-THC on effector cell populations
involved in the immune response to influenza virus infection.

A) Contributions of CB1 and CB2 to Cannabinoid Modulation of APC
It has been observed that both CB1 and CB2 contribute to the modulation of DC function
by Δ9-THC with the use of CB1- and CB2–specific antagonists [265] and that CB1 specifically
diminishes K+ outward currents through Kv channels to suppress DC function [266].
Furthermore, suppression of DC function by Δ9-THC might, in part, be independent of the Gprotein coupled receptors as evidenced by Δ9-THC-mediated suppression of maturation after in
bmDC generated from CB1-/-CB2-/- mice. In support of this argument, pertussis toxin, which
blocks the Gαi subunit necessary for CB1 and CB2 signaling [267, 268], did not completely
abrogate the suppression of IL-12p40 production in bmDC [269]. Also selective antagonists to
either CB1 and CB2 were not able to block the suppression of IL-12p40 by Δ9-THC [270]. This
suggests that although CB1 and CB2 are involved in Δ9-THC modulation of DC, this is not the

165

exclusive mechanism. In the present study, Δ9-THC suppressed LPS-induced maturation of
bmDC generated from CB1-/-CB2-/- mice, however; this suppression of maturation did not alter
elicitation of T cell responses in co-cultures. Collectively, these results suggest that the
maturation level, although suppressed, was sufficient in CB1-/-CB2-/- bmDC to elicit T cell
responses. Alternately, other factors aside from the maturation markers assessed on bmDC might
be involved in the suppression of maturation independently of CB1 and CB2. In support of this
argument, R848 did not induce maturation to the same extent as LPS in bmDC, yet R848-treated
DC were able to elicit T cells in co-cultures. Moreover, cytokine production in lymphocytes was
enhanced in vivo as a result of Δ9-THC treatment in CB1-/-CB2-/- further supporting the view
that targets, in addition to CB1 and CB2, other receptors might contribute to the Δ9-THCmediated effects on DC. Thus although CB1 and/or CB2 appear to play critical roles in the Δ9THC-mediated suppression of DC maturation and impairment of their ability to elicit T cell
responses, additional mechanisms are might be involved in the immue modulation of DC
function by Δ9-THC.
Among the APC populations in the lung, greater sensitivity to Δ9-THC-mediated
immunosuppression was observed in the reduced presence of cDC, pDC, and AM in the lung
after influenza infection. Migration of these three APC populations is regulated by distinct
chemokines and receptors [377-379]. Δ9-THC reduced steady-state mRNA levels of chemokine
receptors but not of chemokines. Taken together with the reduced percentages of APC
populations in the lung, the broad mRNA level changes might be a reflective of a reduced influx
of immune cells into the lung.

B) Putative Mechanisms by Which Δ9-THC Might Alter APC Function

166

While not as clear in AM, it has been demonstrated that cDC and pDC migration is
influenced by their maturation status [329-331, 378]. Aside from MHC II in pDC no overt
changes in the maturation status of the DC subsets was observed as a result of Δ9-THC
treatment. However, broad transcriptional suppression of influenza-induced genes associated
with immune response by Δ9-THC was observed in RNA isolated from the lung. Especially
proinflammatory cytokines are involved in the maturation of cDC, pDC, and AM and their
attraction to the site of infection [380-383]. Thus, indirect effects of reduced tissue inflammation
may be involved in the suppression of maturation and maturation-induced migration. In vitro,
Δ9-THC treatment impaired the LPS-induced maturation of DC as evidenced by lower
expression of CD86, for example. Taken together, it is plausible that reduced maturation of DC
by Δ9-THC, as demonstrated in vitro, decreases the ability of DC to migrate in vivo, which could
interfere with trafficking to or persisting in the lung, thus enriching for a DC population with a
mature phenotype. Furthermore, leukocyte trafficking, to a large extent, is dependent on
chemokine receptors which are part of the G-protein coupled receptor (GPCR) family [384]. It
has been shown that Δ9-THC and other cannabinoids alter the migration of multiple immune
cells. Although it is not clear under which conditions migration is augmented or impaired;
however, involvement of phosphatidylinositol 3-kinase/Akt and ERK1/2 has been demonstrated
[294, 385-388]. Moreover, crosstalk between GPCRs exists [389], and a role for Gαi has been
suggested in CB2 signaling which in turn inhibited signaling by other GPCRs [390]. Thus,
results in this investigation can be explained, in part, by a mechanism involving CB2-mediated
diminution of signaling through chemokine receptors. However, the intracellular signaling
pathways contributing to altered leukocyte trafficking by Δ9-THC remain to be elucidated and
the putative involvement of CB2 at this time is therefore speculative.

167

In contrast, it has been proposed that ligation of CB1 and CB2 induces apoptosis in DC,
which could account for the lower percentage of DC observed in the lung [265]. In the present
study, differences in bmDC viability in vitro using the LIVE/DEAD stain were not observed
(data not shown). It is noteworthy that there were several differences in the experimental
conditions in our study and that used by Do et. al.. Differences include higher LPS
concentrations (10 µg/mL), extended preincubation periods with Δ9-THC (2 hr preincubation),
sequential instead of concurrent treatment with LPS and Δ9-THC, and use of serum free media
[265], which by itself is a stressor to immune cells in culture [391].
It was previously observed that the synthetic CB1 and CB2 agonist, R(+)WIN55,212-2,
suppressed proinflammatory cytokine production in a multiple sclerosis model [392] and other
studies suggest that the mechanism by which cannabinoids suppress proinflammatory cytokine
production might involve IRF3-dependent upregulation of IFNβ [393]. Both IRF3 activation and
IFNβ and other type I interferon production are activated by influenza’s innate signaling in
separate pathways from pro-inflammatory cytokines [394-396]. While there were beneficial
effects observed with R(+)WIN55,212-2 in an experimental autoimmune encephalomyelitis
model, which mimics some aspects of multiple sclerosis by upregulation of type I IFN
production, one could hypothesize that Δ9-THC produces similar events in the influenza model
[393]. Although low in number, pDC greatly contribute to the inflammatory milieu and are
involved in the early phases of the anti-influenza immune response and are known to produce
large amounts of type I IFNs [397-399]. Indeed, IFNβ has been shown to suppress Th1
differentiation by DC when present prior to maturation [400, 401]. Furthermore, DC migration
was reduced by IFNβ [402] and the differentiation of naïve CD4+ T cells into the Th1 phenotype
was also suppressed [403, 404]. However, these studies suggest that pDC-produced IFNβ may be

168

beneficial in ameliorating inflammation. In this influenza model, pDC are one of the main
producers of type I IFNs including IFNβ, and Δ9-THC suppressed their presence and maturation.
Furthemore, steady-state mRNA levels of type 1 IFNs were not enhanced by Δ9-THC treatment
(Figure 36). Thus it is unlikely that in the influenza model IFNβ plays a role in the reduction in
proinflammatory cytokines.

C) Concluding Remarks on the Role of CB1 and CB2 on APC
Collectively, the present studies demonstrate that Δ9-THC suppressed the host immune
response against influenza virus challenge. Furthermore, after influenza infection, cDC, pDC,
and AM are sensitive to Δ9-THC-mediated immune suppression, which requires the presence of
CB1 and/or CB2. In vitro it was demonstrated that cDC-like bmDC play a critical role in
suppressing T cell responses after treatment with Δ9-THC, which was also found to be
dependent on CB1 and/or CB2. In conclusion, signaling through CB1 and CB2 might present an
avenue for therapeutic intervention in order to specifically attenuate APC function.

I.

Significance, Relevance, and Future Studies

The work presented in this dissertation makes a significant contribution to our understanding
of the role of CB1 and CB2 in the immune system. By characterizing the immune response in
WT and CB1-/-CB2-/- mice after infection with the pathogen influenza in vivo and using
immune cells isolated from these mice in vitro, the role of CB1 and CB2 in regulating immune
function was further elucidated.

169

Figure 37 Δ9-THC Does Not Alter Steady-state Whole Lung mRNA Levels of Type I
IFNs. RNA was isolated from whole lungs and reverse transcribed into cDNA (n=5). IFNα
and IFNβ mRNA levels were assessed with respect to the WT-CO-SAL samples, set to 1 fold.
Factorial ANOVA was used to analyze fold mRNA change: +++ (p≤0.001) difference
between NA (SAL) and PR8; ### (p≤0.001) difference between WT and CB1-/-CB2-/-.

170

Figure 37. Δ9-THC Does Not Alter Steady-state Whole
Lung mRNA Levels of Type I IFNs.

171

Since CD8+ T cells express relatively little transcript for CB1 and CB2, it was not surprising that
the effects mediated by Δ9-THC were independent of CB1 and/or CB2, since non-CB1/CB2mediated effects on T cells had been previously reported by this laboratory [231, 234, 284]. In
addition, CD8+ T cells from CB1-/-CB2-/- mice upregulated the surface expression of CD69 in
the presence of Δ9-THC alone, while CD8+ T cells from WT did not. It had been argued that
there might be compensatory mechanisms in CB1-/-CB2-/- mice as evidenced by upregulation of
the non-deleted portion of the CB2 gene transcript [334]. It is unknown what receptor(s), if any,
caused the Δ9-THC-mediated increase in CD69 expression in the absence of stimulation. Studies
identifying this unknown target and defining the transcriptional regulation of CB1 and CB2 will
provide further information on the mechanisms by which cannabinoids alter immune function.
Furthermore, the apparent increase in CD69 by Δ9-THC was inversely correlated with the
magnitude of CTL activity, suggesting that signaling pathways leading to an increase in CD69
expression and suppression of CTL function might share a common mechanism of regulation.
A second novel finding is that the deletion of CB1 and CB2 mainly affects APC function,
suggesting that the modulation of T lymphocyte function by cannabinoids is independent of CB1
and CB2 as described for CTL above. Although there were differences in the percent of
cytokine-producing lymphocytes in vivo, in vitro studies showed no difference in T cell cytokine
production between CB1-/-CB2-/- and WT mice, suggesting involvement of APC populations,
which elicit T cells. As a logical consequence, subsequent studies focused on APC populations
such as bmDC and AM. In the presence and absence of TLR stimulation the magnitude of
maturation in bmDC and AM CB1-/-CB2-/- mice exceeded the maturation observed in cells
from WT mice under the same conditions. In addition, the ability of bmDC to elicit T cells was
greater in bmDC from CB1-/-CB2-/- and did not require prior stimulation with a TLR ligand.

172

These results lead to the conclusion that not only the magnitude of response, but also the
maintenance of an unresponsive state of APC is governed by CB1 and/or CB2.
A third unique contribution of these studies is the finding of dysregulated DC responses in
the absence of immune stimulation in CB1-/-CB2-/- compared to WT mice after GMCSF
expansion of DC precursors in vitro. This was evidenced by greater levels of maturation markers
(e.g. MHC II) and the ability to elicit antigen-specific T cell responses. These results suggest
CB1 and/or CB2 play an important role in maintaining DC homeostasis, potentially by providing
a mechanism for self-regulation by endocannabinoids generated during the in vitro culture. The
endocannabinoids likely signal in an autocrine or paracrine manner since GMCSF-expanded DC
cultures are over 95% pure after 9 days. Cannabinoids might be one of many ways to fine tune
DC homeostasis and provide negative feedback to reduce “leaky” transcription [405] of
proinflammatory transcripts and other genes involved in DC homeostasis. While this idea is
speculative, reduction of “leaky” transcription is thought to occur as a result of cAMP response
element binding protein 2 activity, for example [406]. Furthermore, in the absence of negative
feedback provided by CB1 and/or CB2 in DC, autocrine-amplifying loops might result in
“leaky” transcription [405] leading to upregulation of CD86 and MHC II and expression of
cytokines, eventually presenting itself as a mature phenotype. Although speculative, these
autocrine-amplifying loops of soluble mediator release have been observed for complement
protein 3 in keratinocytes before [407].
Despite these novel findings of DC immune regulation by CB1 and/or CB2, there is a data
gap in the role of each receptor and its individual contribution to the phenotype observed.
Furthermore, it is unknown whether there are existing differences in GMCSF receptor
(GMCSFR) expression or the differences in the responses elicited as a result of GMCSFR

173

ligation between WT and CB1-/-CB2-/- DC precursors. In addition, there might be differences in
the number and/or development of the DC precursors in the bone marrow of WT and CB1-/CB2-/- mice. Although these studies are beyond the scope of this dissertation, another researcher
might find interest in pursuing these experiments.
Among the APC populations, pDC, and AM were most sensitive to the loss of CB1 and CB2.
Both cell types have a similar integrin profile defined by CD11c+CD11b- and consistently
bmDC generated from CB1-/-CB2-/- showed greater CD11c levels than bmDC generated from
WT (data not shown). Loss of integrin regulation including CD11c was implicated in adverse
immune outcomes in mice, suggesting that loss of CB1 and CB2 results in a multi-faceted
dysregulation in the phenotype of APC, most drastically in DC subpopulations including cDC
and pDC [408]. Potentially, antagonists to CB1 and/or CB2 could be used to temporarily induce
the generation and exacerbate the maturation of pDC and cDC populationsfor a therapeutic
benefit. For example, in HIV positive patients an increase in the number or maturation of cDC
populations might allow for the elicitation of HIV-specific CTL populations without the
requirement of CD4+ T cell help. Moreover, correcting the loss of pDC populations as observed
during HIV infection with antagonists to CB1 and/or CB2 might slow or halt disease progression
in HIV positive patients as regulating the balance of pDC has been implicated in the successful
control of HIV infection [409].
Since the dysregulation of APC immune function in CB1-/-CB2-/- compared to WT was
found in vivo and in vitro, it is less likely that the cells other than those of the immune system
significantly contribute to the immune exacerbation to influenza infection in vivo. However, to
test this, bone-marrow chimeras using adoptive transfer of CB1-/-CB2-/- bone marrow into WT
recipients and WT bone marrow into WT recipients would have to be generated. Differences in

174

immune reactivity between CB1-/-CB2-/- and WT bone marrow transfers would confirm that the
immune exacerbation can be attributed to the immune cells of CB1-/-CB2-/- mice. Furthermore,
using adoptive transfers of CB1-/-CB2-/- bmDC into WT donors and WT bmDC into WT donors
in a T cell activation or influenza challenge model would determine whether these cDC-like cells
are sufficient in inducing exacerbated immune responses. The described experiments are
technically challenging and the adoptive transfer of bmDC was attempted, however, the data
were equivocal as little to no T cell activation was observed and are not presented in this
dissertation.
The prevalence of marijuana consumption in industrialized countries is a reason for concern
regarding the findings presented in this dissertation. As a result of globalization the spread of
influenza virus is very rapid and poses difficulties to national governments in preparing for a
pandemic or epidemic. In years of accurate antigen prediction, annual vaccination to influenza
virus is the most effective method in halting viral spread and limiting its mortality. The concept
of herd immunity describes the idea that the vaccination of a significant portion of the population
prevents viral spread within the entire population (including non-vaccinated), because vaccinated
individuals do not transmit the pathogen. In a murine influenza challenge model the immune
response magnitude was drastically reduced by Δ9-THC treatment. These results suggest that
vaccinations might be rendered less effective in marijuana users and could thereby reduce herd
immunity to influenza hence leading to reduced protection of a greater population from a viral
pandemic or epidemic [410, 411].
The dysregulation of DC responses as a result of the loss of CB1 and CB2 are particularly
alarming considering the important role that DC play in the orchestration of immune responses.
DC can induce and maintain tolerance, as well as contribute to the elicitation of effector immune

175

cells. While DC elicited immune responses are important in producing successful vaccinations
and protection from pathogens, unwanted DC immune responses can lead to autoimmune disease
[96, 376, 412-414]. Due to the identification of SNPs in CB1 and CB2 in humans, the potential
for reduced ability to regulate the magnitude of immune response by endocannabinoids exists
potentially resulting in an exacerbated immune phenotype and manifesting itself as autoimmune
disease. Given the association of SNPs in CB1 and CB2 with autoimmune disorders it is likely
that DC can play a role in the development of the phenotypes of autoimmune disease [318, 319].
The system and cells isolated from humans with SNPs in either or both CB1 and CB2 need to be
further characterized to determine a human correlate between the findings presented in this
dissertation and human health. The concepts described in this section are summarized in Figure
37.

176

Figure 38. DC Cannabinoid Immune Modulation Model. For interpretation of the references to color in this and
all other figures, the reader is referred to the electronic version of this dissertation.

177

REFERENCES

178

1.

Vander, A.J., J.H. Sherman, and D.S. Luciano, Respiration, in Human Physiology: The
Mechanism of Body Function. 2001, McGraw−Hill: New York.

2.

Lusuardi, M., et al., Lung mucosal immunity: immunoglobulin-A revisited. Eur Respir J,
2002. 19(4): p. 785; author reply 785-6.

3.

Holmgren, J. and C. Czerkinsky, Mucosal immunity and vaccines. Nat Med, 2005. 11(4
Suppl): p. S45-53.

4.

Bessa, J., et al., Efficient induction of mucosal and systemic immune responses by viruslike particles administered intranasally: implications for vaccine design. Eur J Immunol,
2008. 38(1): p. 114-26.

5.

Gomes, R.F., et al., Comparative respiratory system mechanics in rodents. J Appl
Physiol, 2000. 89(3): p. 908-16.

6.

Harkema, J.R., Comparative aspects of nasal airway anatomy: relevance to inhalation
toxicology. Toxicol Pathol, 1991. 19(4 Pt 1): p. 321-36.

7.

Hautamaki, R.D., et al., Requirement for macrophage elastase for cigarette smokeinduced emphysema in mice. Science, 1997. 277(5334): p. 2002-4.

8.

Reznik, G.K., Comparative anatomy, physiology, and function of the upper respiratory
tract. Environ Health Perspect, 1990. 85: p. 171-6.

9.

Miller, M.J., et al., Oral breathing in newborn infants. J Pediatr, 1985. 107(3): p. 465-9.

10.

Morgan, D.L., et al., Respiratory toxicity of diacetyl in C57BL/6 mice. Toxicol Sci, 2008.
103(1): p. 169-80.

11.

Boers, J.E., A.W. Ambergen, and F.B. Thunnissen, Number and proliferation of clara
cells in normal human airway epithelium. Am J Respir Crit Care Med, 1999. 159(5 Pt 1):
p. 1585-91.

12.

Plopper, C.G., L.H. Hill, and A.T. Mariassy, Ultrastructure of the nonciliated
bronchiolar epithelial (Clara) cell of mammalian lung. III. A study of man with
comparison of 15 mammalian species. Exp Lung Res, 1980. 1(2): p. 171-80.

179

13.

Plopper, C.G., et al., Comparison of nonciliated tracheal epithelial cells in six
mammalian species: ultrastructure and population densities. Exp Lung Res, 1983. 5(4):
p. 281-94.

14.

Holt, P.G., et al., Regulation of immunological homeostasis in the respiratory tract. Nat
Rev Immunol, 2008. 8(2): p. 142-52.

15.

Carlson, D., et al., Oropharyngeal flora in healthy infants: observations and implications
for cystic fibrosis care. Pediatr Pulmonol, 2009. 44(5): p. 497-502.

16.

Konno, M., et al., Study of upper respiratory tract bacterial flora: first report. Variations
in upper respiratory tract bacterial flora in patients with acute upper respiratory tract
infection and healthy subjects and variations by subject age. J Infect Chemother, 2006.
12(2): p. 83-96.

17.

Hilty, M., et al., Disordered microbial communities in asthmatic airways. PLoS One,
2010. 5(1): p. e8578.

18.

Wiley, J.A., M.P. Tighe, and A.G. Harmsen, Upper respiratory tract resistance to
influenza infection is not prevented by the absence of either nasal-associated lymphoid
tissue or cervical lymph nodes. J Immunol, 2005. 175(5): p. 3186-96.

19.

Randall, T.D., Bronchus-associated lymphoid tissue (BALT) structure and function. Adv
Immunol, 2010. 107: p. 187-241.

20.

Sminia, T., G.J. van der Brugge-Gamelkoorn, and S.H. Jeurissen, Structure and function
of bronchus-associated lymphoid tissue (BALT). Crit Rev Immunol, 1989. 9(2): p. 11950.

21.

Wu, H.Y., H.H. Nguyen, and M.W. Russell, Nasal lymphoid tissue (NALT) as a mucosal
immune inductive site. Scand J Immunol, 1997. 46(5): p. 506-13.

22.

Kiyono, H. and S. Fukuyama, NALT- versus Peyer's-patch-mediated mucosal immunity.
Nat Rev Immunol, 2004. 4(9): p. 699-710.

23.

Xu, B., et al., Lymphocyte homing to bronchus-associated lymphoid tissue (BALT) is
mediated by L-selectin/PNAd, alpha4beta1 integrin/VCAM-1, and LFA-1 adhesion
pathways. J Exp Med, 2003. 197(10): p. 1255-67.

180

24.

Haslett, C., Lung Defenses and Immunology. Crofton and Douglas's Respiratory
Diseases. 2000: Blackwell Science Ltd. 83-90.

25.

Canning, B.J., Anatomy and neurophysiology of the cough reflex: ACCP evidence-based
clinical practice guidelines. Chest, 2006. 129(1 Suppl): p. 33S-47S.

26.

Vareille, M., et al., The airway epithelium: soldier in the fight against respiratory
viruses. Clin Microbiol Rev, 2011. 24(1): p. 210-29.

27.

Gerrity, T.R., et al., Calculated deposition of inhaled particles in the airway generations
of normal subjects. J Appl Physiol, 1979. 47(4): p. 867-73.

28.

Schlesinger, R.B., Comparative deposition of inhaled aerosols in experimental animals
and humans: a review. J Toxicol Environ Health, 1985. 15(2): p. 197-214.

29.

Ganz, T., Antimicrobial polypeptides in host defense of the respiratory tract. J Clin
Invest, 2002. 109(6): p. 693-7.

30.

Stripp, B.R., et al., Clara cell secretory protein deficiency alters clara cell secretory
apparatus and the protein composition of airway lining fluid. Am J Respir Cell Mol Biol,
2002. 27(2): p. 170-8.

31.

Devereux, T.R., Alveolar type II and Clara cells: isolation and xenobiotic metabolism.
Environ Health Perspect, 1984. 56: p. 95-101.

32.

Williams, M.C., Alveolar type I cells: molecular phenotype and development. Annu Rev
Physiol, 2003. 65: p. 669-95.

33.

Johnson, M.D., et al., Alveolar epithelial type I cells contain transport proteins and
transport sodium, supporting an active role for type I cells in regulation of lung liquid
homeostasis. Proc Natl Acad Sci U S A, 2002. 99(4): p. 1966-71.

34.

Fehrenbach, H., Alveolar epithelial type II cell: defender of the alveolus revisited. Respir
Res, 2001. 2(1): p. 33-46.

35.

Castranova, V., et al., The alveolar type II epithelial cell: a multifunctional pneumocyte.
Toxicol Appl Pharmacol, 1988. 93(3): p. 472-83.

181

36.

McCormack, F.X. and J.A. Whitsett, The pulmonary collectins, SP-A and SP-D,
orchestrate innate immunity in the lung. J Clin Invest, 2002. 109(6): p. 707-12.

37.

Bals, R., et al., The peptide antibiotic LL-37/hCAP-18 is expressed in epithelia of the
human lung where it has broad antimicrobial activity at the airway surface. Proc Natl
Acad Sci U S A, 1998. 95(16): p. 9541-6.

38.

Schutte, B.C. and P.B. McCray, Jr., [beta]-defensins in lung host defense. Annu Rev
Physiol, 2002. 64: p. 709-48.

39.

Hiemstra, P.S., The role of epithelial beta-defensins and cathelicidins in host defense of
the lung. Exp Lung Res, 2007. 33(10): p. 537-42.

40.

Suzuki, T., C.W. Chow, and G.P. Downey, Role of innate immune cells and their
products in lung immunopathology. Int J Biochem Cell Biol, 2008. 40(6-7): p. 1348-61.

41.

Actor, J.K., S.A. Hwang, and M.L. Kruzel, Lactoferrin as a natural immune modulator.
Curr Pharm Des, 2009. 15(17): p. 1956-73.

42.

Akinbi, H.T., et al., Bacterial killing is enhanced by expression of lysozyme in the lungs
of transgenic mice. J Immunol, 2000. 165(10): p. 5760-6.

43.

Dajani, R., et al., Lysozyme secretion by submucosal glands protects the airway from
bacterial infection. Am J Respir Cell Mol Biol, 2005. 32(6): p. 548-52.

44.

Opitz, B., et al., Innate immune recognition in infectious and noninfectious diseases of
the lung. Am J Respir Crit Care Med, 2010. 181(12): p. 1294-309.

45.

Medzhitov, R., Toll-like receptors and innate immunity. Nat Rev Immunol, 2001. 1(2): p.
135-45.

46.

Takeda, K., T. Kaisho, and S. Akira, Toll-like receptors. Annu Rev Immunol, 2003. 21:
p. 335-76.

47.

Mayer, A.K., et al., Differential recognition of TLR-dependent microbial ligands in
human bronchial epithelial cells. J Immunol, 2007. 178(5): p. 3134-42.

182

48.

Sha, Q., et al., Activation of airway epithelial cells by toll-like receptor agonists. Am J
Respir Cell Mol Biol, 2004. 31(3): p. 358-64.

49.

Schmeck, B., et al., Pneumococci induced TLR- and Rac1-dependent NF-kappaBrecruitment to the IL-8 promoter in lung epithelial cells. Am J Physiol Lung Cell Mol
Physiol, 2006. 290(4): p. L730-L737.

50.

Beg, A.A., Endogenous ligands of Toll-like receptors: implications for regulating
inflammatory and immune responses. Trends Immunol, 2002. 23(11): p. 509-12.

51.

Tsan, M.F. and B. Gao, Endogenous ligands of Toll-like receptors. J Leukoc Biol, 2004.
76(3): p. 514-9.

52.

Kariko, K., et al., mRNA is an endogenous ligand for Toll-like receptor 3. J Biol Chem,
2004. 279(13): p. 12542-50.

53.

Ohashi, K., et al., Cutting edge: heat shock protein 60 is a putative endogenous ligand of
the toll-like receptor-4 complex. J Immunol, 2000. 164(2): p. 558-61.

54.

Muruve, D.A., et al., The inflammasome recognizes cytosolic microbial and host DNA
and triggers an innate immune response. Nature, 2008. 452(7183): p. 103-7.

55.

Takaoka, A. and T. Taniguchi, Cytosolic DNA recognition for triggering innate immune
responses. Adv Drug Deliv Rev, 2008. 60(7): p. 847-57.

56.

Yoneyama, M., et al., The RNA helicase RIG-I has an essential function in doublestranded RNA-induced innate antiviral responses. Nat Immunol, 2004. 5(7): p. 730-7.

57.

Guarda, G., et al., Differential expression of NLRP3 among hematopoietic cells. J
Immunol, 2011. 186(4): p. 2529-34.

58.

Allen, I.C., et al., The NLRP3 inflammasome mediates in vivo innate immunity to
influenza A virus through recognition of viral RNA. Immunity, 2009. 30(4): p. 556-65.

59.

Guo, Z., et al., NS1 protein of influenza A virus inhibits the function of intracytoplasmic
pathogen sensor, RIG-I. Am J Respir Cell Mol Biol, 2007. 36(3): p. 263-9.

183

60.

Kato, H., et al., Cell type-specific involvement of RIG-I in antiviral response. Immunity,
2005. 23(1): p. 19-28.

61.

Kato, H., et al., Differential roles of MDA5 and RIG-I helicases in the recognition of RNA
viruses. Nature, 2006. 441(7089): p. 101-5.

62.

Gitlin, L., et al., Essential role of mda-5 in type I IFN responses to
polyriboinosinic:polyribocytidylic acid and encephalomyocarditis picornavirus. Proc
Natl Acad Sci U S A, 2006. 103(22): p. 8459-64.

63.

Berghall, H., et al., The interferon-inducible RNA helicase, mda-5, is involved in measles
virus-induced expression of antiviral cytokines. Microbes Infect, 2006. 8(8): p. 2138-44.

64.

Kang, D.C., et al., mda-5: An interferon-inducible putative RNA helicase with doublestranded RNA-dependent ATPase activity and melanoma growth-suppressive properties.
Proc Natl Acad Sci U S A, 2002. 99(2): p. 637-42.

65.

Groskreutz, D.J., et al., Respiratory syncytial virus induces TLR3 protein and protein
kinase R, leading to increased double-stranded RNA responsiveness in airway epithelial
cells. J Immunol, 2006. 176(3): p. 1733-40.

66.

Nakamura, T., et al., Double-stranded RNA-dependent protein kinase links pathogen
sensing with stress and metabolic homeostasis. Cell, 2010. 140(3): p. 338-48.

67.

Ichinohe, T., Respective roles of TLR, RIG-I and NLRP3 in influenza virus infection and
immunity: impact on vaccine design. Expert Rev Vaccines, 2010. 9(11): p. 1315-24.

68.

Siren, J., et al., Retinoic acid inducible gene-I and mda-5 are involved in influenza A
virus-induced expression of antiviral cytokines. Microbes Infect, 2006. 8(8): p. 2013-20.

69.

Tschopp, J. and K. Schroder, NLRP3 inflammasome activation: The convergence of
multiple signalling pathways on ROS production? Nat Rev Immunol, 2010. 10(3): p.
210-5.

70.

Iyer, S.S., et al., Necrotic cells trigger a sterile inflammatory response through the Nlrp3
inflammasome. Proc Natl Acad Sci U S A, 2009. 106(48): p. 20388-93.

184

71.

Schroder, K., R. Zhou, and J. Tschopp, The NLRP3 inflammasome: a sensor for
metabolic danger? Science, 2010. 327(5963): p. 296-300.

72.

Baccala, R., et al., TLR-dependent and TLR-independent pathways of type I interferon
induction in systemic autoimmunity. Nat Med, 2007. 13(5): p. 543-51.

73.

Moynagh, P.N., TLR signalling and activation of IRFs: revisiting old friends from the
NF-kappaB pathway. Trends Immunol, 2005. 26(9): p. 469-76.

74.

Takeda, K. and S. Akira, Toll-like receptors in innate immunity. Int Immunol, 2005.
17(1): p. 1-14.

75.

Horng, T., G.M. Barton, and R. Medzhitov, TIRAP: an adapter molecule in the Toll
signaling pathway. Nat Immunol, 2001. 2(9): p. 835-41.

76.

Sato, S., et al., Toll/IL-1 receptor domain-containing adaptor inducing IFN-beta (TRIF)
associates with TNF receptor-associated factor 6 and TANK-binding kinase 1, and
activates two distinct transcription factors, NF-kappa B and IFN-regulatory factor-3, in
the Toll-like receptor signaling. J Immunol, 2003. 171(8): p. 4304-10.

77.

Petrilli, V., et al., The inflammasome: a danger sensing complex triggering innate
immunity. Curr Opin Immunol, 2007. 19(6): p. 615-22.

78.

Bauernfeind, F.G., et al., Cutting edge: NF-kappaB activating pattern recognition and
cytokine receptors license NLRP3 inflammasome activation by regulating NLRP3
expression. J Immunol, 2009. 183(2): p. 787-91.

79.

Martinon, F., K. Burns, and J. Tschopp, The inflammasome: a molecular platform
triggering activation of inflammatory caspases and processing of proIL-beta. Mol Cell,
2002. 10(2): p. 417-26.

80.

Wen, H., et al., Fatty acid-induced NLRP3-ASC inflammasome activation interferes with
insulin signaling. Nat Immunol, 2011. 12(5): p. 408-15.

81.

Kanneganti, T.D., M. Lamkanfi, and G. Nunez, Intracellular NOD-like receptors in host
defense and disease. Immunity, 2007. 27(4): p. 549-59.

185

82.

Dostert, C., et al., Innate immune activation through Nalp3 inflammasome sensing of
asbestos and silica. Science, 2008. 320(5876): p. 674-7.

83.

Vandanmagsar, B., et al., The NLRP3 inflammasome instigates obesity-induced
inflammation and insulin resistance. Nat Med, 2010. 17(2): p. 179-88.

84.

Thomas, P.G., et al., The intracellular sensor NLRP3 mediates key innate and healing
responses to influenza A virus via the regulation of caspase-1. Immunity, 2009. 30(4): p.
566-75.

85.

Masters, S.L. and L.A. O'Neill, Disease-associated amyloid and misfolded protein
aggregates activate the inflammasome. Trends Mol Med, 2011. 17(5): p. 276-82.

86.

Masters, S.L., et al., Activation of the NLRP3 inflammasome by islet amyloid polypeptide
provides a mechanism for enhanced IL-1beta in type 2 diabetes. Nat Immunol, 2011.
11(10): p. 897-904.

87.

Kalali, B.N., et al., Double-stranded RNA induces an antiviral defense status in
epidermal keratinocytes through TLR3-, PKR-, and MDA5/RIG-I-mediated differential
signaling. J Immunol, 2008. 181(4): p. 2694-704.

88.

Schleimer, R.P., et al., Epithelium: at the interface of innate and adaptive immune
responses. J Allergy Clin Immunol, 2007. 120(6): p. 1279-84.

89.

Cleret, A., et al., Lung dendritic cells rapidly mediate anthrax spore entry through the
pulmonary route. J Immunol, 2007. 178(12): p. 7994-8001.

90.

Jakubzick, C., et al., Blood monocyte subsets differentially give rise to CD103+ and
CD103- pulmonary dendritic cell populations. J Immunol, 2008. 180(5): p. 3019-27.

91.

Osterholzer, J.J., et al., Role of dendritic cells and alveolar macrophages in regulating
early host defense against pulmonary infection with Cryptococcus neoformans. Infect
Immun, 2009. 77(9): p. 3749-58.

92.

Lambrecht, B.N., Alveolar macrophage in the driver's seat. Immunity, 2006. 24(4): p.
366-8.

186

93.

Power, C.A., et al., Cloning and characterization of a specific receptor for the novel CC
chemokine MIP-3alpha from lung dendritic cells. J Exp Med, 1997. 186(6): p. 825-35.

94.

Wareing, M.D., et al., Chemokine expression during the development and resolution of a
pulmonary leukocyte response to influenza A virus infection in mice. J Leukoc Biol, 2004.
76(4): p. 886-95.

95.

Saenz, S.A., B.C. Taylor, and D. Artis, Welcome to the neighborhood: epithelial cellderived cytokines license innate and adaptive immune responses at mucosal sites.
Immunol Rev, 2008. 226: p. 172-90.

96.

Banchereau, J. and R.M. Steinman, Dendritic cells and the control of immunity. Nature,
1998. 392(6673): p. 245-52.

97.

Thompson, W.W., et al., Estimating influenza-associated deaths in the United States. Am
J Public Health, 2009. 99 Suppl 2: p. S225-30.

98.

Smith, N.M., et al., Prevention and Control of Influenza: recommendations of the
Advisory Committee on Immunization Practices (ACIP). MMWR Recomm Rep, 2006.
55(RR-10): p. 1-42.

99.

Molinari, N.A., et al., The annual impact of seasonal influenza in the US: measuring
disease burden and costs. Vaccine, 2007. 25(27): p. 5086-96.

100.

Taubenberger, J.K. and D.M. Morens, The pathology of influenza virus infections. Annu
Rev Pathol, 2008. 3: p. 499-522.

101.

Lamb, R.A. and R.M. Krug, Orthomyxoviridae: the viruses and their replication., in
Fields Virology. 2001, Lippincott Williams & Wilkins: Philadelphia. p. 1487–1531.

102.

Basler, C.F. and P.V. Aguilar, Progress in identifying virulence determinants of the 1918
H1N1 and the Southeast Asian H5N1 influenza A viruses. Antiviral Res, 2008. 79(3): p.
166-78.

103.

McGeoch, D., P. Fellner, and C. Newton, Influenza virus genome consists of eight
distinct RNA species. Proc Natl Acad Sci U S A, 1976. 73(9): p. 3045-9.

187

104.

Bouvier, N.M. and P. Palese, The biology of influenza viruses. Vaccine, 2008. 26 Suppl
4: p. D49-53.

105.

Konig, R., et al., Human host factors required for influenza virus replication. Nature,
2010. 463(7282): p. 813-7.

106.

Weis, W., et al., Structure of the influenza virus haemagglutinin complexed with its
receptor, sialic acid. Nature, 1988. 333(6172): p. 426-31.

107.

Rust, M.J., et al., Assembly of endocytic machinery around individual influenza viruses
during viral entry. Nat Struct Mol Biol, 2004. 11(6): p. 567-73.

108.

Mercer, J., M. Schelhaas, and A. Helenius, Virus entry by endocytosis. Annu Rev
Biochem, 2010. 79: p. 803-33.

109.

Chu, V.C. and G.R. Whittaker, Influenza virus entry and infection require host cell Nlinked glycoprotein. Proc Natl Acad Sci U S A, 2004. 101(52): p. 18153-8.

110.

Cros, J.F. and P. Palese, Trafficking of viral genomic RNA into and out of the nucleus:
influenza, Thogoto and Borna disease viruses. Virus Res, 2003. 95(1-2): p. 3-12.

111.

Palese, P., et al., Characterization of temperature sensitive influenza virus mutants
defective in neuraminidase. Virology, 1974. 61(2): p. 397-410.

112.

Colman, P.M., J.N. Varghese, and W.G. Laver, Structure of the catalytic and antigenic
sites in influenza virus neuraminidase. Nature, 1983. 303(5912): p. 41-4.

113.

Varghese, J.N., W.G. Laver, and P.M. Colman, Structure of the influenza virus
glycoprotein antigen neuraminidase at 2.9 A resolution. Nature, 1983. 303(5912): p. 3540.

114.

Drake, J.W., Rates of spontaneous mutation among RNA viruses. Proc Natl Acad Sci U S
A, 1993. 90(9): p. 4171-5.

115.

Carrat, F. and A. Flahault, Influenza vaccine: the challenge of antigenic drift. Vaccine,
2007. 25(39-40): p. 6852-62.

188

116.

Cottey, R., C. Rowe, and B. Bender, Influenza Virus, in Current Protocols in
Immunology, R. Coico, Editor. 2001, Wiley.

117.

Ibricevic, A., et al., Influenza virus receptor specificity and cell tropism in mouse and
human airway epithelial cells. J Virol, 2006. 80(15): p. 7469-80.

118.

Buchweitz, J.P., J.R. Harkema, and N.E. Kaminski, Time-dependent airway epithelial
and inflammatory cell responses induced by influenza virus A/PR/8/34 in C57BL/6 mice.
Toxicol Pathol, 2007. 35(3): p. 424-35.

119.

Buchweitz, J.P., et al., Modulation of airway responses to influenza A/PR/8/34 by Delta9tetrahydrocannabinol in C57BL/6 mice. J Pharmacol Exp Ther, 2007. 323(2): p. 675-83.

120.

Buchweitz, J.P., et al., Targeted deletion of cannabinoid receptors CB1 and CB2
produced enhanced inflammatory responses to influenza A/PR/8/34 in the absence and
presence of Delta9-tetrahydrocannabinol. J Leukoc Biol, 2008. 83(3): p. 785-96.

121.

Brown, D.M., E. Roman, and S.L. Swain, CD4 T cell responses to influenza infection.
Semin Immunol, 2004. 16(3): p. 171-7.

122.

Cambridge, G., J.S. Mackenzie, and D. Keast, Cell-mediated immune response to
influenza virus infections in mice. Infect Immun, 1976. 13(1): p. 36-43.

123.

Wijburg, O.L., et al., Alveolar macrophages regulate the induction of primary cytotoxic
T-lymphocyte responses during influenza virus infection. J Virol, 1997. 71(12): p. 9450-7.

124.

La Gruta, N.L., et al., A question of self-preservation: immunopathology in influenza
virus infection. Immunol Cell Biol, 2007. 85(2): p. 85-92.

125.

Legge, K.L. and T.J. Braciale, Accelerated migration of respiratory dendritic cells to the
regional lymph nodes is limited to the early phase of pulmonary infection. Immunity,
2003. 18(2): p. 265-77.

126.

Manicassamy, B., et al., Analysis of in vivo dynamics of influenza virus infection in mice
using a GFP reporter virus. Proc Natl Acad Sci U S A, 2010. 107(25): p. 11531-6.

189

127.

GeurtsvanKessel, C.H., et al., Clearance of influenza virus from the lung depends on
migratory langerin+CD11b- but not plasmacytoid dendritic cells. J Exp Med, 2008.
205(7): p. 1621-34.

128.

Pulendran, B., H. Tang, and T.L. Denning, Division of labor, plasticity, and crosstalk
between dendritic cell subsets. Curr Opin Immunol, 2008. 20(1): p. 61-7.

129.

Liu, Y.J., Dendritic cell subsets and lineages, and their functions in innate and adaptive
immunity. Cell, 2001. 106(3): p. 259-62.

130.

Shortman, K. and Y.J. Liu, Mouse and human dendritic cell subtypes. Nat Rev Immunol,
2002. 2(3): p. 151-61.

131.

Nakano, H., M. Yanagita, and M.D. Gunn, CD11c(+)B220(+)Gr-1(+) cells in mouse
lymph nodes and spleen display characteristics of plasmacytoid dendritic cells. J Exp
Med, 2001. 194(8): p. 1171-8.

132.

Wu, L. and Y.J. Liu, Development of dendritic-cell lineages. Immunity, 2007. 26(6): p.
741-50.

133.

Gonzalez-Juarrero, M., et al., Dynamics of macrophage cell populations during murine
pulmonary tuberculosis. J Immunol, 2003. 171(6): p. 3128-35.

134.

Diao, J., et al., Characterization of distinct conventional and plasmacytoid dendritic cellcommitted precursors in murine bone marrow. J Immunol, 2004. 173(3): p. 1826-33.

135.

Naik, S.H., et al., Development of plasmacytoid and conventional dendritic cell subtypes
from single precursor cells derived in vitro and in vivo. Nat Immunol, 2007. 8(11): p.
1217-26.

136.

Fonteneau, J.F., et al., Activation of influenza virus-specific CD4+ and CD8+ T cells: a
new role for plasmacytoid dendritic cells in adaptive immunity. Blood, 2003. 101(9): p.
3520-6.

137.

Kaisho, T., Molecular mechanisms for plasmacytoid dendritic cell function and
development. Vaccine, 2010. 28(50): p. 8046-7.

190

138.

Teixeira, L., et al., Plasmacytoid and conventional dendritic cells are early producers of
IL-12 in Neospora caninum-infected mice. Immunol Cell Biol, 2010. 88(1): p. 79-86.

139.

Grayson, M.H., et al., Controls for lung dendritic cell maturation and migration during
respiratory viral infection. J Immunol, 2007. 179(3): p. 1438-48.

140.

Di Cesare, A., P. Di Meglio, and F.O. Nestle, The IL-23/Th17 axis in the
immunopathogenesis of psoriasis. J Invest Dermatol, 2009. 129(6): p. 1339-50.

141.

Iwakura, Y. and H. Ishigame, The IL-23/IL-17 axis in inflammation. J Clin Invest, 2006.
116(5): p. 1218-22.

142.

Isaksson, M., et al., Plasmacytoid DC promote priming of autoimmune Th17 cells and
EAE. Eur J Immunol, 2009. 39(10): p. 2925-35.

143.

Kumagai, Y., et al., Alveolar macrophages are the primary interferon-alpha producer in
pulmonary infection with RNA viruses. Immunity, 2007. 27(2): p. 240-52.

144.

Pribul, P.K., et al., Alveolar macrophages are a major determinant of early responses to
viral lung infection but do not influence subsequent disease development. J Virol, 2008.
82(9): p. 4441-8.

145.

Kirby, A.C., M.C. Coles, and P.M. Kaye, Alveolar macrophages transport pathogens to
lung draining lymph nodes. J Immunol, 2009. 183(3): p. 1983-9.

146.

Thepen, T., G. Kraal, and P.G. Holt, The role of alveolar macrophages in regulation of
lung inflammation. Ann N Y Acad Sci, 1994. 725: p. 200-6.

147.

Lin, K.L., et al., CCR2+ monocyte-derived dendritic cells and exudate macrophages
produce influenza-induced pulmonary immune pathology and mortality. J Immunol,
2008. 180(4): p. 2562-72.

148.

De Santo, C., et al., Invariant NKT cells reduce the immunosuppressive activity of
influenza A virus-induced myeloid-derived suppressor cells in mice and humans. J Clin
Invest, 2008.

149.

Corzo, C.A., et al., Mechanism regulating reactive oxygen species in tumor-induced
myeloid-derived suppressor cells. J Immunol, 2009. 182(9): p. 5693-701.

191

150.

Gabrilovich, D.I. and S. Nagaraj, Myeloid-derived suppressor cells as regulators of the
immune system. Nat Rev Immunol, 2009. 9(3): p. 162-74.

151.

Fairweather, D. and D. Cihakova, Alternatively activated macrophages in infection and
autoimmunity. J Autoimmun, 2009. 33(3-4): p. 222-30.

152.

Pirhonen, J., S. Matikainen, and I. Julkunen, Regulation of virus-induced IL-12 and IL-23
expression in human macrophages. J Immunol, 2002. 169(10): p. 5673-8.

153.

Swain, S.L., R.W. Dutton, and D.L. Woodland, T cell responses to influenza virus
infection: effector and memory cells. Viral Immunol, 2004. 17(2): p. 197-209.

154.

Kim, T.S., J. Sun, and T.J. Braciale, T cell responses during influenza infection: getting
and keeping control. Trends Immunol, 2011. 32(5): p. 225-31.

155.

McKinstry, K.K., et al., IL-10 deficiency unleashes an influenza-specific Th17 response
and enhances survival against high-dose challenge. J Immunol, 2009. 182(12): p. 735363.

156.

Kolls, J.K. and A. Linden, Interleukin-17 family members and inflammation. Immunity,
2004. 21(4): p. 467-76.

157.

Korn, T., et al., IL-17 and Th17 Cells. Annu Rev Immunol, 2009. 27: p. 485-517.

158.

Ye, P., et al., Requirement of interleukin 17 receptor signaling for lung CXC chemokine
and granulocyte colony-stimulating factor expression, neutrophil recruitment, and host
defense. J Exp Med, 2001. 194(4): p. 519-27.

159.

Kohlmeier, J.E. and D.L. Woodland, Immunity to respiratory viruses. Annu Rev
Immunol, 2009. 27: p. 61-82.

160.

Strieter, R.M., J.A. Belperio, and M.P. Keane, Cytokines in innate host defense in the
lung. J Clin Invest, 2002. 109(6): p. 699-705.

161.

Crowe, C.R., et al., Critical role of IL-17RA in immunopathology of influenza infection. J
Immunol, 2009. 183(8): p. 5301-10.

192

162.

Wiley, J.A., et al., Production of interferon-gamma by influenza hemagglutinin-specific
CD8 effector T cells influences the development of pulmonary immunopathology. Am J
Pathol, 2001. 158(1): p. 119-30.

163.

Moskophidis, D. and D. Kioussis, Contribution of virus-specific CD8+ cytotoxic T cells
to virus clearance or pathologic manifestations of influenza virus infection in a T cell
receptor transgenic mouse model. J Exp Med, 1998. 188(2): p. 223-32.

164.

Sun, J., et al., Effector T cells control lung inflammation during acute influenza virus
infection by producing IL-10. Nat Med, 2009. 15(3): p. 277-84.

165.

Jenkins, M.R., et al., Heterogeneity of effector phenotype for acute phase and memory
influenza A virus-specific CTL. J Immunol, 2007. 179(1): p. 64-70.

166.

Zhong, W., et al., Genome-wide characterization of a viral cytotoxic T lymphocyte
epitope repertoire. J Biol Chem, 2003. 278(46): p. 45135-44.

167.

Hamada, H., et al., Tc17, a unique subset of CD8 T cells that can protect against lethal
influenza challenge. J Immunol, 2009. 182(6): p. 3469-81.

168.

Lauzon, N.M., et al., The direct effects of Toll-like receptor ligands on human NK cell
cytokine production and cytotoxicity. Cell Immunol, 2006. 241(2): p. 102-12.

169.

Fernandez, N.C., et al., Dendritic cells directly trigger NK cell functions: cross-talk
relevant in innate anti-tumor immune responses in vivo. Nat Med, 1999. 5(4): p. 405-11.

170.

Passos, S.T., et al., IL-6 promotes NK cell production of IL-17 during toxoplasmosis. J
Immunol, 2010. 184(4): p. 1776-83.

171.

Lanier, L.L., NK cell receptors. Annu Rev Immunol, 1998. 16: p. 359-93.

172.

Kos, F.J. and E.G. Engleman, Role of natural killer cells in the generation of influenza
virus-specific cytotoxic T cells. Cell Immunol, 1996. 173(1): p. 1-6.

173.

Jegerlehner, A., et al., Influenza A vaccine based on the extracellular domain of M2:
weak protection mediated via antibody-dependent NK cell activity. J Immunol, 2004.
172(9): p. 5598-605.

193

174.

Tate, M.D., et al., Neutrophils ameliorate lung injury and the development of severe
disease during influenza infection. J Immunol, 2009. 183(11): p. 7441-50.

175.

Narasaraju, T., et al., Excessive neutrophils and neutrophil extracellular traps contribute
to acute lung injury of influenza pneumonitis. Am J Pathol, 2011. 179(1): p. 199-210.

176.

Wang, J.P., et al., Toll-like receptor-mediated activation of neutrophils by influenza A
virus. Blood, 2008. 112(5): p. 2028-34.

177.

Fialkow, L., Y. Wang, and G.P. Downey, Reactive oxygen and nitrogen species as
signaling molecules regulating neutrophil function. Free Radic Biol Med, 2007. 42(2): p.
153-64.

178.

McDonald, B., et al., Intravascular danger signals guide neutrophils to sites of sterile
inflammation. Science, 2010. 330(6002): p. 362-6.

179.

Belshe, R.B., et al., Correlates of immune protection induced by live, attenuated, coldadapted, trivalent, intranasal influenza virus vaccine. J Infect Dis, 2000. 181(3): p. 11337.

180.

Yu, X., et al., Neutralizing antibodies derived from the B cells of 1918 influenza
pandemic survivors. Nature, 2008. 455(7212): p. 532-6.

181.

Nayak, B., et al., Contributions of the avian influenza virus HA, NA, and M2 surface
proteins to the induction of neutralizing antibodies and protective immunity. J Virol,
2010. 84(5): p. 2408-20.

182.

Topham, D.J. and P.C. Doherty, Clearance of an influenza A virus by CD4+ T cells is
inefficient in the absence of B cells. J Virol, 1998. 72(1): p. 882-5.

183.

Fazekas, G., et al., IgG isotype distribution of local and systemic immune responses
induced by influenza virus infection. Eur J Immunol, 1994. 24(12): p. 3063-7.

184.

Quan, F.S., et al., Induction of heterosubtypic immunity to influenza virus by intranasal
immunization. J Virol, 2008. 82(3): p. 1350-9.

185.

Heer, A.K., et al., TLR signaling fine-tunes anti-influenza B cell responses without
regulating effector T cell responses. J Immunol, 2007. 178(4): p. 2182-91.

194

186.

Vossen, M.T., et al., Viral immune evasion: a masterpiece of evolution. Immunogenetics,
2002. 54(8): p. 527-42.

187.

Burns-Naas, L.A., B.J. Meade, and A.E. Munson, TOXIC RESPONSES OF THE
IMMUNE SYSTEM, in CASARETT AND DOULL’S TOXICOLOGY
THE BASIC SCIENCE OF POISONS, C.D. Klaassen, Editor. 2001, McGraw-Hill: New York.
188.

Compton, W.M., et al., Prevalence of marijuana use disorders in the United States:
1991-1992 and 2001-2002. Jama, 2004. 291(17): p. 2114-21.

189.

Administration, S.A.a.M.H.S., Results from the 2007 National Survey on Drug Use and
Health: National Findings, D.O.H.A.H. SERVICES, Editor. 2007: RocKville, MD.

190.

Hall, W. and L. Degenhardt, Adverse health effects of non-medical cannabis use. Lancet,
2009. 374(9698): p. 1383-91.

191.

Cinti, S., Medical marijuana in HIV-positive patients: what do we know? J Int Assoc
Physicians AIDS Care (Chic), 2009. 8(6): p. 342-6.

192.

Ogborne, A.C., R.G. Smart, and E.M. Adlaf, Self-reported medical use of marijuana: a
survey of the general population. Cmaj, 2000. 162(12): p. 1685-6.

193.

Gaoni, Y. and R. Mechoulam, Isolation, Structure, and Partial Synthesis of an Active
Constituent of Hashish. Journal of the American Chemical Society, 1964. 86(8): p. 1646 1647.

194.

Turner, J.C., J.K. Hemphill, and P.G. Mahlberg, Cannabinoid composition and gland
distribution in clones of Cannabis sativa L. (Cannabaceae). Bull Narc, 1978. 30(1): p.
55-65.

195.

Pertwee, R.G., The diverse CB1 and CB2 receptor pharmacology of three plant
cannabinoids:
delta9-tetrahydrocannabinol,
cannabidiol
and
delta9tetrahydrocannabivarin. Br J Pharmacol, 2008. 153(2): p. 199-215.

196.

Howlett, A.C., et al., International Union of Pharmacology. XXVII. Classification of
cannabinoid receptors. Pharmacol Rev, 2002. 54(2): p. 161-202.

195

197.

Hanus, L.O., Discovery and isolation of anandamide and other endocannabinoids. Chem
Biodivers, 2007. 4(8): p. 1828-41.

198.

Piomelli, D., The molecular logic of endocannabinoid signalling. Nat Rev Neurosci,
2003. 4(11): p. 873-84.

199.

Pacher, P., S. Batkai, and G. Kunos, The endocannabinoid system as an emerging target
of pharmacotherapy. Pharmacol Rev, 2006. 58(3): p. 389-462.

200.

Stapleton, J.A., Trial comes too late as psychiatric side effects end hope for rimonabant.
Addiction, 2009. 104(2): p. 277-8.

201.

Christensen, R., et al., Efficacy and safety of the weight-loss drug rimonabant: a metaanalysis of randomised trials. Lancet, 2007. 370(9600): p. 1706-13.

202.

Di Marzo, V. and J.P. Despres, CB1 antagonists for obesity--what lessons have we
learned from rimonabant? Nat Rev Endocrinol, 2009. 5(11): p. 633-8.

203.

Uchiyama, N., et al., Chemical analysis of synthetic cannabinoids as designer drugs in
herbal products. Forensic Sci Int, 2010. 198(1-3): p. 31-8.

204.

Huestis, M.A., Pharmacokinetics and metabolism of the plant cannabinoids, delta9tetrahydrocannabinol, cannabidiol and cannabinol. Handb Exp Pharmacol, 2005(168): p.
657-90.

205.

McGilveray, I.J., Pharmacokinetics of cannabinoids. Pain Res Manag, 2005. 10 Suppl A:
p. 15A-22A.

206.

Hunault, C.C., et al., Disposition of smoked cannabis with high Delta(9)tetrahydrocannabinol content: A kinetic model. Toxicol Appl Pharmacol, 2010.

207.

Klausner, H.A. and J.V. Dingell, The metabolism and excretion of delta 9tetrahydrocannabinol in the rat. Life Sci I, 1971. 10(1): p. 49-59.

208.

Watanabe, K., et al., Cytochrome P450 enzymes involved in the metabolism of
tetrahydrocannabinols and cannabinol by human hepatic microsomes. Life Sci, 2007.
80(15): p. 1415-9.

196

209.

Mazur, A., et al., Characterization of human hepatic and extrahepatic UDPglucuronosyltransferase enzymes involved in the metabolism of classic cannabinoids.
Drug Metab Dispos, 2009. 37(7): p. 1496-504.

210.

Menetrey, A., et al., Assessment of driving capability through the use of clinical and
psychomotor tests in relation to blood cannabinoids levels following oral administration
of 20 mg dronabinol or of a cannabis decoction made with 20 or 60 mg Delta9-THC. J
Anal Toxicol, 2005. 29(5): p. 327-38.

211.

Kemp, P.M., et al., Cannabinoids in humans. I. Analysis of delta 9-tetrahydrocannabinol
and six metabolites in plasma and urine using GC-MS. J Anal Toxicol, 1995. 19(5): p.
285-91.

212.

Klein, C., et al., Cannabidiol potentiates Delta(9)-tetrahydrocannabinol (THC)
behavioural effects and alters THC pharmacokinetics during acute and chronic treatment
in adolescent rats. Psychopharmacology (Berl), 2011.

213.

Maccarrone, M., et al., The endocannabinoid system and its relevance for nutrition. Annu
Rev Nutr, 2010. 30: p. 423-40.

214.

Nomura, D.K., et al., Activation of the endocannabinoid system by organophosphorus
nerve agents. Nat Chem Biol, 2008. 4(6): p. 373-8.

215.

Long, J.Z., D.K. Nomura, and B.F. Cravatt, Characterization of monoacylglycerol lipase
inhibition reveals differences in central and peripheral endocannabinoid metabolism.
Chem Biol, 2009. 16(7): p. 744-53.

216.

Munro, S., K.L. Thomas, and M. Abu-Shaar, Molecular characterization of a peripheral
receptor for cannabinoids. Nature, 1993. 365(6441): p. 61-5.

217.

Shire, D., et al., An amino-terminal variant of the central cannabinoid receptor resulting
from alternative splicing. J Biol Chem, 1995. 270(8): p. 3726-31.

218.

Matsuda, L.A., et al., Structure of a cannabinoid receptor and functional expression of
the cloned cDNA. Nature, 1990. 346(6284): p. 561-4.

219.

Kaminski, N.E., et al., Identification of a functionally relevant cannabinoid receptor on
mouse spleen cells that is involved in cannabinoid-mediated immune modulation. Mol
Pharmacol, 1992. 42(5): p. 736-42.

197

220.

Gomez Del Pulgar, T., et al., Cannabinoids protect astrocytes from ceramide-induced
apoptosis through the phosphatidylinositol 3-kinase/protein kinase B pathway. J Biol
Chem, 2002. 277(39): p. 36527-33.

221.

Ehrhart, J., et al., Stimulation of cannabinoid receptor 2 (CB2) suppresses microglial
activation. J Neuroinflammation, 2005. 2: p. 29.

222.

Fernandez-Ruiz, J., et al., Cannabinoid CB2 receptor: a new target for controlling neural
cell survival? Trends Pharmacol Sci, 2007. 28(1): p. 39-45.

223.

Galiegue, S., et al., Expression of central and peripheral cannabinoid receptors in human
immune tissues and leukocyte subpopulations. Eur J Biochem, 1995. 232(1): p. 54-61.

224.

Schatz, A.R., et al., Cannabinoid receptors CB1 and CB2: a characterization of
expression and adenylate cyclase modulation within the immune system. Toxicol Appl
Pharmacol, 1997. 142(2): p. 278-87.

225.

Carayon, P., et al., Modulation and functional involvement of CB2 peripheral
cannabinoid receptors during B-cell differentiation. Blood, 1998. 92(10): p. 3605-15.

226.

Jiang, S., et al., Expression and function of cannabinoid receptors CB1 and CB2 and
their cognate cannabinoid ligands in murine embryonic stem cells. PLoS ONE, 2007.
2(7): p. e641.

227.

Batkai, S., et al., Cannabinoid antagonist SR-141716 inhibits endotoxic hypotension by a
cardiac mechanism not involving CB1 or CB2 receptors. Am J Physiol Heart Circ
Physiol, 2004. 287(2): p. H595-600.

228.

Jarai, Z., et al., Cannabinoid-induced mesenteric vasodilation through an endothelial site
distinct from CB1 or CB2 receptors. Proc Natl Acad Sci U S A, 1999. 96(24): p. 1413641.

229.

Kaplan, B.L., et al., 2-Arachidonoyl-glycerol suppresses interferon-gamma production in
phorbol ester/ionomycin-activated mouse splenocytes independent of CB1 or CB2. J
Leukoc Biol, 2005. 77(6): p. 966-74.

230.

Rockwell, C.E., et al., Interleukin-2 suppression by 2-arachidonyl glycerol is mediated
through peroxisome proliferator-activated receptor gamma independently of cannabinoid
receptors 1 and 2. Mol Pharmacol, 2006. 70(1): p. 101-11.

198

231.

Rao, G.K. and N.E. Kaminski, Cannabinoid-mediated elevation of intracellular calcium:
a structure-activity relationship. J Pharmacol Exp Ther, 2006. 317(2): p. 820-9.

232.

Brown, A.J., Novel cannabinoid receptors. Br J Pharmacol, 2007. 152(5): p. 567-75.

233.

Rao, G.K., W. Zhang, and N.E. Kaminski, Cannabinoid receptor-mediated regulation of
intracellular calcium by delta(9)-tetrahydrocannabinol in resting T cells. J Leukoc Biol,
2004. 75(5): p. 884-92.

234.

Kaplan, B.L., C.E. Rockwell, and N.E. Kaminski, Evidence for cannabinoid receptordependent and -independent mechanisms of action in leukocytes. J Pharmacol Exp Ther,
2003. 306(3): p. 1077-85.

235.

Kaplan, B.L., A.E. Springs, and N.E. Kaminski, The profile of immune modulation by
cannabidiol (CBD) involves deregulation of nuclear factor of activated T cells (NFAT).
Biochem Pharmacol, 2008. 76(6): p. 726-37.

236.

Raman, P., et al., 15-Deoxy-{Delta}12,14-prostaglandin J2-Glycerol Ester, a Putative
Metabolite of 2-Arachidonyl Glycerol, Activates Peroxisome Proliferator Activated
Receptor {gamma}. Mol Pharmacol, 2011. 80(1): p. 201-9.

237.

Irving, A., New blood brothers: the GPR55 and CB(2) partnership. Cell Res, 2011.

238.

McHugh, D., et al., Delta(9) -THC and N-arachidonyl glycine are full agonists at GPR18
and cause migration in the human endometrial cell line, HEC-1B. Br J Pharmacol, 2011.

239.

Berdyshev, E.V., Cannabinoid receptors and the regulation of immune response. Chem
Phys Lipids, 2000. 108(1-2): p. 169-90.

240.

Bayewitch, M., et al., The peripheral cannabinoid receptor: adenylate cyclase inhibition
and G protein coupling. FEBS Lett, 1995. 375(1-2): p. 143-7.

241.

Schatz, A.R., et al., Cannabinoid receptors CB1 and CB2: a characterization of
expression and adenylate cyclase modulation within the immune system. Toxicol Appl
Pharmacol, 1996. 142(2): p. 278-87.

242.

Vroon, A., C.J. Heijnen, and A. Kavelaars, GRKs and arrestins: regulators of migration
and inflammation. J Leukoc Biol, 2006. 80(6): p. 1214-21.

199

243.

Hsieh, C., et al., Internalization and recycling of the CB1 cannabinoid receptor. J
Neurochem, 1999. 73(2): p. 493-501.

244.

Bouaboula, M., et al., Signaling pathway associated with stimulation of CB2 peripheral
cannabinoid receptor. Involvement of both mitogen-activated protein kinase and
induction of Krox-24 expression. Eur J Biochem, 1996. 237(3): p. 704-11.

245.

Suzuki, A., et al., Role of PTEN/PI3K pathway in endothelial cells. Biochem Soc Trans,
2007. 35(Pt 2): p. 172-6.

246.

Milligan, G. and E. Kostenis, Heterotrimeric G-proteins: a short history. Br J Pharmacol,
2006. 147 Suppl 1: p. S46-55.

247.

Dupre, D.J., et al., The role of Gbetagamma subunits in the organization, assembly, and
function of GPCR signaling complexes. Annu Rev Pharmacol Toxicol, 2009. 49: p. 3156.

248.

Luttrell, L.M. and R.J. Lefkowitz, The role of beta-arrestins in the termination and
transduction of G-protein-coupled receptor signals. J Cell Sci, 2002. 115(Pt 3): p. 45565.

249.

Miller, W.E. and R.J. Lefkowitz, Expanding roles for beta-arrestins as scaffolds and
adapters in GPCR signaling and trafficking. Curr Opin Cell Biol, 2001. 13(2): p. 139-45.

250.

Hupfeld, C.J., et al., Insulin-induced beta-arrestin1 Ser-412 phosphorylation is a
mechanism for desensitization of ERK activation by Galphai-coupled receptors. J Biol
Chem, 2005. 280(2): p. 1016-23.

251.

Abramow-Newerly, M., et al., RGS proteins have a signalling complex: interactions
between RGS proteins and GPCRs, effectors, and auxiliary proteins. Cell Signal, 2006.
18(5): p. 579-91.

252.

Mackie, K., Cannabinoid receptor homo- and heterodimerization. Life Sci, 2005. 77(14):
p. 1667-73.

253.

Barnes, P.J., Receptor heterodimerization: a new level of cross-talk. J Clin Invest, 2006.
116(5): p. 1210-2.

200

254.

Prinster, S.C., C. Hague, and R.A. Hall, Heterodimerization of g protein-coupled
receptors: specificity and functional significance. Pharmacol Rev, 2005. 57(3): p. 289-98.

255.

Henstridge, C.M., et al., The GPR55 ligand L-alpha-lysophosphatidylinositol promotes
RhoA-dependent Ca2+ signaling and NFAT activation. Faseb J, 2009. 23(1): p. 183-93.

256.

Ning, Y., et al., Endogenous and synthetic agonists of GPR119 differ in signalling
pathways and their effects on insulin secretion in MIN6c4 insulinoma cells. Br J
Pharmacol, 2008. 155(7): p. 1056-65.

257.

Ong, H.L., et al., Dynamic assembly of TRPC1-STIM1-Orai1 ternary complex is involved
in store-operated calcium influx. Evidence for similarities in store-operated and calcium
release-activated calcium channel components. J Biol Chem, 2007. 282(12): p. 9105-16.

258.

Beech, D.J., TRPC1: store-operated channel and more. Pflugers Arch, 2005. 451(1): p.
53-60.

259.

Shipley, J.M. and D.J. Waxman, Down-regulation of STAT5b transcriptional activity by
ligand-activated peroxisome proliferator-activated receptor (PPAR) alpha and
PPARgamma. Mol Pharmacol, 2003. 64(2): p. 355-64.

260.

Klein, T.W. and G.A. Cabral, Cannabinoid-induced immune suppression and modulation
of antigen-presenting cells. J Neuroimmune Pharmacol, 2006. 1(1): p. 50-64.

261.

Zheng, Z.M., S. Specter, and H. Friedman, Inhibition by delta-9-tetrahydrocannabinol of
tumor necrosis factor alpha production by mouse and human macrophages. Int J
Immunopharmacol, 1992. 14(8): p. 1445-52.

262.

Jeon, Y.J., et al., Attenuation of inducible nitric oxide synthase gene expression by delta
9-tetrahydrocannabinol is mediated through the inhibition of nuclear factor- kappa B/Rel
activation. Mol Pharmacol, 1996. 50(2): p. 334-41.

263.

Raborn, E.S., et al., The cannabinoid delta-9-tetrahydrocannabinol mediates inhibition of
macrophage chemotaxis to RANTES/CCL5: linkage to the CB2 receptor. J Neuroimmune
Pharmacol, 2008. 3(2): p. 117-29.

264.

Carlisle, S.J., et al., Differential expression of the CB2 cannabinoid receptor by rodent
macrophages and macrophage-like cells in relation to cell activation. Int
Immunopharmacol, 2002. 2(1): p. 69-82.

201

265.

Do, Y., et al., Activation through cannabinoid receptors 1 and 2 on dendritic cells
triggers NF-kappaB-dependent apoptosis: novel role for endogenous and exogenous
cannabinoids in immunoregulation. J Immunol, 2004. 173(4): p. 2373-82.

266.

Wacnik, P.W., et al., Cannabinoids affect dendritic cell (DC) potassium channel function
and modulate DC T cell stimulatory capacity. J Immunol, 2008. 181(5): p. 3057-66.

267.

Kaminski, N.E., et al., Suppression of the humoral immune response by cannabinoids is
partially mediated through inhibition of adenylate cyclase by a pertussis toxin-sensitive
G-protein coupled mechanism. Biochem Pharmacol, 1994. 48(10): p. 1899-908.

268.

Howlett, A.C., J.M. Qualy, and L.L. Khachatrian, Involvement of Gi in the inhibition of
adenylate cyclase by cannabimimetic drugs. Mol Pharmacol, 1986. 29(3): p. 307-13.

269.

Lu, T., et al., Role of cannabinoid receptors in Delta-9-tetrahydrocannabinol suppression
of IL-12p40 in mouse bone marrow-derived dendritic cells infected with Legionella
pneumophila. Eur J Pharmacol, 2006. 532(1-2): p. 170-7.

270.

Lu, T., et al., Cannabinoid treatment suppresses the T-helper cell-polarizing function of
mouse dendritic cells stimulated with Legionella pneumophila infection. J Pharmacol Exp
Ther, 2006. 319(1): p. 269-76.

271.

Hegde, V.L., M. Nagarkatti, and P.S. Nagarkatti, Cannabinoid receptor activation leads
to massive mobilization of myeloid-derived suppressor cells with potent
immunosuppressive properties. Eur J Immunol, 2010. 40(12): p. 3358-71.

272.

McHugh, D., et al., Inhibition of human neutrophil chemotaxis by endogenous
cannabinoids and phytocannabinoids: evidence for a site distinct from CB1 and CB2.
Mol Pharmacol, 2008. 73(2): p. 441-50.

273.

Murikinati, S., et al., Activation of cannabinoid 2 receptors protects against cerebral
ischemia by inhibiting neutrophil recruitment. Faseb J, 2010. 24(3): p. 788-98.

274.

Kraft, B., W. Wintersberger, and H.G. Kress, Cannabinoid receptor-independent
suppression of the superoxide generation of human neutrophils (PMN) by CP55 940, but
not by anandamide. Life Sci, 2004. 75(8): p. 969-77.

202

275.

Kraft, B. and H.G. Kress, Indirect CB2 receptor and mediator-dependent stimulation of
human whole-blood neutrophils by exogenous and endogenous cannabinoids. J
Pharmacol Exp Ther, 2005. 315(2): p. 641-7.

276.

Rosenkrantz, H., A.J. Miller, and H.J. Esber, delta9-Tetrahydrocannabinol suppression
of the primary immune response in rats. J Toxicol Environ Health, 1975. 1(1): p. 119-25.

277.

Nahas, G.G., A. Morishima, and B. Desoize, Effects of cannabinoids on macromolecular
synthesis and replication of cultured lymphocytes. Fed Proc, 1977. 36(5): p. 1748-52.

278.

Klein, T.W., et al., The effect of delta-9-tetrahydrocannabinol and 11-hydroxy-delta-9tetrahydrocannabinol on T-lymphocyte and B-lymphocyte mitogen responses. J
Immunopharmacol, 1985. 7(4): p. 451-66.

279.

Springs, A.E., et al., Effects of targeted deletion of cannabinoid receptors CB1 and CB2
on immune competence and sensitivity to immune modulation by Delta9tetrahydrocannabinol. J Leukoc Biol, 2008. 84(6): p. 1574-84.

280.

Zhu, W., et al., delta-9-Tetrahydrocannabinol (THC) decreases the number of high and
intermediate affinity IL-2 receptors of the IL-2 dependent cell line NKB61A2. Int J
Immunopharmacol, 1993. 15(3): p. 401-8.

281.

Condie, R., et al., Cannabinoid inhibition of adenylate cyclase-mediated signal
transduction and interleukin 2 (IL-2) expression in the murine T-cell line, EL4.IL-2. J
Biol Chem, 1996. 271(22): p. 13175-83.

282.

Yea, S.S., K.H. Yang, and N.E. Kaminski, Role of nuclear factor of activated T-cells and
activator protein-1 in the inhibition of interleukin-2 gene transcription by cannabinol in
EL4 T-cells. J Pharmacol Exp Ther, 2000. 292(2): p. 597-605.

283.

Faubert, B.L. and N.E. Kaminski, AP-1 activity is negatively regulated by cannabinol
through inhibition of its protein components, c-fos and c-jun. J Leukoc Biol, 2000. 67(2):
p. 259-66.

284.

Rao, G.K. and N.E. Kaminski, Induction of intracellular calcium elevation by Delta9tetrahydrocannabinol in T cells involves TRPC1 channels. J Leukoc Biol, 2006. 79(1): p.
202-13.

203

285.

Lu, H., et al., Suppression of T cell costimulator ICOS by Delta9-tetrahydrocannabinol. J
Leukoc Biol, 2009. 85(2): p. 322-9.

286.

Newton, C.A., et al., CB(1) and CB(2) cannabinoid receptors mediate different aspects of
delta-9-tetrahydrocannabinol (THC)-induced T helper cell shift following immune
activation by Legionella pneumophila infection. J Neuroimmune Pharmacol, 2009. 4(1):
p. 92-102.

287.

Bouaboula, M., et al., Cannabinoid-receptor expression in human leukocytes. Eur J
Biochem, 1993. 214(1): p. 173-80.

288.

Daaka, Y., H. Friedman, and T.W. Klein, Cannabinoid receptor proteins are increased in
Jurkat, human T-cell line after mitogen activation. J Pharmacol Exp Ther, 1996. 276(2):
p. 776-83.

289.

Pross, S.H., et al., Differential suppression of T-cell subpopulations by thc (delta-9tetrahydrocannabinol). Int J Immunopharmacol, 1990. 12(5): p. 539-44.

290.

Klein, T.W., et al., Marijuana components suppress induction and cytolytic function of
murine cytotoxic T cells in vitro and in vivo. J Toxicol Environ Health, 1991. 32(4): p.
465-77.

291.

Fischer-Stenger, K., A.W. Updegrove, and G.A. Cabral, Delta 9-tetrahydrocannabinol
decreases cytotoxic T lymphocyte activity to herpes simplex virus type 1-infected cells.
Proc Soc Exp Biol Med, 1992. 200(3): p. 422-30.

292.

Klein, T.W., C. Newton, and H. Friedman, Inhibition of natural killer cell function by
marijuana components. J Toxicol Environ Health, 1987. 20(4): p. 321-32.

293.

Kawakami, Y., et al., Suppression by cannabinoids of a cloned cell line with natural
killer cell activity. Proc Soc Exp Biol Med, 1988. 187(3): p. 355-9.

294.

Kishimoto, S., et al., Endogenous cannabinoid receptor ligand induces the migration of
human natural killer cells. J Biochem, 2005. 137(2): p. 217-23.

295.

Kusher, D.I., et al., Effect of the psychoactive metabolite of marijuana, delta 9tetrahydrocannabinol (THC), on the synthesis of tumor necrosis factor by human large
granular lymphocytes. Cell Immunol, 1994. 154(1): p. 99-108.

204

296.

Derocq, J.M., et al., Cannabinoids enhance human B-cell growth at low nanomolar
concentrations. FEBS Lett, 1995. 369(2-3): p. 177-82.

297.

Lee, M., K.H. Yang, and N.E. Kaminski, Effects of putative cannabinoid receptor
ligands, anandamide and 2-arachidonyl-glycerol, on immune function in B6C3F1 mouse
splenocytes. J Pharmacol Exp Ther, 1995. 275(2): p. 529-36.

298.

Srivastava, M.D., B.I. Srivastava, and B. Brouhard, Delta9 tetrahydrocannabinol and
cannabidiol alter cytokine production by human immune cells. Immunopharmacology,
1998. 40(3): p. 179-85.

299.

Mishkin, E.M. and G.A. Cabral, delta-9-Tetrahydrocannabinol decreases host resistance
to herpes simplex virus type 2 vaginal infection in the B6C3F1 mouse. J Gen Virol, 1985.
66 ( Pt 12): p. 2539-49.

300.

Cabral, G.A., et al., Effect of delta 9-tetrahydrocannabinol on herpes simplex virus type 2
vaginal infection in the guinea pig. Proc Soc Exp Biol Med, 1986. 182(2): p. 181-6.

301.

Zhu, L.X., et al., Delta-9-tetrahydrocannabinol inhibits antitumor immunity by a CB2
receptor-mediated, cytokine-dependent pathway. J Immunol, 2000. 165(1): p. 373-80.

302.

Morahan, P.S., et al., Effects of cannabinoids on host resistance to Listeria
monocytogenes and herpes simplex virus. Infect Immun, 1979. 23(3): p. 670-4.

303.

Newton, C.A., T.W. Klein, and H. Friedman, Secondary immunity to Legionella
pneumophila and Th1 activity are suppressed by delta-9-tetrahydrocannabinol injection.
Infect Immun, 1994. 62(9): p. 4015-20.

304.

Klein, T.W., et al., Delta 9-tetrahydrocannabinol treatment suppresses immunity and
early IFN-gamma, IL-12, and IL-12 receptor beta 2 responses to Legionella pneumophila
infection. J Immunol, 2000. 164(12): p. 6461-6.

305.

Cabral, G.A., J.C. Lockmuller, and E.M. Mishkin, Delta 9-tetrahydrocannabinol
decreases alpha/beta interferon response to herpes simplex virus type 2 in the B6C3F1
mouse. Proc Soc Exp Biol Med, 1986. 181(2): p. 305-11.

306.

Orr, M.T., et al., Cutting edge: recombinant Listeria monocytogenes expressing a single
immune-dominant peptide confers protective immunity to herpes simplex virus-1
infection. J Immunol, 2007. 178(8): p. 4731-5.

205

307.

Zimmer, A., et al., Increased mortality, hypoactivity, and hypoalgesia in cannabinoid
CB1 receptor knockout mice. Proc Natl Acad Sci U S A, 1999. 96(10): p. 5780-5.

308.

Kunos, G. and P. Pacher, Cannabinoids cool the intestine. Nat Med, 2004. 10(7): p. 6789.

309.

Massa, F., et al., The endogenous cannabinoid system protects against colonic
inflammation. J Clin Invest, 2004. 113(8): p. 1202-9.

310.

Yoshida, T., et al., The cannabinoid CB1 receptor mediates retrograde signals for
depolarization-induced suppression of inhibition in cerebellar Purkinje cells. J Neurosci,
2002. 22(5): p. 1690-7.

311.

Ganta, C.K., et al., Central angiotensin II-enhanced splenic cytokine gene expression is
mediated by the sympathetic nervous system. Am J Physiol Heart Circ Physiol, 2005.
289(4): p. H1683-91.

312.

Buckley, N.E., et al., Immunomodulation by cannabinoids is absent in mice deficient for
the cannabinoid CB(2) receptor. Eur J Pharmacol, 2000. 396(2-3): p. 141-9.

313.

Ziring, D., et al., Formation of B and T cell subsets require the cannabinoid receptor
CB2. Immunogenetics, 2006. 58(9): p. 714-25.

314.

Ofek, O., et al., Peripheral cannabinoid receptor, CB2, regulates bone mass. Proc Natl
Acad Sci U S A, 2006. 103(3): p. 696-701.

315.

Karsak, M., et al., Attenuation of allergic contact dermatitis through the
endocannabinoid system. Science, 2007. 316(5830): p. 1494-7.

316.

Ghosh, M., et al., COX-2 suppresses tissue factor expression via endocannabinoiddirected PPARdelta activation. J Exp Med, 2007. 204(9): p. 2053-61.

317.

Kaplan, B.L., et al., The effects of targeted deletion of cannabinoid receptors CB1 and
CB2 on intranasal sensitization and challenge with adjuvant-free ovalbumin. Toxicol
Pathol, 2010. 38(3): p. 382-92.

206

318.

Sipe, J.C., et al., Reduced endocannabinoid immune modulation by a common
cannabinoid 2 (CB2) receptor gene polymorphism: possible risk for autoimmune
disorders. J Leukoc Biol, 2005. 78(1): p. 231-8.

319.

Storr, M., et al., The cannabinoid 1 receptor (CNR1) 1359 G/A polymorphism modulates
susceptibility to ulcerative colitis and the phenotype in Crohn's disease. PLoS One, 2010.
5(2): p. e9453.

320.

Kaplan, B.L., et al., The Effects of Targeted Deletion of Cannabinoid Receptors CB1 and
CB2 on Intranasal Sensitization and Challenge with Adjuvant-Free Ovalbumin. Toxicol
Pathol, 2010.

321.

Eisen, M.B., et al., Cluster analysis and display of genome-wide expression patterns.
Proc Natl Acad Sci U S A, 1998. 95(25): p. 14863-8.

322.

Lutz, M.B., et al., An advanced culture method for generating large quantities of highly
pure dendritic cells from mouse bone marrow. J Immunol Methods, 1999. 223(1): p. 7792.

323.

Brandt, K., et al., Interleukin-21 inhibits dendritic cell activation and maturation. Blood,
2003. 102(12): p. 4090-8.

324.

Ivanov, II, et al., The orphan nuclear receptor RORgammat directs the differentiation
program of proinflammatory IL-17+ T helper cells. Cell, 2006. 126(6): p. 1121-33.

325.

Lopez-Cabrera, M., et al., Transcriptional regulation of the gene encoding the human Ctype lectin leukocyte receptor AIM/CD69 and functional characterization of its tumor
necrosis factor-alpha-responsive elements. J Biol Chem, 1995. 270(37): p. 21545-51.

326.

Brenchley, J.M., et al., Expansion of activated human naive T-cells precedes effector
function. Clin Exp Immunol, 2002. 130(3): p. 432-40.

327.

Akira, S., S. Uematsu, and O. Takeuchi, Pathogen recognition and innate immunity. Cell,
2006. 124(4): p. 783-801.

328.

Asselin-Paturel, C., et al., Mouse type I IFN-producing cells are immature APCs with
plasmacytoid morphology. Nat Immunol, 2001. 2(12): p. 1144-50.

207

329.

De Vries, I.J., et al., Effective migration of antigen-pulsed dendritic cells to lymph nodes
in melanoma patients is determined by their maturation state. Cancer Res, 2003. 63(1): p.
12-7.

330.

Lin, C.L., et al., Dendritic cell chemotaxis and transendothelial migration are induced by
distinct chemokines and are regulated on maturation. Eur J Immunol, 1998. 28(12): p.
4114-22.

331.

De Smedt, T., et al., Regulation of dendritic cell numbers and maturation by
lipopolysaccharide in vivo. J Exp Med, 1996. 184(4): p. 1413-24.

332.

Poltorak, A., et al., Defective LPS signaling in C3H/HeJ and C57BL/10ScCr mice:
mutations in Tlr4 gene. Science, 1998. 282(5396): p. 2085-8.

333.

Jurk, M., et al., Human TLR7 or TLR8 independently confer responsiveness to the
antiviral compound R-848. Nat Immunol, 2002. 3(6): p. 499.

334.

Liu, Q.R., et al., Species differences in cannabinoid receptor 2 (CNR2 gene):
identification of novel human and rodent CB2 isoforms, differential tissue expression and
regulation by cannabinoid receptor ligands. Genes Brain Behav, 2009. 8(5): p. 519-30.

335.

Suri-Payer, E., et al., CD4+CD25+ T cells inhibit both the induction and effector
function of autoreactive T cells and represent a unique lineage of immunoregulatory
cells. J Immunol, 1998. 160(3): p. 1212-8.

336.

Piccirillo, C.A. and E.M. Shevach, Cutting edge: control of CD8+ T cell activation by
CD4+CD25+ immunoregulatory cells. J Immunol, 2001. 167(3): p. 1137-40.

337.

Bettelli, E., et al., Reciprocal developmental pathways for the generation of pathogenic
effector TH17 and regulatory T cells. Nature, 2006. 441(7090): p. 235-8.

338.

Fontenot, J.D. and A.Y. Rudensky, A well adapted regulatory contrivance: regulatory T
cell development and the forkhead family transcription factor Foxp3. Nat Immunol,
2005. 6(4): p. 331-7.

339.

Friedman, H., C. Newton, and T.W. Klein, Microbial infections, immunomodulation, and
drugs of abuse. Clin Microbiol Rev, 2003. 16(2): p. 209-19.

208

340.

Ashton, C.H., Pharmacology and effects of cannabis: a brief review. Br J Psychiatry,
2001. 178: p. 101-6.

341.

Azorlosa, J.L., et al., Marijuana smoking: effect of varying delta 9-tetrahydrocannabinol
content and number of puffs. J Pharmacol Exp Ther, 1992. 261(1): p. 114-22.

342.

Huestis, M.A. and E.J. Cone, Relationship of Delta 9-tetrahydrocannabinol
concentrations in oral fluid and plasma after controlled administration of smoked
cannabis. J Anal Toxicol, 2004. 28(6): p. 394-9.

343.

Sundrud, M.S., et al., Genetic reprogramming of primary human T cells reveals
functional plasticity in Th cell differentiation. J Immunol, 2003. 171(7): p. 3542-9.

344.

Tucker, A.N., S.J. Vore, and M.I. Luster, Suppression of B cell differentiation by 2,3,7,8tetrachlorodibenzo-p-dioxin. Mol Pharmacol, 1986. 29(4): p. 372-7.

345.

Piccolella, E., et al., Effect of glucocorticoids on the development of suppressive activity
in human lymphocyte response to a polysaccharide purified from Candida albicans. J
Immunol, 1985. 134(2): p. 1166-71.

346.

Hess, A.D. and P.J. Tutschka, Effect of cyclosporin A on human lymphocyte responses in
vitro. I. CsA allows for the expression of alloantigen-activated suppressor cells while
preferentially inhibiting the induction of cytolytic effector lymphocytes in MLR. J
Immunol, 1980. 124(6): p. 2601-8.

347.

Castellanos, M.C., et al., Expression of the leukocyte early activation antigen CD69 is
regulated by the transcription factor AP-1. J Immunol, 1997. 159(11): p. 5463-73.

348.

Castellanos Mdel, C., et al., Multiple cis-acting elements regulate the expression of the
early T cell activation antigen CD69. Eur J Immunol, 2002. 32(11): p. 3108-17.

349.

Vazquez, B.N., et al., CD69 gene is differentially regulated in T and B cells by
evolutionarily conserved promoter-distal elements. J Immunol, 2009. 183(10): p. 651321.

350.

Testi, R., et al., The CD69 receptor: a multipurpose cell-surface trigger for
hematopoietic cells. Immunol Today, 1994. 15(10): p. 479-83.

209

351.

Rodrigues Mascarenhas, S., et al., CD69 expression induced by thapsigargin, phorbol
ester and ouabain on thymocytes is dependent on external Ca2+ entry. Life Sci, 2003.
73(8): p. 1037-51.

352.

Ryberg, E., et al., The orphan receptor GPR55 is a novel cannabinoid receptor. Br J
Pharmacol, 2007. 152(7): p. 1092-101.

353.

Lauckner, J.E., et al., GPR55 is a cannabinoid receptor that increases intracellular
calcium and inhibits M current. Proc Natl Acad Sci U S A, 2008. 105(7): p. 2699-704.

354.

Oka, S., et al., Identification of GPR55 as a lysophosphatidylinositol receptor. Biochem
Biophys Res Commun, 2007. 362(4): p. 928-34.

355.

Parish, I.A., et al., The molecular signature of CD8+ T cells undergoing deletional
tolerance. Blood, 2009. 113(19): p. 4575-85.

356.

Macian, F., et al., Transcriptional mechanisms underlying lymphocyte tolerance. Cell,
2002. 109(6): p. 719-31.

357.

Faubert Kaplan, B.L. and N.E. Kaminski, Cannabinoids inhibit the activation of ERK
MAPK in PMA/Io-stimulated mouse splenocytes. Int Immunopharmacol, 2003. 3(10-11):
p. 1503-10.

358.

Cabral, G.A. and A. Staab, Effects on the immune system. Handb Exp Pharmacol,
2005(168): p. 385-423.

359.

Beck-Schimmer, B., et al., Alveolar macrophages regulate neutrophil recruitment in
endotoxin-induced lung injury. Respir Res, 2005. 6: p. 61.

360.

Nathan, C., Neutrophils and immunity: challenges and opportunities. Nat Rev Immunol,
2006. 6(3): p. 173-82.

361.

Wang, S.Z., et al., Clara cell secretory protein modulates lung inflammatory and immune
responses to respiratory syncytial virus infection. J Immunol, 2003. 171(2): p. 1051-60.

362.

Sandoval-Montes, C. and L. Santos-Argumedo, CD38 is expressed selectively during the
activation of a subset of mature T cells with reduced proliferation but improved potential
to produce cytokines. J Leukoc Biol, 2005. 77(4): p. 513-21.

210

363.

Cox, C.A., et al., Both Th1 and Th17 are immunopathogenic but differ in other key
biological activities. J Immunol, 2008. 180(11): p. 7414-22.

364.

Chang, M., et al., NF-kappa B1 p105 regulates T cell homeostasis and prevents chronic
inflammation. J Immunol, 2009. 182(5): p. 3131-8.

365.

Jenner, R.G. and R.A. Young, Insights into host responses against pathogens from
transcriptional profiling. Nat Rev Microbiol, 2005. 3(4): p. 281-94.

366.

Ludwiczek, S., et al., Cytokine-mediated regulation of iron transport in human monocytic
cells. Blood, 2003. 101(10): p. 4148-54.

367.

Kim, S. and P. Ponka, Effects of interferon-gamma and lipopolysaccharide on
macrophage iron metabolism are mediated by nitric oxide-induced degradation of iron
regulatory protein 2. J Biol Chem, 2000. 275(9): p. 6220-6.

368.

McCoy, K.L., D. Gainey, and G.A. Cabral, delta 9-Tetrahydrocannabinol modulates
antigen processing by macrophages. J Pharmacol Exp Ther, 1995. 273(3): p. 1216-23.

369.

McCoy, K.L., et al., Cannabinoid inhibition of the processing of intact lysozyme by
macrophages: evidence for CB2 receptor participation. J Pharmacol Exp Ther, 1999.
289(3): p. 1620-5.

370.

Chuchawankul, S., et al., Role of cannabinoid receptors in inhibiting macrophage
costimulatory activity. Int Immunopharmacol, 2004. 4(2): p. 265-78.

371.

Clements, D.J., M. Matveyeva, and K.L. McCoy, Delta9-tetrahydrocannabinol
suppresses macrophage costimulation by decreasing heat-stable antigen expression. Int J
Immunopharmacol, 1998. 20(8): p. 415-28.

372.

Baskin, C.R., et al., Early and sustained innate immune response defines pathology and
death in nonhuman primates infected by highly pathogenic influenza virus. Proc Natl
Acad Sci U S A, 2009. 106(9): p. 3455-60.

373.

Brydon, E.W., S.J. Morris, and C. Sweet, Role of apoptosis and cytokines in influenza
virus morbidity. FEMS Microbiol Rev, 2005. 29(4): p. 837-50.

211

374.

Middel, P., et al., Increased number of mature dendritic cells in Crohn's disease:
evidence for a chemokine mediated retention mechanism. Gut, 2006. 55(2): p. 220-7.

375.

Serafini, B., et al., Intracerebral recruitment and maturation of dendritic cells in the
onset and progression of experimental autoimmune encephalomyelitis. Am J Pathol,
2000. 157(6): p. 1991-2002.

376.

Ludewig, B., et al., Dendritic cells induce autoimmune diabetes and maintain disease via
de novo formation of local lymphoid tissue. J Exp Med, 1998. 188(8): p. 1493-501.

377.

Penna, G., S. Sozzani, and L. Adorini, Cutting edge: selective usage of chemokine
receptors by plasmacytoid dendritic cells. J Immunol, 2001. 167(4): p. 1862-6.

378.

Asselin-Paturel, C., et al., Type I interferon dependence of plasmacytoid dendritic cell
activation and migration. J Exp Med, 2005. 201(7): p. 1157-67.

379.

Kirby, A.C., J.G. Raynes, and P.M. Kaye, CD11b regulates recruitment of alveolar
macrophages but not pulmonary dendritic cells after pneumococcal challenge. J Infect
Dis, 2006. 193(2): p. 205-13.

380.

Kohrgruber, N., et al., Survival, maturation, and function of CD11c- and CD11c+
peripheral blood dendritic cells are differentially regulated by cytokines. J Immunol,
1999. 163(6): p. 3250-9.

381.

Sallusto, F. and A. Lanzavecchia, Mobilizing dendritic cells for tolerance, priming, and
chronic inflammation. J Exp Med, 1999. 189(4): p. 611-4.

382.

Randolph, G.J., J. Ochando, and S. Partida-Sanchez, Migration of dendritic cell subsets
and their precursors. Annu Rev Immunol, 2008. 26: p. 293-316.

383.

Laskin, D.L., B. Weinberger, and J.D. Laskin, Functional heterogeneity in liver and lung
macrophages. J Leukoc Biol, 2001. 70(2): p. 163-70.

384.

Allen, S.J., S.E. Crown, and T.M. Handel, Chemokine: receptor structure, interactions,
and antagonism. Annu Rev Immunol, 2007. 25: p. 787-820.

212

385.

Montecucco, F., et al., CB2 cannabinoid receptor agonist JWH-015 modulates human
monocyte migration through defined intracellular signaling pathways. Am J Physiol
Heart Circ Physiol, 2008. 294(3): p. H1145-55.

386.

Kishimoto, S., et al., 2-arachidonoylglycerol induces the migration of HL-60 cells
differentiated into macrophage-like cells and human peripheral blood monocytes through
the cannabinoid CB2 receptor-dependent mechanism. J Biol Chem, 2003. 278(27): p.
24469-75.

387.

Miller, A.M. and N. Stella, CB2 receptor-mediated migration of immune cells: it can go
either way. Br J Pharmacol, 2008. 153(2): p. 299-308.

388.

Sacerdote, P., et al., In vivo and in vitro treatment with the synthetic cannabinoid CP55,
940 decreases the in vitro migration of macrophages in the rat: involvement of both CB1
and CB2 receptors. J Neuroimmunol, 2000. 109(2): p. 155-63.

389.

Cervantes, D., C. Crosby, and Y. Xiang, Arrestin orchestrates crosstalk between G
protein-coupled receptors to modulate the spatiotemporal activation of ERK MAPK. Circ
Res, 2010. 106(1): p. 79-88.

390.

Bouaboula, M., et al., Gi protein modulation induced by a selective inverse agonist for
the peripheral cannabinoid receptor CB2: implication for intracellular signalization
cross-regulation. Mol Pharmacol, 1999. 55(3): p. 473-80.

391.

Perandones, C.E., et al., Regulation of apoptosis in vitro in mature murine spleen T cells.
J Immunol, 1993. 151(7): p. 3521-9.

392.

Croxford, J.L. and S.D. Miller, Immunoregulation of a viral model of multiple sclerosis
using the synthetic cannabinoid R+WIN55,212. J Clin Invest, 2003. 111(8): p. 1231-40.

393.

Downer, E.J., et al., Identification of the Synthetic Cannabinoid R(+)WIN55,212-2 as a
Novel Regulator of IFN Regulatory Factor 3 Activation and IFN-{beta} Expression:
RELEVANCE TO THERAPEUTIC EFFECTS IN MODELS OF MULTIPLE SCLEROSIS.
J Biol Chem, 2011. 286(12): p. 10316-28.

394.

Le Goffic, R., et al., Cutting Edge: Influenza A virus activates TLR3-dependent
inflammatory and RIG-I-dependent antiviral responses in human lung epithelial cells. J
Immunol, 2007. 178(6): p. 3368-72.

213

395.

Meylan, E. and J. Tschopp, Toll-like receptors and RNA helicases: two parallel ways to
trigger antiviral responses. Mol Cell, 2006. 22(5): p. 561-9.

396.

Honda, K. and T. Taniguchi, IRFs: master regulators of signalling by Toll-like receptors
and cytosolic pattern-recognition receptors. Nat Rev Immunol, 2006. 6(9): p. 644-58.

397.

Wolf, A.I., et al., Plasmacytoid dendritic cells are dispensable during primary influenza
virus infection. J Immunol, 2009. 182(2): p. 871-9.

398.

Cella, M., et al., Plasmacytoid dendritic cells activated by influenza virus and CD40L
drive a potent TH1 polarization. Nat Immunol, 2000. 1(4): p. 305-10.

399.

Colonna, M., G. Trinchieri, and Y.J. Liu, Plasmacytoid dendritic cells in immunity. Nat
Immunol, 2004. 5(12): p. 1219-26.

400.

Nagai, T., et al., Interferon-beta mediates opposing effects on interferon-gammadependent Interleukin-12 p70 secretion by human monocyte-derived dendritic cells.
Scand J Immunol, 2007. 65(2): p. 107-17.

401.

Nagai, T., et al., Timing of IFN-beta exposure during human dendritic cell maturation
and naive Th cell stimulation has contrasting effects on Th1 subset generation: a role for
IFN-beta-mediated regulation of IL-12 family cytokines and IL-18 in naive Th cell
differentiation. J Immunol, 2003. 171(10): p. 5233-43.

402.

Yen, J.H., W. Kong, and D. Ganea, IFN-beta inhibits dendritic cell migration through
STAT-1-mediated transcriptional suppression of CCR7 and matrix metalloproteinase 9. J
Immunol, 2010. 184(7): p. 3478-86.

403.

Guo, B., E.Y. Chang, and G. Cheng, The type I IFN induction pathway constrains Th17mediated autoimmune inflammation in mice. J Clin Invest, 2008. 118(5): p. 1680-90.

404.

Ramgolam, V.S., et al., IFN-beta inhibits human Th17 cell differentiation. J Immunol,
2009. 183(8): p. 5418-27.

405.

Blossey, R., L. Cardelli, and A. Phillips, Compositionality, stochasticity, and
cooperativity in dynamic models of gene regulation. Hfsp J, 2008. 2(1): p. 17-28.

214

406.

Karpinski, B.A., et al., Molecular cloning of human CREB-2: an ATF/CREB
transcription factor that can negatively regulate transcription from the cAMP response
element. Proc Natl Acad Sci U S A, 1992. 89(11): p. 4820-4.

407.

Purwar, R., et al., Induction of C3 and CCL2 by C3a in keratinocytes: a novel autocrine
amplification loop of inflammatory skin reactions. J Immunol, 2006. 177(7): p. 4444-50.

408.

Sadhu, C., et al., CD11c/CD18: novel ligands and a role in delayed-type hypersensitivity.
J Leukoc Biol, 2007. 81(6): p. 1395-403.

409.

Fitzgerald-Bocarsly, P. and E.S. Jacobs, Plasmacytoid dendritic cells in HIV infection:
striking a delicate balance. J Leukoc Biol, 2010. 87(4): p. 609-20.

410.

Kunisaki, K.M. and E.N. Janoff, Influenza in immunosuppressed populations: a review of
infection frequency, morbidity, mortality, and vaccine responses. Lancet Infect Dis, 2009.
9(8): p. 493-504.

411.

Anderson, R.M. and R.M. May, Vaccination and herd immunity to infectious diseases.
Nature, 1985. 318(6044): p. 323-9.

412.

Banchereau, J. and A.K. Palucka, Dendritic cells as therapeutic vaccines against cancer.
Nat Rev Immunol, 2005. 5(4): p. 296-306.

413.

Banchereau, J., et al., Dendritic cells: controllers of the immune system and a new
promise for immunotherapy. Novartis Found Symp, 2003. 252: p. 226-35; discussion
235-8, 257-67.

414.

Kapsenberg, M.L., Dendritic-cell control of pathogen-driven T-cell polarization. Nat Rev
Immunol, 2003. 3(12): p. 984-93.

215