University of Southern California Dissertations and Theses

Engineering chimeric antigen receptor (CAR) -modified T cells for enhanced cancer immunotherapy

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Engineering chimeric antigen receptor (CAR) -modified T cells for enhanced cancer immunotherapy

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Engineering Chimeric Antigen Receptor (CAR) -modified T cells for
Enhanced Cancer Immunotherapy

by
Si Li

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(PHARMACEUTICAL SCIENCES)

August 2017

Copyright 2017 Si Li

ii

Dedication

This thesis is dedicated to my family.

iii

Acknowledgements
First, I would like to thank my mentor Dr. Pin Wang for accepting me as part of his research
group and having been supporting and guiding me throughout the years. His patients and
encouragement during my graduate studies help to fully tap my potential. Thank you Dr. Wang
for providing me various opportunities to enrich my knowledge and experience in the doctoral
training. Meanwhile, I’d like to extend my gratitude to my committee Dr. Wei-Chiang Shen and
Dr. Curtis Okamoto for their suggestion, support and encouragement.
I am immensely thankful to all my colleagues and friends in the Wang lab (Dr. Paul Bryson, Dr.
Chupei Zhang, Dr. Jinxu Fang, Dr. Xiaolu Han, Yu-Jeong Kim, John Mac, Elizabeth Siegler,
Jennifer Rohrs, and Shuai Yang). I’d like to show my special thanks to Dr. Biliang Hu for
tutoring me when I joined the lab and Natnaree Siriwon and Xiaoyang Zhang for collaboration
on the study. It has been such a blessing to work with them.
Last but not the least, I’d like to express my deepest gratitude to my beloved family. Thank you
my parents, Shuqing Li and Meie Tang, for your endless love and encouragement. I could not
reach this far without your support and understanding. Thank you my husband, Zhengfei Lu, for
your support and company in all these years. Thank you for taking care of Ryan and me. You
and Ryan make every day of my life filled with love and happiness.

iv

Table of Contents
Dedication ...................................................................................................................................... ii
Acknowledgements ...................................................................................................................... iii
List of Tables ............................................................................................................................... vii
List of Figures ............................................................................................................................. viii
List of Abbreviations .................................................................................................................... x
Abstract ........................................................................................................................................ xii
Chapter 1. Introduction ............................................................................................................... 1
1.1 Cancer Immunotherapy .................................................................................................... 2
1.2 Immune checkpoints ........................................................................................................ 5
1.2.1 PD-1 and its signaling ............................................................................................... 5
1.2.2 PD-1/PD-L1 blockade in cancer therapy ................................................................ 10
1.2.3 Safety of PD-1/PD-L1 blockade ............................................................................. 14
1.3 Adoptive T cell transfer (ACT) ...................................................................................... 15
1.3.1 Chimeric Antigen Receptor (CAR) ........................................................................ 16
1.3.2 CAR in cancer therapy ............................................................................................ 16
1.3.3 Safety of CAR-T cell therapy ................................................................................. 21
1.4 Combination therapy ...................................................................................................... 22
1.4.1 Combining immunotherapies .................................................................................. 23
1.4.2 Combining immunotherapy and chemotherapy ...................................................... 23
1.5 Summary and Thesis work ............................................................................................. 24
Chapter 2. Anti-PD-1 antibody Enhances Antitumor Immunity of CAR-modified T cells in
B-cell Lymphoma ........................................................................................................................ 26
2.1 Abstract ............................................................................................................................... 27
2.2 Introduction ......................................................................................................................... 28
2.3 Materials and methods ........................................................................................................ 29
2.4 Results ................................................................................................................................. 34
2.4.1 Expression and activation of 1D3-28Z.1-3 anti-CD19 CAR in mouse splenic T cells 34
2.4.2 PD-1 expression is upregulated on anti-CD19-CAR-transduced T cells following
antigen-specific stimulation .................................................................................................. 36
2.4.3 PD-1 blockade enhances the antigen-specific immune responses of 1D3-28Z.1-3 CAR
T cells .................................................................................................................................... 37
2.4.4 PD-1 blockade enhances 1D3-28Z.1-3 CAR T cell-mediated tumor regression in
established tumor model ....................................................................................................... 39

v

2.4.5 The antitumor efficacy of combined therapy is associated with tumor
microenvironment modulation .............................................................................................. 41
2.4.6 Combination therapy specifically regulates local immune responses within tumor .... 42
2.5 Discussion ........................................................................................................................... 44
2.6 Conclusion .......................................................................................................................... 47
Chapter 3. Enhanced Cancer Immunotherapy by Chimeric Antigen Receptor-Modified T
Cells Engineered to Secrete Checkpoint Inhibitors ................................................................. 48
3.1 Abstract ............................................................................................................................... 49
3.2 Introduction ......................................................................................................................... 50
3.3 Materials and methods ........................................................................................................ 52
3.4 Results ................................................................................................................................. 59
3.4.1 Characterization of anti-CD19 CAR-T cells secreting anti-PD-1 antibody ................ 59
3.4.2 Secreting anti-PD-1 antibody enhances the antigen-specific immune responses of
CAR-T cells .......................................................................................................................... 61
3.4.3 Secreting anti-PD-1 alleviates CAR T cell exhaustion after antigen stimulation ........ 64
3.4.4 Anti-PD-1 engineered CAR T cells exhibit enhanced antitumor reactivity ................ 67
3.4.5 Anti-PD-1 engineered CAR T cells can expand more in vivo than parental CAR T
cells ....................................................................................................................................... 69
3.4.6 Anti-PD-1 engineered CAR T cells lead to reversal of T cell exhaustion and higher T
cell effector function at the established tumor site ............................................................... 70
3.5 Discussion ........................................................................................................................... 73
3.6 Conclusion .......................................................................................................................... 79
Chapter 4. Inhibition of Stat3 Enhances Antitumor Activity of PD-L1 Blockade in a
Murine Melanoma Model ........................................................................................................... 80
4.1 Abstract ............................................................................................................................... 81
4.2 Introduction ......................................................................................................................... 82
4.3 Materials and methods ........................................................................................................ 85
4.4 Results ................................................................................................................................. 89
4.4.1 PD-L1 blockade enhances Stat3 phosphorylation ....................................................... 89
4.4.2 BP-1-102 inhibits Stat3 activation and induces apoptotic cell death in murine
melanoma cells ...................................................................................................................... 89
4.4.3 BP-1-102 enhances the infiltration of mature DCs and cytotoxic CD8
+
T cells to local
tumor tissue ........................................................................................................................... 92
4.4.4 BP-1-102 enhances PD-L1 blockade-mediated tumor regression ............................... 92
4.4.5 Combination therapy modulates the infiltration of immune cells into tumor tissues .. 95
4.4.6 Combination therapy improves local immune responses at molecular level ............... 97

vi

4.5 Discussion ........................................................................................................................... 98
4.6 Conclusion ........................................................................................................................ 102
Chapter 5. Conclusion and Future perspectives .................................................................... 103
References .................................................................................................................................. 108

vii

List of Tables
Chapter 1
Table 1-1 Tumor-associated antigens in different tumor types.
...............................................
3

Table 1-2 Summary of current PD-1 and PD-L1 inhibitors in clinical trials.
.......................
14

Table 1-3 Therapeutic targets in treating solid tumors with CAR-T cells.
...........................
20

viii

List of Figures
Chapter 1
Figure 1-1 Multiple co-inhibitory interactions regulate T-cell responses.
...............................
6

Figure 1-2 Programmed death-1 (PD-1) signaling.
....................................................................
7

Figure 1-3 Mechanisms of PD-1 signaling-induced immunosuppression in the tumor
microenvironment.
.......................................................................................................................
9

Figure 1-4 Tumor adaptive resistance model through PD-1/PD-L1 engagement.
................
10

Figure 1-5 Building blocks of chimeric antigen receptors (CARs).
........................................
17

Figure 1-6 Overview of CAR-T cell therapy in clinic.
.............................................................
18

Chapter 2
Figure 2-1 Expression and activation of 1D3-28Z.1-3 anti-CD19 CAR in mouse splenic T
cells.
.............................................................................................................................................
35

Figure 2-2 Upregulation of PD-1 expression on CAR T cells following antigen-specific
stimulation.
..................................................................................................................................
36

Figure 2-3 Anti-PD-1 enhanced the antigen-specific immune responses of CAR T cells.
....
38

Figure 2-4 PD-1 blockade enhanced CAR19 T cells mediated tumor regression of
established tumor.
......................................................................................................................
40

Figure 2-5 The enhanced antitumor efficacy of combined therapy is correlated with tumor
microenvironment modulation.
.................................................................................................
42

Figure 2-6 Combined therapy specifically regulates immune responses at local tumor site.

......................................................................................................................................................
43

Chapter 3
Figure 3-1 Construction and characterization of CAR19 and CAR19.αPD1.
.......................
60

Figure 3-2 Anti-PD-1 expression enhanced the antigen-specific immune responses of CAR
T cells.
..........................................................................................................................................
63

Figure 3-3 Secreting anti-PD-1 scFv protected CAR T cells from being exhausted.
............
66

Figure 3-4 Adoptive transfer of CAR T cells secreting anti-PD-1 scFv enhanced the growth
inhibition of established tumor.
.................................................................................................
68

Figure 3-5 CAR T cells secreting anti-PD-1 were expanded more efficiently than parental
CAR T cells in vivo.
....................................................................................................................
70

Figure 3-6 CAR T cells secreting anti-PD-1 were more functional than parental CAR T
cells at local tumor site.
..............................................................................................................
72

Chapter 4
Figure 4-1 PD-L1 blockade increases Stat3 phosphorylation.
................................................
90

Figure 4-2 BP-1-102 inhibits Stat3 phosphorylation and induces apoptotic cell death.
.......
91

Figure 4-3 BP-1-102 modulates the tumor microenvironment.
..............................................
93

ix

Figure 4-4 BP-1-102 enhances the antitumor efficacy of PD-L1 blockade in B16-F10 murine
melanoma tumor model.
............................................................................................................
94

Figure 4-5 The combined therapy of anti-PD-L1 antibody and BP-1-102 enhanced tumor-
infiltrating T cells and decreased regulatory T cells within tumor.
.......................................
96

Figure 4-6 The combined therapy altered the tumor immune microenvironment.
..............
98

x

List of Abbreviations
ACT Adoptive T Cell Transfer
AML Acute Myeloid Leukemia
CAR Chimeric Antigen Receptor
CAIX Carbonic Anhydrase IX
CEA Carcinoembryonic Antigen
CLL Chronic Lymphocytic Leukemia
CPIs Immune Checkpoint Inhibitors
CRC Colorectal Cancer
CR Complete Response
CTLA-4 Cytotoxic T Lymphocyte-associated Antigen-4
FAP Fibroblast Activation Protein
FRα α-Folate Receptor
GM-CSF Granulocyte Macrophage Colony-stimulating Factor
HL Hodgkin’s Lymphoma
HER2 Human Epidermal Growth Factor Receptor 2
IFN-γ Interferon-gamma
IL-1β Interleukin-1 beta
IL-2 Interleukin-2
IL-4 Interleukin-4
irAE immune-related Adverse Event
ITIM Immunoreceptor Tyrosine-based Inhibitory Motif
ITSM Immunoreceptor Tyrosine-based Switch Motif
LAG-3 Lymphocyte-activation Gene-3

xi

LPS Lipopolysaccharide
L1-CAM L1-Cell Adhesion Molecule
MM Multiple Myeloma
NHL Non-Hodgkin’s Lymphoma
NSCLC Non-Small Cell Lung Cancer
ORR Objective Response Rate
PD-1 Programmed Cell Death-1
PD-L1 Programmed Death-ligand 1
PKA Protein Kinase A
PSMA Prostate-specific Membrane Antigen
RCC Renal Cell Carcinoma
STAT3 Signal Transducer and Activator of Transcription 3
scFv single chain variable Fragment
TAA Tumor-associated Antigen
TCR T Cell Receptor
TIL Tumor-infiltrated Lymphocyte
TIM-3 T-cell Immunoglobulin and Mucin-domain Containing-3
TNF-α Tumor Necrosis Factor-alpha
TRAIL TNF-related Apoptosis-induced Ligand
Treg Regulatory T Cell
VEGF Vascular Endothelial Growth Factor

xii

Abstract
Chimeric antigen receptor (CAR) T-cell based therapy has shown promise as an
immunotherapeutic modality for cancers. It has achieved successful responses in patients with
certain hematopoietic malignancies. However, the outcome has been less promising in the
treatment of lymphoma or solid tumors, partly owing to the immunosuppressive properties and
establishment of an immunosuppressive microenvironment. The PD-1/PD-L1 regulatory
pathway has demonstrated particularly antagonistic effects on the antitumor response of TILs. As
a result, we were trying to investigate the effects of PD-1/PD-L1 blockade on modulating the
tumor microenvironment and the function of infused CAR T cells.
In my first study, to overcome the limitation of CAR-T cells in the suppressive tumor
microenvironment, we combined CAR-T cells with anti-PD-1 antibody and evaluated the
antitumor efficacy in a murine lymphoma model. We observed that PD-1 blockade significantly
enhanced CAR T cell expansion and its effector function in vitro. In the animal study, we
demonstrated that the anti-PD-1 antibody significantly enhanced the antitumor activity of CAR-
T cells and prolonged the overall survival.
Though anti-PD-1 antibody significantly enhanced antitumor immunity of CAR-T cells,
antibody treatment has several limitations. For example, it requires multiple and continuous
antibody administration to obtain a sustained efficacy. Also, the large size of antibodies prevents
them from entering the tumor mass and encountering the infiltrated PD-1-positive T cells. To
account for these inefficiencies, multiple high-dose treatments with immunomodulatory drugs or
antibodies are required, but this can result in side effects that range from mild diarrhea to
autoimmune hepatitis, pneumonitis and colitis. Moreover, it has been shown that the Fc portion
of antibodies may cause immune cell depletion by activating cytotoxic signals within

xiii

macrophages and natural killer cells, which usually express FcαRI and FcγRIIIA/FcγRIIC,
respectively.
Therefore, in my second study, we focused our efforts on engineering CAR T cells to secrete and
deliver high concentrations of human scFvs against PD-1 (CAR.αPD1-T), aiming to change the
immunosuppressive tumor microenvironment, prevent tumor-induced hypofunctionality and
enhance the antitumor immunity of infused CAR T cells. We demonstrated for the first time that
PD-1 blockade by continuously secreted anti-PD-1 prevented T cell exhaustion and significantly
enhanced T cell expansion and effector function both in vitro and in vivo. In the xenograft mouse
model, we found that the secretion of anti-PD-1 enhanced the antitumor activity of CAR-T cells
and prolonged overall survival. Collectively, our study presents an important and novel strategy
that enables CAR-T cells to achieve better antitumor immunity, especially in the treatment of
solid tumors.
In order to achieve the best therapeutic efficacy, other than combining immunotherapies, a well-
designed combination of chemotherapy and immunotherapy may also be capable of improving
the response rates, especially for patients with refractory tumors. For example, clinical trials with
anti-PD-1 or anti-PD-L1 monoclonal antibodies alone have shown notable responses in patients
with metastatic melanoma. However, the overall response rate is still low. Therefore, seeking
combination therapy with enhanced efficacy of tumor eradication mediated by immune
checkpoint blockade remains an urgent task. In the present study, we showed that PD-L1
blockade significantly increased Stat3 phosphorylation, a critical factor shown to drive tumor
progression and immune evasion, including modulation of tumor-mediated immune suppression.
By combining BP-1-102, a Stat3 inhibitor, and anti-PD-L1 antibody, we demonstrated a
significant inhibition of tumor growth, compared to either treatment alone. Taken together, the

xiv

current study suggests that Stat3 inhibition by the small-molecule inhibitor BP-1-102 can
markedly enhances the antitumor potency of PD-L1 blockade, improving antitumor immunity
through the enhancement of both tumor microenvironment modulation and local immune
responses.

1

Chapter 1. Introduction

2

1.1 Cancer Immunotherapy
Over the past years, immunotherapy has evolved and become a novel promising modality for
cancer therapy. Unlike the other therapeutic strategies, immunotherapy is a type of cancer
treatment designed to boost, improve and/or restore a person’s own immune system to fight the
cancer (Li et al., 2016). Normally, the immune system is capable to distinguish between body’s
own cells-self and foreign cells-nonself, and quickly launches attack on cells or organisms
carrying foreign markers. In the case of cancer, some of the antigens on the cell surface may
change when the normal cells develop into cancer cells, which makes the tumors similar to
transplanted allogeneic organs even though they usually share the same major histocompatibility
complex (MHC) alleles as the normal tissues (Carmi et al., 2015). The antigens, which are
typically overexpressed on the tumor cell surface, are known as tumor-associated antigens
(TAAs). So far, a number of TAAs developed from different cancers have been identified
(Table 1-1) (Scott et al., 2012). Like infectious agents, the immune system can also attack tumor
cells by recognizing TAAs. For example, mice study has shown that the establishment of
allogeneic tumors can induce the production of tumor-binding IgG antibodies, which enable
dendritic cells (DCs) to internalize tumor antigens and subsequently activate tumor-reactive T
cells (Carmi et al., 2015). More importantly, T cells are also capable of recognizing and
unleashing attack on autologous cells that display tumor antigens (Muul et al., 1987; Rosenberg
et al., 1986). Interestingly, recent research has shown that in addition to T cell, NK cell also can
be an important factor to kill cancer cells (Cheng et al., 2013; Rezvani and Rouce, 2015). Most
of the time, this tumor-eradicating immunity confers to the host the ability to suppress tumor
development. This role of immune system that protects the host against the development of
cancers of non-viral origin is called cancer immunosurveillance, which was firstly proposed by

3

Burnet and Thomas in1950s (Burnet, 1957; Thomas, 1959). However, other times, when the
immune system breaks down or is overwhelmed, the cancer cells may slip through the defenses,
which is also known as immune evasion (Seliger, 2005). In order to improve or enhance the
immune system and make it better at recognizing and eliminating the tumor cells, a number of
immunotherapeutic approaches have been proposed and studied, including vaccines, cytokines,
monoclonal antibodies, checkpoint inhibitors, and adoptive T-cell transfer (ACT).

CAIX, carbonic anhydrase IX; CEA, CAIX, carbonic anhydrase IX; CEA, carcinoembryonic antigen; EGFR,
epidermal growth factor receptor; EpCAM, epithelial cell adhesion molecule; EPHA3, ephrin receptor A3; FAP,
fibroblast activation protein; gpA33, glycoprotein A33; IGF1R, insulin-like growth factor 1 receptor; Le
y
, Lewis Y
antigen; mAbs, monoclonal antibodies; PSMA, prostate-specific membrane antigen; RANKL, receptor activator of
nuclear factor-κB ligand; TAG-72, tumour-associated glycoprotein 72; TRAILR, tumour necrosis factor-related
apoptosis-inducing ligand receptor; VEGF, vascular endothelial growth factor; VEGFR, VEGF receptor.
Table 1-1 Tumor-associated antigens in different tumor types.
(Scott AM (2012) Nat Rev Cancer. 12: 278-287)

Therapeutic cancer vaccines, may administered as peptides, proteins, or tumor-antigen-pulsed
DCs with adjuvants, are designed to specifically attack and eradicate tumor cells through
boosting patient’s own immune system (Guo et al., 2013). In contrast to the traditional vaccines,

4

which are usually preventative and used to protect the body against foreign invaders, therapeutic
cancer vaccines aim to stimulate and strengthen the body’s natural immune response to fight
cancer. So far, several cancer treatment vaccines have been approved by the FDA. For example,
in 2010, sipuleucel-T was approved for the treatment of metastatic prostate cancer. Recently, in
2015, talimogene laherparepvec, the first oncolytic virus therapy, was also approved for the
treatment of metastatic melanoma.
Cytokines are a unique class of intercellular regulatory proteins that play a crucial role in
initiating, maintaining, and regulating immunologic homeostatic and inflammatory process
(Cutler and Brombacher, 2005). The first cytokine used for cancer therapy is interleukin-2 (IL-2).
The administration of IL-2 leads to durable, completed, and apparently curative regressions in
patients with metastatic melanoma and renal cancer (Rosenberg, 2014). Unfortunately, the
clinical use of IL-2 is limited due to its serious toxic effects at the dosage needed to sufficiently
elicit antitumor T cell responses. Later, a lot of other cytokines have also been studied and tested
in clinic, the overall response rate, however, is unimpressive (Abbas, 2011).
Antibodies are a type of proteins that can specifically recognize and bind to antigens expressed
on the surface of tumor cells. Tumor-specific monoclonal antibodies are capable to bind to TAAs
and elicit direct or indirect immune responses to destroy the tumor cells (Scott et al., 2012). The
mechanism of tumor cell killing mediated by antibodies summarized by Andrew Scott et al.
consists of direct action of the antibody (through blocking signaling pathways necessary for
tumor cell growth or survival), boost immune responses (through triggering an immune-mediated
cytotoxic response), and inhibit angiogenesis (Scott et al., 2012). So far, several monoclonal
antibodies have been approved by the FDA for cancer immunotherapy, including anti-CD52
antibody for CLL treatment, anti-CD20 antibody for metastatic melanoma, NHL and CLL

5

treatment, anti-CTLA-4 antibody for metastatic melanoma, anti-PD-1 antibody for metastatic
melanoma and unresectable or metastatic squamous non-small cell lung cancer, and anti-PD-L1
antibody for bladder cancer treatment. Among these approved therapeutic antibodies, more than
half of them are immune checkpoint inhibitors.
1.2 Immune checkpoints
The activity of immune system is mainly regulated by T cells. In T cell biology, the ultimate
amplitude and quality of the T cell response, which is initiated through TCR recognition of
specific antigen, is regulated by a balance between co- stimulatory and inhibitory signals (Pardoll,
2012). The primary role of immune checkpoints is to negatively regulate immune system and
maintain self-tolerance, protecting tissues from autoimmune attack during inflammatory
responses. However, in the case of cancer, this translates into a major immune resistance
mechanism (Pardoll, 2012). So far, several immune checkpoints have been identified, including
cytotoxic T-lymphocyte-associated antigen-4 (CTLA-4), lymphocyte-activation gene-3 (LAG-3),
T-cell immunoglobulin and mucin-domain containing-3 (TIM-3), and programmed cell death-1
(PD-1). Among them, CTLA-4 and PD-1 are well studied targets in cancer therapy (Figure 1-1)
(Pardoll, 2012).
1.2.1 PD-1 and its signaling
PD-1, also known as CD279, is a type I trans-membrane protein consisting of an extracellular
immunoglobulin superfamily domain, a transmembrane domain, and an intracellular domain
containing an immune receptor tyrosine-based inhibitory motif (ITIM) and an immunoreceptor
tyrosine-based switch motif (ITSM) (Ceeraz et al., 2013). PD-1 is one of the most important co-

6

Figure 1-1 Multiple co-inhibitory interactions regulate T-cell responses.
(Freeman G. J. (2012) Nat Immunol. 13(2): 113-115)

inhibitory receptors in T cells and highly expressed in activated T cells, B cells, and myeloid
cells (Freeman et al., 2000). It functions as an immune checkpoint and is primarily believed to
limit the activity of T cells by binding to its cognate ligands. Normally, when PD-1 binds to its
ligand, it can activate the intracellular phosphatase SHP
2
, which will inhibit kinase-signaling
pathways involved in T-cell activation, proliferation and apoptosis (Figure 1-2). This process
prevents over activation during inflammatory responses and protects tissues from autoimmune
attack. However, in the case of cancer, this translates into a major immune resistance mechanism
(Pardoll, 2012). Unlike the other inhibitory receptors such as CTLA-4, which usually inhibits
early T-cell activation. PD-1 primarily inhibits T-cell activity in the effector phase within tissues
and tumors (Santarpia and Karachaliou, 2015). It has been shown that the expression of PD-1 is
upregulated shortly when T cell is activated. Additionally, in the tumor microenvironment, PD-1
expression on tumor infiltrating T lymphocytes (TILs) has been shown to be significantly higher
compared to that in normal tissue T-cell infiltrates and peripheral blood T lymphocytes, which

7

Figure 1-2 Programmed death-1 (PD-1) signaling.
(Jordan M. C. (2015) Trends in Pharmacological Sciences. 36: 587-595)

indicates that the tumor microenvironment can lead to upregulation of PD-1 on tumor-reactive T
cells and can contribute to impaired antitumor immune responses (Ahmed et al., 2015; Bonifant
et al., 2016). In addition, PD-1 is also expressed on regulatory T cells (Tregs) and capable to
promote the proliferation of Treg and inhibit T cell responses (Francisco et al., 2009).
The two ligands for PD-1 are PD-L1 and PD-L2. PD-L1 is constitutively expressed on resting T
cells, B cells, macrophages, and dendritic cells (DCs) (Yamazaki et al., 2002). In addition to
hematopoietic lineage cells, PD-L1 is also shown abundantly in a wide variety of solid tumors,
including melanoma, ovarian cancer, renal cell carcinoma, gastric cancer, lung cancer, various
leukemias and so on (Brown et al., 2003; Dong et al., 2002; Konishi et al., 2004). In contrast, the
expression of PD-L1 in normal tissues is undetectable (Dong et al., 2002). In comparison, PD-L2
expression is more restricted on macrophages and dendritic cells and less prevalent than PD-L1
across different tumors (He et al., 2015). The restricted expression patterns of PD-L2 determines

8

that the primary role of PD-L2 is to regulate T-cell priming or polarization, whereas the broad
distribution of PD-L1 implies a more general role in protecting peripheral tissues from excessive
inflammation (He et al., 2015).
It has been shown that PD-1/PD-L1 signaling induces immune suppression in multiple ways,
including mediating apoptosis of activated T cells, stimulating IL-10 production in human
peripheral blood T cells, inducing T cell anergy and exhaustion, increasing Treg function, and
inhibiting T cell proliferation (Figure 1-3) (Chen and Han, 2015). It has shown that the
expression of PD-L1 would promote CD8
+
T cells apoptosis and blocking PD-L1 could reverse
this effect, suggesting that PD-1/PD-L1 pathway may play an important role in inducing T cell
apoptosis (Thomas and Massague, 2005). T cell anergy refers to the hyporesponsive state of T
cells with low IL-2 production or incomplete activation, which is usually caused by low co-
stimulatory and/or high co-inhibitory stimulation (Crespo et al., 2013). The exact molecular
mechanism controlling T cell anergy is still not clear, however, many studies have shown that
PD-1/PD-L1 signaling inhibits the effector phase of CD8
+
T cells and blockade of PD-L1
improves tumor-specific T cell responses (Blank et al., 2004; Blank et al., 2006). Tregs are
involved in mediating peripheral tolerance by actively suppressing effector T cells and inhibiting
immune-mediated tissue damage (Francisco et al., 2010). In the progression of cancer, Tregs also
play an active and significant role in suppressing tumor-specific immunity. The ligation of PD-1
with PD-L1 is capable to enhance Foxp3 expression and induce the conversion of naive CD4
+
T
cells to induced Tregs (iTregs) by attenuation of Akt-mTOR signaling and concomitant
upregulation of PTEN (Francisco et al., 2010). Additionally, PD-1 can also inhibit the induction

9

Figure 1-3 Mechanisms of PD-1 signaling-induced immunosuppression in the tumor
microenvironment.
(Chen L. (2015) J Clin Invest. 125(9): 3384-91)

of PI3K activity and downstream activation of Akt, the signaling of which is widely known to be
involved in cell proliferation. Thereby, PD-1 can also regulate T cell function by inhibiting cell
proliferation

(Blank et al., 2006).
The expression of PD-L1 can be induced by multiple pro-inflammatory mediators, including
IFN-γ, TNF-α, IL-2, LPS, GM-CSF, VEGF, and IL-4 (Curiel et al., 2003; Kondo et al., 2010;
Voron et al., 2015; Yamazaki et al., 2002). In particular, IFN-γ is a potent inducer of PD-L1
expression. In tumor microenvironment, the production of these inflammatory molecules may
promote immune suppression by upregulating PD-L1 expression, which is also called “adaptive
immune resistance” since the tumor acquires the ability to protect itself by inducing PD-L1 in
response to IFN-γ produced by activated T cells (Figure 1-4) (Chen and Han, 2015; He et al.,

10

2015). PD-L1 is also regulated by other proteins such as PTEN and Stat3. Studies have shown
that the frequently mutated PTEN in cancer cells would increase both the mRNA and protein
expression of PD-L1 by upregulating S6K1 gene and activating the PI3K/Akt downstream
mTOR-S6K1 signaling pathway (Crane et al., 2009; Parsa et al., 2007). In addition, it is also
found that Stat3 can bind to PD-L1 promoter and regulate its expression (Wolfle et al., 2011). It
is important to note that in addition to PD-1, PD-L1 can also bind to CD80, which is a co-
stimulatory receptor on T cell and does not transmit negative regulated signals (Butte et al.,
2007).
1.2.2 PD-1/PD-L1 blockade in cancer therapy
The findings of increased PD-1 expression on TILs and increased PD-L1 expression on tumor
cells have provided important rationale for blockade this pathway to enhance antitumor
immunity (Pardoll, 2012). To date, more than 200 clinical trials using antibodies targeting PD-1
pathway have been carried out in a variety of cancers, including both solid and hematological
tumors (Table 1-2) (Ribas et al., 2016).

Figure 1-4 Tumor adaptive resistance model through PD-1/PD-L1 engagement.
(Chen L. (2015) J Clin Invest. 125(9): 3384-91)

11

In 2008, the first phase I clinical trial of a humanized anti-PD-1 antibody, CT-001, was
conducted in 17 patients with advanced hematologic malignancies such as acute myeloid
leukemia (AML), chronic lymphocytic leukemia (CLL), non-Hodgkin’s lymphoma (NHL),
Hodgkin’s lymphoma (HL), and multiple myeloma (MM). The clinical benefit was observed in
33% of the patients with one complete remission. No severe adverse event was observed in this
study, so CT-011 was considered generally safe and well tolerated (Berger et al., 2008).
Subsequently, in 2010, a phase I study of MDX-1106, also known as nivolumab, was carried out
in 39 patients with advanced metastatic melanoma, colorectal cancer (CRC), castrate-resistant
prostate cancer, non-small-cell lung cancer (NSCLC) or renal cell carcinoma (RCC). In this
study, only one serious adverse event, inflammatory colitis, was observed and the overall
objective response rate (ORR) was 7.7%, including one durable complete response in CRC and
two partial responses in melanoma and RCC (Brahmer et al., 2010). Two years later, in 2012,
another phase I trial of nivolumab (BMS-936558) was reported. In this study, 296 patients with
selected advanced solid tumors including melanoma, lung cancer, and renal cell carcinoma,
received treatment. It shows that the ORR is 28% for melanoma, 18% for lung cancer, and 27%
for renal cell carcinoma. Overall, nivolumab was well tolerated with low-grade fatigue,
decreased appetite, diarrhea, nausea, cough, dyspnea, constipation, vomiting, rash, pyrexia, and
headache. Grade 3 or 4 treatment-related adverse events were observed in 41 of 296 patients
(14%) (Topalian et al., 2012). Long-term safety of nivolumab was further followed up in a study,
which involves 107 patients with advanced melanoma. It shows that median overall survival in
nivolumab-treated patients was 16.8 months, and 1- and 2-year survival rates were 62% and 43%,
respectively. The long-term safety was also acceptable (Topalian et al., 2014). In December 2014,
the FDA approved nivolumab for patients with unresectable or metastatic melanoma based on a

12

phase III clinical trial of 120 participants showing that 32% of subjects receiving nivolumab had
their tumor shrink. The effect lasts for more than 6 months in approximately one-third of the
participants who experienced tumor shrinkage (National Cancer Institute, 2015). Later, the FDA
subsequently approved nivolumab for NSCLC, RCC, classical HL, head and neck cancer, and
urothelial carcinoma (Institute, 2015).
MK-3475, also known as pembrolizumab, is a highly selective, humanized monoclonal IgG4
against PD-1. The first clinical trial was published in 2013 to evaluate the safety and tumor
responses of MK-3475. It revealed that of the 135 participants, 79% reported drug-related
adverse events of any grade, and 13% reported grade 3 or 4 drug-related adverse events.
Generalized symptoms, including fatigue and asthenia, fever, myalgias, and headaches, were
reported frequently but were of low grade in more than 95% of the cases. Grade 3 or 4 pruritus
was reported in 1% of the patients, and grade 3 or 4 rash in 2%. Therefore, pembrolizumab is
considered to be well tolerated. The overall ORR during receipt of therapy was 37% (Hamid et
al., 2013). Subsequently, pembrolizumab was evaluated as a treatment for metastatic melanoma
in three major clinical trials: KEYNOTE-001 (in patients who were ipilimumab-naïve or
pretreated), KEYNOTE-002 (in patients who were ipilimumab pretreated) and KEYNOTE-006
(in patients who were ipilimumab naïve)

(Boutros et al., 2016). The results from KEYNOTE-001
phase I clinical trial showed that 34% of the cohort of 655 patients had an objective response to
pembrolizumab, with 6% having a complete response. On the basis of these results, in 2014, the
FDA approved pembrolizumab to be used in treatment for metastatic melanoma with non BRAF-
mutant. Later on, pembrolizumab was also approved to be used in treatment for NSCLC, head
and neck cancer and HL.

13

AMP-224, a PD-L2 Fc fusion protein, binds specifically to PD-1
HI
T cells. A recent clinical
study of AMP-224 has been conducted in 17 patients with refractory metastatic CRC. However,
the trial is still ongoing and no objective responses have been released yet (Austin G. Duffy,
2016).
In addition to anti-PD-1 antibody, to block PD-1/PD-L1 axis, an alternative approach is to block
the ligand, PD-L1. Thus far, four anti-PD-L1 monoclonal antibodies (MEDI4736, BMS-936559,
MPDL3280A, MSB001071C) are in clinical trial to evaluate their safety and antitumor efficacy.
MEDI4736, also known as atezolizumab, has been approved by the FDA for the treatment of
patients with urothelial carcinoma or NSCLC in 2016 (Institute, 2016; www.FDA.gov, 2016).
The safety and efficacy of atezolizumab was studied in a clinical trial, which involves 310
participants. It showed that 14.8% of patients experienced tumor shrink. By classifying the
expression of PD-L1 in tumor-infiltrating immune cells, it was found that 26% of the participants
that expressed PD-L1 had a tumor response, whereas only 9.5% of participants who were
classified as “negative” for PD-L1 expression showed responses (www.FDA.gov, 2016). In
addition, atezolizumab has been well tolerated due to the fact that most of the side effects are
very low-grade (www.FDA.gov, 2016). BMS-936559, is a high-affinity, fully human, PD-L1-
specific, IgG4 monoclonal antibody. It has been evaluated in 207 patients with advanced solid
tumors, including NSCLC, melanoma, colorectal cancer, RCC, and ovarian cancer, showing that
the overall ORR was observed in melanoma (17%), NSCLC (8-11%), Ovarian cancer (6%), and
RCC (12%) (Brahmer et al., 2010). The safety and efficacy of MPDL3280A, a human
monoclonal antibody targeting PD-L1, was evaluated in a phase I clinical trial, showing an ORR
of 21% in nonselected solid tumors, including NSCLC, RCC, melanoma, CRC, and gastric
cancer. The safety profile of MPDL3280A showed that it could be well tolerated, with no

14

pneumonitis-related deaths (Roy S. Herbst, 2013). MSB0010718C, a fully human IgG1
monoclonal antibody targeting PD-L1, was also evaluated in a phase I open-label, multiple
ascending dose trial. The trial is still under evaluation and no safety date or objective responses
have been released yet (Christopher Ryan Heery, 2014).

Table 1-2 Summary of current PD-1 and PD-L1 inhibitors in clinical trials.

1.2.3 Safety of PD-1/PD-L1 blockade
In preclinical study, it shows that the phenotype of PD-1 or PD-L1 mice is mild. For example,
different from CTLA-4, the genetic ablation of which would lead to profound
lymphoproliferation and death at 3-4 weeks of age, PD-1 knockout mice develop tissue- and
strain-specific autoimmunity at approximately 9 months of age, and PD-L1 knockout mice
display virtually no phenotype (Quezada and Peggs, 2013). In the clinical trials, the adverse
events of utilizing PD-1 or PD-L1 inhibitors are mostly immune-related (irAEs). For example,

15

irAEs occur in 70% of patients treated with a PD-1/PD-L1 antibody (Michot et al., 2016). These
adverse events are mostly grade I or II that mainly affects the skin (43-45% of patients) and the
gastrointestinal tract (29-32% of patients) (Boutros et al., 2016). Grade III or V irAEs are barely
observed in patients treated with PD-1/PD-L1 antibodies, whereas, 9-15% was seen in patients
treated with anti-CTLA-4 antibodies. The majority of these irAEs occurs within 3-6 months of
the initiation of anti-PD-1 or anti-PD-1 treatment (Michot et al., 2016).

1.3 Adoptive T cell transfer (ACT)
T cells are believed to play a major role in immunosurveillance and tumor eradication. Based on
this paradigm, efforts have been made to generate, educate, and/or enhance T cells against
tumors. Since the group of Steve Rosenberg pioneered the isolation and expansion of tumor-
infiltrating T lymphocytes (TILs), and showed that patient-derived T cells could be grown and
selected in culture, and then infused back into patients for the treatment of advanced melanoma,
T-cell therapy has been becoming the most popular therapeutic modality in cancer
immunotherapy (Bonini and Mondino, 2015). It revealed that following lymphodepleting
chemotherapy, adoptive transfer of TILs could mediate objective tumor regression in 50-70% of
patients with metastatic melanoma, implying the potential of patient-derived TILs in the
treatment of solid tumors. However, even though tumor-specific lymphocytes can be recovered
from several solid tumors, poorer antigenicity, tumor growth at less-accessible sites, and
technical difficulties in obtaining sufficient numbers of cells limited the clinical application of
this strategy (Bonini and Mondino, 2015).

16

With the advances in gene transfer technology and insights in T-cell biology, to further exploit
alloreactive T cells, and improve the safety and efficacy of autologous ACT therapy, patient-
derived T cells have been engineered to express a T-cell receptor (TCR) or a chimeric antigen
receptor (CAR) specific for TAAs. TCRs and CARs confer to T cells the ability to recognize
specific TAAs and kill tumor cells via MHC-dependent (TCR) or MHC-independent (CAR)
mechanisms more efficiently (Bonini and Mondino, 2015).
1.3.1 Chimeric Antigen Receptor (CAR)
Typically, a CAR is genetically engineered on the T cell to enable tumor antigen recognition in a
MHC-independent manner. CARs are composed of an extracellular antigen-recognition domain,
a hinge, a transmembrane domain, and an intracellular signaling domain. The extracellular
ligand-binding domain is usually a single-chain variable fragment (scFv) derived from a
monoclonal antibody, which is usually designed to target a specific TAA (Figure 1-5). When the
CAR and its ligand are engaged, a signal will be transmitted into the intracellular T cell
machinery through a signaling domain, typically the CD3ζ chain of the T-cell receptor (TCR). A
CAR with CD3ζ as its endodomain is known as the “first generation”. To enhance T cell
proliferation and persistence, co-stimulatory receptors such as CD28, 4-1BB, or OX40 are
cooperated to the CD3ζ intracellular domain. CAR with one of the co-stimulatory receptors is
known as “second generation” and CAR with more than one co-stimulatory molecule is known
as “third generation” (Gill et al., 2016) (Figure 1-5).
1.3.2 CAR in cancer therapy
CAR-T cell therapy uses gene transfer technology to reprogram a patient’s T cells to express

17

Figure 1-5 Building blocks of chimeric antigen receptors (CARs).
(A) Designed monoclonal antibody recognizes specific TAA. (B) TCR and co-stimulatory
molecules of T cells. (C) Anatomy of CARs.

CAR, thereby directing T cells specifically against tumor cells (Figure 1-6) (Maus and June,
2016). Since 1989, when CAR was firstly reported by Gross et al. (Gross et al., 1989), a variety
of studies have been published, using CARs targeting various tumor antigens. In the past a few
years, a number of different CARs have been evaluated in clinic trials. So far, the studies have
provided important proof-of-principle demonstration to emphasize the potential of CAR-T cell
for cancer therapy, especially for advanced malignancies. For example, a trial of 16 ALL patients
treated with CAR-T cells targeting CD19 showed 88% of complete response (CR) (Brentjens et
al., 2013; Davila et al., 2014). Later on, more trials were conducted in the other blood
malignancies such as NHL, showing that anti-CD19 CAR-T cells were also capable to induce

18

more than 50% of CR (Kochenderfer et al., 2015). Encouraged by the results of CAR-T cell

Figure 1-6 Overview of CAR-T cell therapy in clinic.
(Marcela V. Maus, and Carl H. June Clin Cancer Res 2016;22:1875-1884)

therapy in blood-borne tumors, a growing number of studies have focused on solid tumors,
targeting surface proteins including carcinoembryonic antigen (CEA), the diganglioside GD2,
mesothelin, interleukin 13 receptor α (IL13Rα), human epidermal growth factor receptor 2
(HER2), fibroblast activation protein (FAP), epithelial growth factor receptor (EGFR), and L1
cell adhesion molecule (L1CAM) (Table 1-3) (Zhang et al., 2016). Unfortunately, despite
encouraging results have been achieved in blood malignancies, the antitumor efficacy of CAR-T
cell therapy in solid tumors is less inspiring (Newick et al., 2016a). For example, two trials using

19

CAR-T cells specifically targeting α-folate receptor or CAIX for ovarian cancer patients or renal
cell carcinoma patients, respectively, failed to achieve effective antitumor responses (Kershaw et
al., 2006; Lamers et al., 2006).

Unlike the “liquid tumor”, the solid tumor has a highly
immunosuppressive microenvironment, which is extremely inhospitable to CAR-T cells. To
achieve effective antitumor activity, the CAR-T cells firstly has to infiltrate into the tumor in
order to get access to the tumor cells and elicit TAA-specific cytotoxicity. Even after successful
trafficking, T cells still have to overcome various challenges, including harsh physical and
metabolic barriers, highly immunosuppressive tumor microenvironment, and tumor-induced T
cell exhaustion. The solid tumor presents a markedly stressful and inhospitable
microenvironment toward T cells, including high tissue pressure, hypoxia, and nutrient starvation,
which would inhibit T-cell extravasation, proliferation, and cytokine production. It has been
shown that reducing tumor fibroblast numbers using FAP-CAR-T cells or having CAR-T cell
secret an enzyme that degrades matrix could augment CAR-T cell function in animal models
(Newick et al., 2016a). Furthermore, as stated before, T cell itself expresses a number of
inhibitory receptors in order to prevent autoimmunity in the cases of inflammation or chronic
disease. In the case of tumor, it takes advantage of this characterize and overexpress ligands such
as PD-L1 to inhibit T-cell function and avoid tumor surveillance. In addition, in the tumor
microenvironment, the presence of suppressive factors, cytokines and immune cells, may limit
the efficacy of CAR-T cell therapy.
The trafficking of CAR-T cells to the tumor microenvironment is essential for the success of
cancer immunotherapy. In general, T-cell trafficking to the tumor is regulated by a series of
distinct processes, involving attachment/adhesion, rolling/tethering, chemotaxis, and

20

extravasation (Slaney et al., 2014; Zhang et al., 2016). When the naïve T cells get activated, they
will become effector T cells and gain expression of a number of homing molecules that confer

FRα, α-folate receptor; L1-CAM, L1-cell adhesion molecule; CAIX, carbonic anhydrase IX; FAP, Fibroblast
activation protein; HER2, human epidermal growth factor receptor 2; CEA, carcinoembryonic antigen; PSMA,
prostate-specific membrane antigen; CEA, Carcino Embryonie Antigen.
Table 1-3 Therapeutic targets in treating solid tumors with CAR-T cells.
(Zhang H (2016) Int J Biol Sci. 12(6): 718-729)
them to migrate to lesions, such as cancer. These molecules include ligand for E- and P-selectin,
and chemokine receptors such as CXCR3 and CXCR6. The mismatch of chemokine-chemokine
receptor pairs has been shown to contribute to the poor homing of CAR-T cells into the tumor
(Slaney et al., 2014; Zhang et al., 2016). Other than that, the downregulation of adhesion
molecules and aberrant vasculature may also play important roles. It has been reported that the
low expression of some critical chemokines, such as CXCL9 and CXCL10 (ligands for CXCR3),

21

by tumor cells may limit the infiltration of effector T cells, resulting in poor antitumor immunity.
Increasing the expression of CXCL9 and CXCL10 on tumor cells resulted in an increased
infiltration of CXCR3
+
effector T cells and an enhanced antitumor response (Hong et al., 2011).
Therefore, to overcome this problem, one approach is to design CAR-T cells that can coexpress
“better matched” chemokine receptors. For example, it has been shown that mesothelioma
tumors overexpress CCL2. Therefore, to engineer mesothelin CAR-T cells and make it coexpress
CCR2b has resulted in enhanced intra-tumoral migration of CAR-T cells and antitumor activity
(Moon et al., 2011). More importantly, people found that PD-1 blockade was capable to enhance
T-cell migration to tumors by elevating IFN-γ inducible chemokines, including CXCL10,
thereby inspiring researchers to focus on combining PD-1 blockade and CAR-T cell therapy
trying to achieve better antitumor activity. In addition, it has also been shown that the expression
of protein kinase A (PKA) is capable to cause poor T-cell trafficking by interacting with ezrin
and inhibiting T-cell receptor (TCR) activation. To engineer the CAR-T cells and make them
express a small peptide that inhibits the association of PKA and ezrin, the CAR-T cells revealed
enhanced antitumor activity due to increased T-cell infiltration of established tumors (Newick et
al., 2016b).
1.3.3 Safety of CAR-T cell therapy
So far, based on the dramatic antitumor efficacy in blood malignancies, more than 110 CAR-T
cell clinical trials have been being conducted worldwide. Also, the US FDA is expected to
approve the first CAR-T cell therapy in 2017 (BOCK, 2016). Despite some promising results,
there are a number of side effects associated with CAR-T cell therapy. The most serious is
cytokine release syndrome (CRS). It has been shown that a potential life-threatening condition
can be mediated by the release of IL-6, TNF-α, IFN-γ following immune cell activation.

22

Fortunately, it reveals that the effects of CRS can be mitigated by reducing the number of
infused CAR T-cells and the use of anti-IL-6 receptor antibodies and steroids (Michot et al.,
2016). Another side effect associated with CAR-T cell therapy is B-cell aplasia when CAR-T
cell specifically targets B cell-specific marker CD19. To address this issue, people found that
infusion of gamma globulin, naturally-occurring antibodies that are required to provide
protective immunity could alleviate the effect of loss of B cells. In addition, for CAR-T cell-
targeting TAAs that are also expressed by normal tissues might have some on-target off-tumor
toxicity (Michot et al., 2016). In order to address this problem, our lab engineered and masked
the CAR and demonstrated that by masking the CARs it could improve the safety profile of
conventional CARs (Chen and Han, 2015).

1.4 Combination therapy
More and more oncologists realize that a one-size-fits-all approach to medicine is too simple to
achieve the optimal anti-tumor activity. Given the development of drug resistance, genetic
difference in patients, and the requirement of more effective but less toxic treatment options, it is
important to seek other therapeutic strategies as combination therapy upon present options
(ONCOSEC, 2013). Each drug or treatment method approved by the FDA for cancer therapy
must prove to be effective in inhibiting tumor growth or prolonging patient survival. However,
monotherapy may not always give the desired result. In order to achieve the best therapeutic
efficacy, large dosages are usually required, with subsequent drug resistances or toxicity. A well-
designed combination therapy may overcome these difficulties and improve the response rates.

23

1.4.1 Combining immunotherapies
Despite single immunotherapy has demonstrated clear anti-tumor activity across multiple tumor
types, the response rates are still low. In order to achieve a good therapeutic efficacy, large
dosages are usually utilized, with subsequent drug resistances or toxicity. For example, the
objective response rate (ORR) of ipilimumab (anti-CTLA-4 antibody) alone in advanced
melanoma is only 10.9%, and the ORR of nivolumab (anti-PD-1 antibody) alone in advanced
melanoma is 40%. However, to combine ipilimumab and nivolumab, at a much lower dosages
for both antibodies, the ORR can also achieve 40% (Callahan et al., 2014). Additionally, in
CAR-T cell based therapy, the trafficking of CAR-T cells to the tumor microenvironment is
essential for the success of cancer immunotherapy. People found that PD-1 blockade was capable
to enhance T-cell migration to tumors by elevating IFN-γ inducible chemokines, including
CXCL10. Therefore, in order to enhance the homing of CAR-T cells into tumors, the researchers
are focused on combining PD-1 blockade and CAR-T cell therapy together to achieve better
antitumor activity (Cherkassky et al., 2016; John et al., 2013).
1.4.2 Combining immunotherapy and chemotherapy
Conventionally, people thought that chemotherapy is a contra-event for immunotherapy as it
may cause lymphocyte depletion (lymphopenia) and the programmed cell death induced by
chemotherapy is tolerogenic. However, more than more evidence has demonstrated that a
number of immune cells killed by the chemotherapy may be beneficial rather than detrimental
for the cancer therapy over long-term treatment (van der Most et al., 2005). For example, the loss
of regulatory CD4
+
T cells caused by chemotherapy may benefit the anti-tumor T cell response
induced by immunotherapy (Ding and Zhou, 2012). In addition, it has been shown that
lymphopenia can trigger a phase of immune system regeneration (van der Most et al., 2005). In

24

addition, it has been shown that chemotherapy, especially targeted therapies, may also directly
regulate immune responses. For example, sunitinib, a multi-targeted receptor tyrosine kinase
(RTK) inhibitor, is shown to attenuate the activities of specific immune cell populations that
restrain cytotoxic T lymphocytes (CTLs), including FoxP3
+
regulatory T cells and myeloid-
derived suppressor cells (MDSCs) (Vanneman and Dranoff, 2012). Other studies also show that
the inhibition of Jak2/Stat3 augment tumor antigen presentation by DCs, thereby enhancing the
priming of tumor-specific immune responses of CTLs (Nefedova et al., 2005a; Nefedova et al.,
2005b). Some studies have been shown that the general response rate of tumor-specific
monoclonal antibody yield 8-10% in patients with advanced cancers or recurrent disease. When
combined with chemotherapy and/or radiotherapy, the response rate is shown to increase to 30%
(Kirkwood et al., 2012).
1.5 Summary and Thesis work
In this study, in order to overcome the highly immunosuppressive tumor microenvironment and
improve the potency of CAR T-cell based therapy in the treatment of solid tumors, we combined
the CAR T cells with immune checkpoint inhibitors (CPIs), aiming to change the
immunosuppressive tumor microenvironment, prevent tumor-induced hypofunctionality and
enhance the antitumor immunity of infused CAR T cells. Moreover, in order to achieve the best
therapeutic efficacy, especially for patients with refractory tumors, other than combining
immunotherapies, we also combined immunotherapy with chemotherapy and evaluate the
antitumor efficacy of combination therapy for the treatment of advanced melanoma.
In chapter 2, to overcome the limitation of CAR-T cells in the suppressive tumor
microenvironment, we combined CAR-T cells with anti-PD-1 antibody and evaluated the
antitumor efficacy in a murine lymphoma model. We observed that PD-1 blockade significantly

25

enhanced CAR T cell expansion and its effector function in vitro. In the animal study, we
demonstrated that the anti-PD-1 antibody significantly enhanced the antitumor activity of CAR-
T cells and prolonged overall survival.
In chapter 3, instead of combining CAR T cells with systemic injection of anti-PD-1 antibody,
we focused our efforts on engineering CAR T cells to secrete and deliver high concentrations of
human scFvs against PD-1 (CAR.αPD1). We demonstrated for the first time that PD-1 blockade
by continuously secreted anti-PD-1 prevented T cell exhaustion and significantly enhanced T cell
expansion and effector function both in vitro and in vivo. In the xenograft mouse model, we
found that the secretion of anti-PD-1 enhanced the antitumor activity of CAR-T cells and
prolonged overall survival.
In chapter 4, for the first time, we showed that in the treatment of melanoma cells, PD-L1
blockade significantly increased Stat3 phosphorylation, a critical factor shown to drive tumor
progression and immune evasion, including modulation of tumor-mediated immune suppression.
In order to overcome the resistance of melanoma cells to PD-L1 blockade, we combined BP-1-
102, a Stat3 inhibitor, and anti-PD-L1 antibody, and demonstrated a significant inhibition of
tumor growth, compared to either treatment alone.

26

Chapter 2. Anti-PD-1 antibody Enhances Antitumor Immunity of
CAR-modified T cells in B-cell Lymphoma

27

2.1 Abstract
Despite favorable responses of chimeric antigen receptor (CAR)-engineered T cell therapy in
patients with certain hematologic malignancies, the outcome has been far from satisfactory in the
treatment of solid tumors or lymphoma, partially owing to the development of an
immunosuppressive tumor microenvironment. To overcome this limitation, we combined CAR-
T cells with checkpoint inhibitor (CPI) anti-PD-1 antibody and evaluated the antitumor efficacy
in a murine lymphoma model. In an effort to evaluate the effector function and expansion
capacity of CAR T cells combining with PD-1 blockade in vitro, we measured the production of
IFN-γ and cell proliferation marker Ki-67 following antigen-specific stimulation. Furthermore,
the antitumor efficacy of CAR-T cells, anti-PD-1 antibody and CAR-T cells combined with anti-
PD-1 antibody was determined using a murine lymphoma model. Finally, the underlying
mechanism of enhanced tumor eradication of combination therapy was investigated by analyzing
the expansion and tumor infiltration of adoptively transferred T cells. We demonstrated that PD-
1 blockade significantly enhanced CAR T cell expansion and effector function in vitro. In the
animal study, we demonstrated that the anti-PD-1 antibody enhanced the antitumor activity of
CAR-T cells and prolonged overall survival. In conclusion, our study presents an important and
novel strategy that enables CAR-T cells to achieve better antitumor immunity, especially in the
treatment of tumors with suppressive microenvironment.

28

2.2 Introduction
The development of ex vivo culture technology and genetically engineered T cells has led to
rapid generation of chimeric antigen receptor (CAR) T cells, thereby broadening the applicability
of cancer immunotherapy (Brentjens et al., 2014; Hollyman et al., 2009; Somerville et al., 2012;
Vera et al., 2010). Typically, a CAR, consisting of an extracellular antigen-recognition domain, a
hinge, a transmembrane domain, and an intracellular signaling domain, is genetically engineered
on the T cell to enable tumor-associated antigen (TAA) recognition in a major histocompatibility
class (MHC)- independent manner (Ahmed et al., 2015).
So far, CAR-T cell therapy has consistently demonstrated significant antitumor capacity in
patients with acute lymphoblastic leukemia (ALL) (Maude et al., 2015). It shows that up to 90%
of children and adults with ALL, who had either relapsed or failed to respond to standard
therapies, achieved a complete remission after CAR-T cell therapy (Maude et al., 2014).
However, the outcome has been less inspiring in treatment of other hematological tumors such as
lymphoma or solid malignancies (Fousek and Ahmed, 2015; Gill et al., 2016; Maude et al.,
2014). For example, in recent clinical trials of non-Hodgkin lymphoma (NHL), only 33%
complete remission were achieved with CAR-T cell therapy (Helwick, 2016; Locke et al., 2015).
This can be attributed, in part, to the establishment of an immunosuppressive microenvironment
within the tumors. Such milieu involves the upregulation of a number of intrinsic inhibitory
pathways mediated by increased expression of inhibitory receptors (IRs) in T cells reacting with
their cognate ligands expressed on cancer cell surface (Pardoll, 2012; Yamazaki et al., 2002).
Programmed dearh-1 (PD-1) is one of the most important IRs characterized in T cells. Unlike
other IRs, PD-1 is upregulated shortly after T cell activation, which in turn, inhibits T cell
effector function via interacting with its two ligands, PD-L1 or PD-L2. PD-L1 is constitutively

29

expressed on T cells, B cells, macrophages, and dendritic cells (DCs) (Kondo et al., 2010). PD-
L1 is also shown to be abundantly expressed in a wide variety of solid tumors (Brown et al.,
2003; Dong et al., 2002; Konishi et al., 2004). In contrast, the expression of PD-L1 in normal
tissues is undetectable (Dong et al., 2002). As a consequence of its critical role in
immunosuppression, PD-1 has been the focus of recent research. It aims to neutralize PD-1’s
negative effect on T cells and enhance their antitumor responses. Clinical studies have
demonstrated that PD-1 blockade significantly enhanced tumor regression in colon, renal and
lung cancers and melanoma (Dong et al., 2002).
A recent study shows tumor-induced hypofunction of CAR T cells as well as upregulation of
PD-1 on the CAR T cells and demonstrates the contribution of PD-1 to the dysfunction of tumor-
infiltrating CAR T cells (Moon et al., 2014), thereby suggesting a potential strategy whereby
CAR T therapy could be combined with PD-1 blockade in cancer treatment (Chong et al., 2017).
Herein, in this study, in order to expand the application of CAR T cells in cancer
immunotherapy, we combined PD-1 blockade and CAR T cells and examined the antitumor
activity of the combined therapy in the treatment of lymphoma by using A20 murine B-cell
subcutaneous lymphoma model. We demonstrated that PD-1 blockade significantly improved the
antitumor capacity and enhanced the antitumor immunity of CAR-T cell therapy.

2.3 Materials and methods
Plasmid. The MSGV-1D3-28Z recombinant retroviral vector was kindly provided as a gift from
Dr. Steven A. Rosenberg. It encodes the MSGV (mouse stem cell virus-based splice-gag vector)
retroviral backbone and the 1D3-28Z.1-3 CAR. The 1D3-28Z.1-3 CAR consists of an anti-CD19

30

scFv, a portion of the murine CD28 molecule, and the intracellular component of the murine
TCR-ζ molecule in which the first and third ITAMs were inactivated.

Mice. BALB/c mice and the congenic strain CByJ.SJL (B6) (The Jackson Laboratories) were
used for in vivo efficacy studies. Mice were fed ad libitum and kept in air-conditioned rooms at
20°C ± 2°C with a 12-hour light–dark period. Animal care and manipulation were in conformity
with USC institutional guidelines, which were in accordance with the Guidelines for the Care
and Use of Laboratory Animals.

Cell culture. 293T cells were maintained in a 5% CO2 environment with Dulbecco’s modified
Eagle’s medium supplemented with 10% FBS, 2 mM L-glutamine, 100 U/ml penicillin and
100µg/ml streptomycin. A20 cells (ATCC number: TIB-208
TM
) was maintained in 5% CO
2

environment with RPMI-1640 medium supplemented with 10% FBS, 2 mM of L-glutamine, 100
U/ml penicillin, 100µg/ml streptomycin and 0.05mM β-mercaptoethanol (Sigma-Aldrich, St.
Louis, MO). All cell culture media and additives were purchased from Hyclone.

Retroviral T-cell Transduction. A total of 8-10 × 10
7
splenocytes were harvested from
CByJ.SJL (B6) mouse and activated in vitro using the T-cell activation Kit from e-Bioscience,
and supplemented with 15ng/ml mIL-2. Two days later, anti-mouse-CD19-CAR-encoding
retroviral vector was harvested, and added to non-tissue culture plates that had been coated
overnight with 15 µg/mL retronectin (Takara Bio Inc) at 4°C. Retroviral vector was spin-loaded
onto plates by centrifuging 2 hours at 2000g at 32°C. The activated T cells were washed and
resuspended at 10
6
/mL in R10, and 10
6
activated T cell were added per well to the retrovirus-

31

loaded plates. Plates were spun at 1000g at 32°C for 10 minutes and incubated overnight at
37°C, 5% CO
2
. The next day (day 3), the transduced T cells were transferred to a new 24-well
plates at 10
6
/mL in R10. Before treatment, CD19 CAR-transduced mouse T cell were evaluated
for expression of the appropriate CD19 CAR by Anti-Fab staining and flow cytometric analysis,
and cell function was evaluated by overnight co-culture with cognate antigen-bearing target cells
(5 × 10
5
:0.5 × 10
5
) and intracellular staining of interferon-γ (IFN-γ) was performed subsequently.
T cells for treatment were washed in saline before infusion into BALB/c mice intravenously.

Anti-CD19 CAR staining. To detect anti-CD19 CAR expression on the cell surface, cells were
stained with goat anti-rat Fab conjugated with FITC (Jackson ImmunoResearch). Before FACs
staining, 5 × 10
5
were harvested and washed three times with FACs buffer (PBS containing 5%
bovine serum albumin fraction V). Cell were then stained with 20 µg of goat anti-rat Fab
conjugated with FITC at 4°C for 30 minutes. Cells were washed and then fixed with transfix
cellular antigen stabilizing reagent (Thermo Scientific, Waltham, MA) at 4°C for 10 minutes.
Cells were then washed twice and stained with PE-anti-CD3, FITC-anti-CD4, PE/Cy5-anti-CD8
(BioLegend, San Diego, CA) at 4°C for 10 minutes. Cells were washed and resuspended in PBS.
Fluorescence was assessed using a MACSquant cytometer (Miltenyi Biotec, San Diego, CA),
and all the FACs data were analyzed using FlowJo software (Tree Star, Ashland, OR).

Surface immunostaining and flow cytometry. Splenic T cells (1 × 10
6
)

were cultured with
target cells for overnight at 37°C and 5% CO
2
in 96-well round bottom plates. PE-Cy5.5-anti-
CD3, Pacific Blue
TM
-anti-CD8, PE-anti-PD-1, FITC-anti-TIM-3 and APC-anti-LAG-3
antibodies were used for the cell surface staining.

32

Intracellular staining. Splenic T cells (1 × 10
6
)

were cultured with target cells for 16 hours at
37°C and 5% CO
2
with (IFN-γ) or without (Ki-67 and p-Akt) GolgiPlug (BD Biosciences, San
Jose, CA) in 96-well round bottom plates. PE-Cy5.5-anti-CD3, FITC-anti-CD4, Pacific blue-
CD8, PE-anti-IFN-γ, PE-anti-Ki-67 and APC-anti-p-Akt antibodies were used for the
intracellular staining. Cytofix/Cytoperm Fixation and Permeabilization Kit (BD Biosciences)
was used to permeabilize the cell membrane and perform intracellular staining according to the
manufacturer’s instruction.

Specific cell lysis assay. A20 cells were labeled by suspending them in PBS+0.1% BSA with 5
µM Carboxyfluorescein succinimidyl ester (CFSE) fluorescent dye at a concentration of 1 × 10
6

cells/mL. The cells were incubated for 30 minutes at 37°C. After incubation, the same volume of
FBS was added into the cell suspension and then incubated for 2 minutes at room temperature.
The cells were then washed twice and suspended in fresh R10 medium. Cocultures were set up in
round bottom 96-well plates in triplicate at the following effector-to-target ratios: 1:1, 5:1 and
10:1. The cultures were incubated for 6 hours at 37°C, followed by 7-AAD labeling, according
to the manufacturer’s instructions (BD Biosciences). Flow cytometric analysis was performed to
quantify the percent survival (7-AAD-negative) of target cells. In the wells containing only target
cells without effector cells, the percentage of viable A20 cells was calculated and used to correct
the variation in the starting cell numbers and spontaneous cell death. The cytotoxicity was
determined in triplicate and presented in mean ± SEM.

33

ELISA. IFN-γ was measured using mouse IFN-γ ELISA kit (BD Biosciences, San Jose, CA)
according to the manufacturer’s instructions respectively. Briefly, 96-well ELISA plates
(Thermo Scientific, Waltham, MA) were coated with 200 ng/well of capture antibodies against
the indicated proteins at 4°C overnight. On the next day, plates were washed with wash buffer
(PBS containing 0.05% Tween 20) and blocked with assay buffer (PBS containing 10% FBS) for
2 hours at room temperature. Equal volume of serum or protein extract was added to the plate
and incubated for 2 hours at room temperature. Plates were then washed and incubated with
detection antibodies for 1 hour at room temperature.

In vivo study. BALB/c mice were preconditioned (5 Gy) and then injected with 10 million A20
cell on the right flank on Day 0. On the day 6, the mice were injected with scFv-anti-CD19-
transduced T cells or non-transduced T cells (6×10
6
/dose) from congenic CByJ.SJL (B6) mice on
days 6 in combination with either anti-PD-1 or isotype antibodies (250 µg/injection on days 6, 9,
and 13). Control groups were left untreated. Tumor growth in mice was monitored every 2 to 4
days. Tumor size was measured by calipers and calculated by the following formula: W
2
× L / 2.
Mice were euthanized when they displayed obvious weight loss, ulceration of tumors, or tumor
size larger than 1000 mm
3
.

Statistical analysis. Statistical analysis was performed in GraphPad Prism (GraphPad Software,
San Diego, CA, USA), version 5.01. One-way ANOVA with Tukey’s multiple comparison was
performed to assess the differences among different groups in the in vitro assays. Tumor growth
curve was analyzed using one-way ANOVA with repeated measures (Tukey’s multiple
comparison method). Mouse survival curve was evaluated by the Kaplan-Meier analysis (log-

34

rank test with Bonferroni correction). A P value less than 0.05 was considered statistically
significant. Significance of findings was defined as: ns = not significant, P > 0.05; *, P < 0.05;
**, P < 0.01; ***, P < 0.001.

2.4 Results
2.4.1 Expression and activation of 1D3-28Z.1-3 anti-CD19 CAR in mouse splenic T cells
The schematic design of the 1D3-28Z.1-3 anti-CD19 CAR used in this study is shown in Figure
2-1A and has also been reported by the others (Kochenderfer et al., 2010). To examine the
expression of 1D3-28Z.1-3 in primary lymphocytes, mouse splenic T cells were transduced by
retrovirus expressing 1D3-28Z.1-3 CAR. As shown in Figure 2-1B, reproducible levels of 1D3-
28Z.1-3 were observed in transduced primary splenic T cells. The median percentage of CAR-
expressing T cells was 76%. Co-stimulation through either CD28 or 4-1BB has been shown to
regulate multiple aspects of T cell function by activating PI3K/Akt signaling pathway (Parry et
al., 1997; Starck et al., 2005). To evaluate the activation of downstream signaling pathway of
CAR19 T cells upon antigen-specific stimulation, non-transduced or CAR19 T cells were co-
cultured with CD19-expressing murine A20 B-cell lymphoma cells without exogenous IL-2 for
16 h before they were subjected to intracellular staining for the expression of phosphorylated
Akt. We found that p-Akt (ser473) in CAR19 CD8
+
T cells was significantly increased upon
antigen-specific stimulation, compared to non-transduced CD8
+
T cells (15.9±3.0% versus
7.6±2.1%; Figure 2-1C and 2-1D).

35

Figure 2-1 Expression and activation of 1D3-28Z.1-3 anti-CD19 CAR in mouse splenic T
cells.
(A) Schematic of retroviral construct encoding 1D3-28Z.1-3 anti-CD19 CAR. scFv, single
chain variable fragment; CD28, a portion of mouse CD28 molecule; CD3, the intracellular
component of the mouse TCR-ζ molecule; LTR, long terminal repeat. (B) CAR expression
was detected in murine splenic T cells by flow cytometry following FITC-conjugated goat
anti-rat Fab staining. A viable CD3
+
lymphocyte gating strategy was used. CD8
+
T cells were
shown in each panel. CAR-expressing CD8 T cells were gated and shown in each scatterplot. NT
indicates non-transduced T cells, which were used as a control. (C) Intracellular expression of
p-Akt in T cells. p-Akt-expressing CD8 T cells were gated and shown in each scatterplot. (D)
The summarized statistics of (C) were shown in bar graphs (n=3, mean ± SEM; **P < 0.01).

NT
1D3 CAR
23
0
5
10
15
20
**
% CD3
+
CD8
+
pAkt
+
A
B
5’LTR 1D3 scFv CD28
CD3ζ 1
st
and 3
rd

ITAMs inactive
3’LTR
Ψ
C
NT CAR
CD8-PE/Cy5
Anti-Fab-FITC
D
CD8-PE/Cy5
p-Akt-APC
NT CAR

36

2.4.2 PD-1 expression is upregulated on anti-CD19-CAR-transduced T cells following
antigen-specific stimulation
It has been shown that PD-1 expression on T cells is increased following T-cell activation via
TCR recognition of MHC/peptide (Day et al., 2006; Petrovas et al., 2006; Trautmann et al.,
2006). In addition, the expression of PD-1 was also shown to be upregulated on anti-HER2 CAR
T cells with antigen-specific stimulation (John et al., 2013). To evaluate if antigen-specific
stimulation of T cells would have similar effect and enhance the PD-1 expression on anti-CD19
CAR T cells, both the non-transduced and CAR19 T cells were cultured with or without A20
cells for overnight. We found that without antigen stimulation, the cell surface expression of PD-
1 on CAR19 T cells is similar to that on the non-transduced T cells. However, upon CAR
stimulation, the expression of PD-1 on CAR19 CD8
+
T cells was increased and significantly

Figure 2-2 Upregulation of PD-1 expression on CAR T cells following antigen-specific
stimulation.
Non-transduced T cell or CAR19 T cells were stimulated with or without A20 B-cell
lymphoma cells at an E/T ratio of 1:1 for overnight. The cell surface expression of (A) PD-1
and (B) LAG-3 was detected by flow cytometry following cell surface staining. The
percentage of PD-1 (A) or LAG-3 (B) expressing CD8 T cells over total CD8
+
T cells was
shown in bar graphs (n=3; ns, not significant, P > 0.05; **P < 0.01).
Media
A20
0
5
10
15
**
ns
% CD3
+
CD8
+
LAG-3
+
A B
Media
A20
0
5
10
15
20
NT
CAR
**
% CD3
+
CD8
+
PD-1
+
Media
A20
0
5
10
15
20
NT
CAR
**
% CD3
+
CD8
+
PD-1
+

37

higher when compared to the non-transduced T cells (15.6±1.9% versus 5.2±0.25%; Figure 2-
2A).
In addition to PD-1, other cell surface inhibitory molecules, including lymphocyte activation
gene 3 protein (LAG-3), T cell immunoglobulin domain and mucin domain-containing protein 3
(TIM-3; also known as HAVCR2) and cytotoxic T-lymphocyte associated protein 4 (CTLA-4),
also play important roles in inducing T cell exhaustion and limiting the antitumor efficacy of
CAR-T cell therapy (Pardoll, 2012). In order to evaluate whether the expression of other T cell
exhaustion markers is regulated by CAR stimulation, we measured the expression of LAG-3 and
TIM-3 in CAR19 T cells. We found that compared to non-transduced T cells, without antigen
stimulation, the basal level of LAG-3 on CAR19 CD8
+
T cells was significantly higher
(8.54±0.5% versus 4.59±1.1%; Figure 2-2B). With antigen-specific stimulation, the expression
of LAG-3 on both T cells was upregulated and the expression between non-transduced and
CAR19 T cells showed little difference (Figure 2-2B). The expression of TIM-3 on both non-
transduced and CAR19 T cells was undetectable (data not shown).

2.4.3 PD-1 blockade enhances the antigen-specific immune responses of 1D3-28Z.1-3 CAR
T cells
We have shown that PD-1, the expression of which plays an important role in inhibiting T cell
activation and inducing T cell exhaustion, was significantly upregulated on CAR19 T cells
following antigen-specific stimulation. Next, to assess the effect of PD-1 blockade on the
functional capacity of CAR19 T cells, the T cells that transduced with 1D3-28Z.1-3 were co-
cultured with A20 cells in combination with either anti-PD-1 or control isotype antibodies and
the expression of T cell function marker IFN-γ and cell proliferation marker Ki-67 was

38

evaluated. Notably, it revealed that PD-1 blockade significantly increased the expression of IFN-
γ and Ki-67 on CAR19 T cells, compared to the group that treated with control isotype antibody
(Figure 2-3A and 2-3B).

Figure 2-3 Anti-PD-1 enhanced the antigen-specific immune responses of CAR T cells.
To examine the effects of PD-1 blockade on T-cell function, non-transduced T cells or
CAR19 T cells were co-cultured with A20 cells at an E/T ratio of 10:1 (IFN-γ) or 1:1 (Ki-
67) for overnight in the presence of anti-PD-1 or isotype control antibodies. The T cells
were then harvested and stained intracellularly for IFN-γ (A) and Ki-67 (B) as determined
by flow cytometry. (C) The ability of CAR19 T cells to lyse A20 cells with or without the
presence of anti-PD-1 antibody was determined by flow cytometry after 6-hour co-culture at
1:1, 5:1, and 10:1 effector-to-target ratios. Non-transduced (NT) T cells were used as a control.
Error bars represent standard error of the mean for each treatment group (n = 3; *P < 0.05;
**P < 0.01).
C
B A
NT
CAR+iso
CAR+αPD-1
0
5
10
15
**
% CD3
+
CD8
+
Ki67
+
NT
CAR+iso
CAR+αPD-1
0
10
20
30
**
% CD3
+
CD8
+
IFNγ
+
1:1
5:1
10:1
0
10
20
30
40 NT
CAR+iso
CAR+αPD-1
E:T ratio
% Cytotoxicity
*

39

Furthermore, the cytolytic activity of CAR19 T cells with or without PD-1 blockade was
examined in a 6-hour cytotoxicity assay. The cytotoxic activity of T cells transduced with 1D3-
28Z.1-3 against A20 cells was evaluated at E/T ratios of 1, 5 and 10. We found that CAR19 T
cells mediated significant cell lysis of target cells at E/T ratios of 5 and 10, but not at E/T ratio of
1, compared to the non-transduced T cells. Interestingly enough, with PD-1 blockade, CAR19 T
cells induced significant cytotoxicity of target cells at E/T ratio of 1, compared to that without
PD-1 blockade (Figure 2-3D).

2.4.4 PD-1 blockade enhances 1D3-28Z.1-3 CAR T cell-mediated tumor regression in
established tumor model
Next, to evaluate the antitumor efficacy of combined PD-1 blockade with anti-CD19 CAR T
cells, we adoptively transferred 6 x 10
6
CAR19 T cells into BALB/C mice bearing established
A20 subcutaneous B-cell lymphoma (~30 mm
3
). 6-hour later on the same day, we intravenously
injected anti-PD-1 or control isotype antibodies. The antibody injection was continued twice a
week for two weeks. The experimental design for animal study is shown in Figure 2-4A (upper
panel). We found that anti-PD-1 antibody or CAR19 T cells treatment alone significantly
inhibited tumor growth, compared to the control group. Notably, combined PD-1 blockade and
CAR19 T cells further enhanced the antitumor efficacy, compared to either treatment given alone
(Figure 2-4A, lower panel). Moreover, the combination therapy significantly enhanced the long-
term survival, compared to monotherapy (Figure 2-4B). Further analysis of the engraftment and
expansion of CAR T cells were assessed. Three days following T cell infusion, mice were
euthanized, and different organs and tissues, including the tumor, blood, and spleen, were

40

harvested for transferred-T cell staining. We found that certain percentage of CD45.1
+
T cells (2-
5%) circulated in the blood, whereas most of the CD45.1
+
T cells (15-20%) homed to the spleen
in all groups. Particularly, compared to CAR19 T cells monotherapy, the combination therapy
resulted in significantly higher infiltration of transferred T cells into the spleen. The infiltration
of CD45.1
+
T cell in the tumor was low and showed little difference across all the examined

Figure 2-4 PD-1 blockade enhanced CAR19 T cells mediated tumor regression of
established tumor.
(A) Adoptive transfer of CAR19 T cells in combination with PD-1 blockade enhances
growth inhibition of established tumor. BALB/c mice were preconditioned (5 Gy) and then
injected with 10 million A20 cell on the right flank on Day 0. On the day 6, the mice were
injected with CAR19 T cells or non-transduced T cells (6×10
6
/dose) from congenic from
congenic CByJ.SJL (B6) mice on days 6 in combination with either anti-PD-1 or isotype
Tumor
Blood
Spleen
0
10
20
30
40
50
Day 9 Posttherapy
ns
*
ns
ns
ns
***
CAR+iso
nontreated
NT+αPD-1
CAR+αPD-1
% CD45.1
+
A
C
X-ray irradiation
Tumor inoculation
A20, 10×10
6
Day -5 0 3
CAR-T, i.v.
6×10
6
9
Euthanize
Monitor tumor growth
B
D
0 5 10 15 20 25
0
20
40
60
80
100
nontreated
NT+αPD1
CAR+iso
CAR+αPD1
Days post T cell injection
Percent survival
P<0.001
Tumor
Blood
Spleen
0
5
10
15
20
25
nontreated
NT+αPD-1
CAR+iso
CAR+αPD-1
Day 3 Posttherapy
**
% CD45.1
+
0
2
4
8
13
0
400
800
1200
nontreated
NT+αPD-1
CAR+iso
CAR+αPD-1
*
*
*
*
Days post T cell injection
Tumor volume (mm
3
)

41

antibodies (250 µg/injection on days 6, 9, and 13). Control groups were left untreated. Tumor
growth in mice was monitored every 2 to 4 days. Tumor size was measured by calipers and
calculated by the following formula: W
2
× L / 2. Mice were euthanized when they displayed
obvious weight loss, ulceration of tumors, or tumor size larger than 1000 mm
3
. Tumor growth
curve was shown for mice without treatment, treated with non-transduced (NT) plus anti-PD-1
injection, CAR19 plus isotype control antibody, or CAR19 plus anti-PD-1 antibody injection.
Data were presented as mean tumor volume ± standard error of the mean (SEM) at indicated
time points (n = 6; *P<0.05). (B) Survival of A20 tumor-bearing mice after indicated treatment.
Overall survival curves were plotted using the Kaplan-Meier method and compared using the
log-rank (Mantel-Cox) test (n ≥ 7). The percentage of transferred CD45.1
+
T cells in the tumor,
blood, and spleen of A20 tumor-bearing mice that were adoptively transferred with non-
transduced (NT) plus anti-PD-1 injection, CAR19 plus isotype control antibody, or CAR19 plus
anti-PD-1 antibody injection was investigated by flow cytometry on day 3 (C) or day 9 (D) post-
therapy (n = 3, mean ± SEM; *P<0.05; **P < 0.01; ***P < 0.001).

tissues (Figure 2-4C). One-week post-T cell infusion, on day 9, we observed a significant
expansion of transferred CD45.1
+
T cells in the circulation of mouse treated with CAR19 T cells
or combined CAR19 T cells and anti-PD-1 antibody. Remarkably, compared to either treatment
given alone, in the combined treatment group, CD45.1
+
T cells specifically accumulated and
expanded in the tumor, whereas the population of CD45.1
+
T cells had little variation across all
the treatment groups (Figure 2-4D).

2.4.5 The antitumor efficacy of combined therapy is associated with tumor
microenvironment modulation
It has been well established that both cytotoxic CD8
+
T cells and helper CD4
+
T cells were
involved in antitumor immunity (Fridman et al., 2012). To further understand the underlying
mechanism of enhanced antitumor activity of combination therapy, we therefore assessed the

42

effects of combined therapy on tumor infiltrating lymphocytes. We found that compared to
CAR19 T cells monotherapy, the combined therapy significantly expanded the population of
CD8
+
T and CD4
+
T in the tumor microenvironment (Figure 2-5A and 2-5B).

Figure 2-5 The enhanced antitumor efficacy of combined therapy is correlated with tumor
microenvironment modulation.
Tumors were taken on day 13 post-therapy from mice treated with non-transduced (NT) T
cells plus anti-PD-1 antibody, CAR19 T cells plus isotype control antibody, or CAR19 T
cells plus anti-PD-1 antibody. Non-treated mice were used as a control. The percentage of
CD8
+
T (A) and CD4
+
T (B) cells at the tumor sites was examined by flow cytometry. Error
bars represent standard error of the mean for each treatment group (n=3; ns, not significant, P
> 0.05; **P < 0.01).

2.4.6 Combination therapy specifically regulates local immune responses within tumor
To further substantiate the observed changes of tumor microenvironment, we examined the
expression of T-cell activation marker in tumor-bearing mice by ELISA. It showed that CAR19
T cells monotherapy significantly increased the expression of IFN-γ in the tumor, compared to
the control group. Moreover, the PD-1 blockade further enhanced CAR19 T cell-mediated IFN-γ
up-regulation (Figure 2-6A). However, the measurement of IFN-γ expression in the serum and
nontreated
NT+αPD-1
CAR+iso
CAR+αPD-1
0
1
2
3
4
5
**
ns
% CD45
+
CD3
+
CD8
+
A B
nontreated
NT+αPD-1
CAR+iso
CAR+αPD-1
0
2
4
6 **
ns
% CD45
+
CD3
+
CD4
+

43

spleen showed that the combined therapy had little effects on IFN-γ expression, compared to
CAR19 T cells monotherapy (Figure 2-6B and 2-6C).

Figure 2-6 Combined therapy specifically regulates immune responses at local tumor site.
(A) Tumors, (B) Serum and (C) Spleens were harvested on day 9 post-therapy from mice
treated with non-transduced (NT) T cells plus anti-PD-1 antibody, CAR19 T cells plus
isotype control antibody, or CAR19 T cells plus anti-PD-1 antibody. Non-treated mice were
used as a control. The expression of IFN-γ was quantified using ELISA. Error bars represent
standard error of the mean for each treatment group (n=3; ns, not significant, P > 0.05; *P <
0.05).

A
C
B
nontreated
NT+αPD1
CAR+iso
CAR+αPD1
0.0
0.5
1.0
1.5
IFN-γ(ng/ml)
*
Tumor
nontreated
NT+αPD1
CAR+iso
CAR+αPD1
0
1
2
3
IFN-γ(ng/ml)
ns
Spleen
nontreated
NT+αPD1
CAR+iso
CAR+αPD1
0.0
0.5
1.0
1.5
2.0
IFN-γ(ng/ml)
ns
Serum

44

2.5 Discussion
Adoptive T cell therapy using gene-modified T cells has emerged as a promising method for
cancer immunotherapy. It has achieved successful responses in patients with certain
hematopoietic malignancies. However, in the treatment of other blood-borne tumors, such as FL
and DLBCL, or solid tumors, the outcome has been less promising, partly owing to the
immunosuppressive properties and establishment of an immunosuppressive microenvironment
within the tumors (Vazquez-Cintron et al., 2010). The PD-1/PD-L1 regulatory pathway has
demonstrated a particularly antagonistic effect on the antitumor response of TILs. The FL TILs
were shown to have high PD-1 expression and PD-L1
+
histiocytes were also found within the T
cell-rich zone of the neoplastic follicles in FL (Myklebust et al., 2013). In addition to FL, the
other lymphomas or solid tumors with poor prognosis also showed upregulation of PD-L1
expression, with increasing PD-1 expression on TILs (Myklebust et al., 2013; Wu et al., 2015;
Xia et al., 2016). The combined effect of these two leads to tumor escape. However, this can be
disrupted by the use of checkpoint inhibitors (CPIs), targeting the PD-1/PD-L1 pathway (Ansell
et al., 2015; Brahmer et al., 2010; Dong et al., 2002). As a result, in this study, we combined
anti-PD-1 antibody and CAR19 T cells to evaluate the effects of PD-1 blockade in infused
CAR19 T cells and CAR19 T cell-mediated antitumor immunity.
Herein we have demonstrated that the expression of PD-1 was significantly increased on CAR19
T cells following antigen-specific stimulation and that PD-1 blockade significantly increased the
functional capacity of CAR19 T cells both in vitro and in vivo. In the adoptive T cell transfer
experiment, we demonstrated that combined PD-1 blockade and CAR19 T cells significantly
enhanced the CAR T cell-mediated inhibition of established tumor growth. Interestingly, we
found that the enhanced antitumor efficacy of combined therapy was correlated with increased

45

local accumulation and proliferation of adoptive transferred T cell and elevated modulation of
tumor microenvironment and immune responses within tumor.
It has been shown that the expression of PD-1 is increased shortly after T-cell activation through
TCR recognition of MHC/peptide (Day et al., 2006; Petrovas et al., 2006; Trautmann et al.,
2006). In order to investigate the possibility that PD-1 can be similarly upregulated on CAR19 T
cells following antigen-specific stimulation, we co-cultured the T cells with target cells for
overnight and measured PD-1 expression. In consistent with previous observation, which showed
that the expression of PD-1 in anti-Her2 CAR T cells is elevated upon antigen-specific activation
(John et al., 2013), we demonstrated that CAR stimulation significantly enhanced PD-1
expression on anti-CD19-CAR-transduced CD8
+
T cells. This observation suggests that the up-
regulation of PD-1 may contribute to limiting the full potential of CAR T cells in inducing anti-
tumor immunity. It has been shown that co-expression of multiple inhibitory receptors is a
cardinal feature of T cell exhaustion (Thommen et al., 2015; Wherry and Kurachi, 2015). Thus,
in addition to PD-1, other cell surface inhibitory molecules were also examined. We found that
the expression level of TIM-3 on both rested T cells and activated T cells was minimal and
undetectable. Interestingly, we observed that without stimulation, the basal level of LAG-3 on
CAR19 T cells was significantly higher than that on non-transduced T cells. Upon antigen
stimulation, LAG-3 expression was upregulated on both T cells, whereas little difference was
seen between non-transduced and CAR19 T cells T cells, suggesting that the upregulation of
LAG-3 may also contribute to limiting the activity of CAR T cells as PD-1 does, but through
different mechanisms (Buchbinder and Desai, 2016).
The PD-1/PD-L1 pathway involves the regulation of cytokine production by T cells, inhibiting
production of IFN-γ, TNF-α and IL-2 (Taylor et al., 2011). In this study, to unleash the inhibitory

46

effect of tumor-induced upregulation of PD-1 on CAR T cells, we combined anti-PD-1 antibody
with CAR19 T cells, and demonstrated that PD-1 blockade significantly enhanced T cell
functional capacity by increasing the production of IFN-γ. In addition to cytokine production,
PD-1 can also inhibit T cell proliferation (Keir et al., 2008). With CAR-specific stimulation in
the presence of PD-L1
+
cancer cells, we found that anti-PD-1 antibody significantly increased
the expression of Ki-67 in CAR19 T cells, implying that PD-1 blockade confers the CAR19 T
cells to be more proliferated. Taken together, these data imply that PD-1/PD-L1 signaling
blockade results in more functional CAR19 T cells with higher proliferation capacity.
Our in vivo study showed that the tumor growth could be inhibited by CAR19 T cells treatment
or anti-PD-1 antibody treatment given alone. However, in comparison, combined PD-1 blockade
and CAR19 T cells further enhanced the inhibitory effect on tumor growth and pro-longed the
survival. To understand the underlying mechanism of enhanced antitumor efficacy of the
combined therapy, we analyzed the expansion and infiltration of adoptively transferred T cells in
different tissues. We found that PD-1 blockade specifically increased the accumulation of
adoptively transferred T cells to local tumor tissues. However, in the other tissues, such as blood
and spleen, the accumulation of T cell is similar in groups with and without PD-1 blockade.
From our current study, it is unclear whether the administrated anti-PD-1 antibody enhances
CAR T cells infiltration, enhances T cell retention, or enhances T cell proliferation in the tumor
microenvironment, but also the effect could be a result of a combination of these effects. A
previous study by Wong et al. (2007) reported that PD-1 blockade augments proliferation (Wong
et al., 2007). This data corresponds with our in vitro experiment that PD-1 blockade enhanced
cell proliferation of CAR19 T cells. Moreover, it has been reported that PD-1/PD-L1 blockade
reduces T cell motility (Zinselmeyer et al., 2013), which may lead to higher T cell retention in

47

the tumor microenvironment. In addition to adoptively transferred T cells, we also examined the
endogenous tumor infiltrating T cells and found that PD-1 blockade significantly increased the
population of tumor infiltrating CD8
+
T and CD4
+
T cells. The population of cytotoxic CD8
+
T
and helper CD4
+
T cells among TILs is critical in eliciting antitumor immunity (Ceeraz et al.,
2013). Taken together, the increased CAR19 T cells trafficking to tumor and the modulation of
tumor microenvironment may both contribute to the enhanced antitumor immunity of combined
therapy, which was also supported by the specifically increased expression of IFN-γ in the
tumor.

2.6 Conclusion
In summary, our study support the immunotherapeutic approaches that combine immune
checkpoint blockade with CAR-T cell therapy. Tumor-induced upregulation of PD-1 on the
surface of CAR T cells and the low tumor infiltration of cytotoxic T cells lead to tumor evasion
to CAR-T cell therapy. In this study, we have demonstrated that PD-1 blockade allows enhanced
CAR T cells functional capacity, increased accumulation of CAR T cells, and infiltration of
endogenous cytotoxic CD8
+
and helper CD4
+
T cell into local tumor tissue, thereby leading to
enhanced antitumor immunity of CAR-T cell therapy.

48

Chapter 3. Enhanced Cancer Immunotherapy by Chimeric Antigen
Receptor-Modified T Cells Engineered to Secrete Checkpoint
Inhibitors

49

3.1 Abstract
Despite favorable responses of chimeric antigen receptor (CAR)-engineered T cell therapy in
patients with hematologic malignancies, the outcome has been far from satisfactory in the
treatment of solid tumors, partially owing to the development of an immunosuppressive tumor
microenvironment. To overcome this limitation, we engineered CAR-T cells secreting
checkpoint inhibitors (CPIs) targeting PD-1 (CAR.αPD1-T) and evaluated their efficacy in a
human lung carcinoma xenograft mouse model. In an effort to evaluate the effector function and
expansion capacity of CAR.αPD1-T cells in vitro, we measured the production of IFN-γ and T
cell proliferation following antigen-specific stimulation. Furthermore, the antitumor efficacy of
CAR.αPD1-T cells, CAR-T cells, and CAR-T cells combined with anti-PD-1 antibody was
determined using a xenograft mouse model. Finally, the underlying mechanism of enhanced
tumor eradication of CAR.αPD1-T cells was investigated by analyzing the expansion and
functional capacity of tumor-infiltrating lymphocytes. We demonstrated that human anti-PD-1
CPIs secreted by CAR.αPD1-T cells efficiently bound to PD-1 and reversed the inhibitory effect
of PD-1/PD-L1 interaction on T cell function. PD-1 blockade by continuously secreted anti-PD-1
prevented T cell exhaustion and significantly enhanced T cell expansion and effector function
both in vitro and in vivo. In the xenograft mouse model, we demonstrated that the secretion of
anti-PD-1 enhanced the antitumor activity of CAR-T cells and prolonged overall survival. In
conclusion, with constitutive anti-PD-1 secretion, CAR.αPD1-T cells are less exhausted, more
functional and expandable, and more efficient at tumor eradication than parental CAR-T cells.
Collectively, our study presents an important and novel strategy that enables CAR-T cells to
achieve better antitumor immunity, especially in the treatment of solid tumors.

50

3.2 Introduction
Adoptive cell transfer (ACT), as a modality of immunotherapy for cancer, has demonstrated
remarkable success in treating hematologic malignancies and malignant melanoma (Grupp et al.,
2013; Khammari et al., 2009; Mackensen et al., 2006; Maus et al., 2014; Park et al., 2016). An
especially effective form of ACT, which uses gene-modified T cells expressing a chimeric
antigen receptor (CAR) to specifically target tumor-associated-antigen (TAA), such as CD19 and
GD2, has displayed encouraging results in clinical trials for treating such diseases as B cell
malignancies and neuroblastoma (Davila et al., 2014; Gui et al., 2016; Louis et al., 2011).
Unlike naturally occurring T cell receptors (TCRs), CARs are artificial receptor consisting of an
extracellular antigen recognition domain fused with intracellular T cell signaling and
costimulatory domains. CARs can directly and selectively recognize cell surface TAAs in a
major histocompatibility class (MHC)-independent manner (Kershaw et al., 2013). Despite the
documented success of CAR T cell therapy in patients with hematologic malignancies, only
modest responses have been observed in solid tumors. This can be attributed, in part, to the
establishment of an immunosuppressive microenvironment in solid tumors. Such milieu involves
the upregulation of a number of intrinsic inhibitory pathways mediated by increased expression
of inhibitory receptors (IRs) in T cells reacting with their cognate ligands within the tumor
(Bonifant et al., 2016; Gill et al., 2016).
So far, several IRs have been characterized in T cells, such as CTLA-4, T cell Ig mucin-3 (TIM-
3), lymphocyte-activation gene 3 (LAG-3), and programmed death-1 (PD-1) (Pardoll, 2012).
These molecules are upregulated following sustained activation of T cells in chronic disease and
cancer, and they promote T cell dysfunction and exhaustion, thus resulting in escape of tumor
from immune surveillance (Pardoll, 2012). Unlike other IRs, PD-1 is upregulated shortly after T

51

cell activation, which in turn, inhibits T cell effector function via interacting with its two ligands,
PD-L1 or PD-L2. PD-L1 is constitutively expressed on T cells, B cells, macrophages, and
dendritic cells (DCs) (Yamazaki et al., 2002). PD-L1 is also shown to be abundantly expressed in
a wide variety of solid tumors (Brown et al., 2003; Dong et al., 2002; Konishi et al., 2004). In
contrast, the expression of PD-L1 in normal tissues is undetectable (Dong et al., 2002). As a
consequence of its critical role in immunosuppression, PD-1 has been the focus of recent
research, aiming to neutralize its negative effect on T cells and enhance antitumor responses.
Clinical studies have demonstrated that PD-1 blockade significantly enhanced tumor regression
in colon, renal and lung cancers and melanoma (Pardoll, 2012).
A recent study shows tumor-induced hypofunction of CAR T cells as well as upregulation of
PD-1 on the CAR T cells and demonstrates the contribution of PD-1 to the dysfunction of tumor-
infiltrating CAR T cells (Moon et al., 2014), thereby suggesting a potential strategy whereby
CAR T therapy could be combined with PD-1 blockade in cancer treatment (Newick et al.,
2016b). Therefore, in this study, in order to overcome the inhibitory effect of PD-1 signaling in
CAR T cells, we genetically engineered CAR T cells with the capacity to continuously produce a
single-chain variable fragment (scFv) form of anti-PD-1 antibody. In our own tumor models, we
found that anti-PD-1 scFv expression and secretion could interrupt the engagement of PD-1 with
its ligand, PD-L1, and prevent CAR T cells from being inhibited and exhausted. Most
importantly, in a CD19 tumor model, we demonstrated for the first time that the secretion of anti-
PD-1 scFv by CAR T cells could significantly improve the capacity of CAR T cells in
eradicating an established solid tumor.

52

3.3 Materials and methods
Mice. Six- to eight-week-old female NOD.Cg-Prkdc
scid
IL2Rg
tm1Wj1
.Sz (NSG) mice were
purchased from Jackson Laboratory (Farmington, CT). All animal studies were performed in
accordance with the Animal Care and Use Committee guidelines of the NIH and were conducted
under protocols approved by the Animal Care and Use Committee of the NCI.

Cell culture and antibodies. Cell lines SKOV3 and 293T were obtained from ATCC. The lung
cancer line NCI-H292 was kindly provided by Dr. Ite Laird-Offringa (University of Southern
California, Los Angeles, CA). The H292-CD19 and SKOV3-CD19 cell lines were generated by
the transduction of parental NCI-H292 and SKOV3 cells with a lentiviral vector encoding the
cDNA of human CD19. The transduced H292 and SKOV3 cells were stained with anti-human
CD19 antibody (BioLegend, San Diego, CA) and sorted to yield a relatively pure population of
CD19-overexpressing cells. SKOV3, SKOV3-CD19, NCI-H292, and H292-CD19 cells were
maintained in R10 medium consisting of RPMI-1640 medium supplemented with 10% fetal
bovine serum (FBS), 2 mM L-glutamine, 10 mM HEPES, 100 U/ml penicillin and 100µg/ml
streptomycin. The 293T cells were cultured in D10 medium consisting of DMEM medium
supplemented with 10% FBS, 2 mM L-glutamine, 10 mM HEPES, 100 U/ml penicillin and
100µg/ml streptomycin. All above cell culture media and supplements were purchased from
Hyclone (Logan, UT). Human peripheral blood mononuclear cells (PBMCs) were cultured in T
cell medium (TCM), which is composed of X-Vivo 15 medium (Lonza, Walkersville, MD)
supplemented with 5% human AB serum (GemCell, West Sacramento, CA), 1% HEPES (Gibco,
Grand Island, NY), 1% Pen-Strep (Gibco), 1% GlutaMax (Gibco), and 0.2% N-Acetyl Cysteine
(Sigma-Aldrich, St. Louis, MO).

53

Primary antibodies used in this study include biotinylated Protein L (GeneScript, Piscataway,
NJ); PE-anti-CD45, PE-Cy5.5-anti-CD3, FITC-anti-CD4, Pacific Blue
TM
-anti-CD8, FITC-anti-
CD8, PE-anti-IFN-γ, Brilliant Violet 421
TM
-anti-PD-1, PE-anti-PD-L1, PerCP/Cy5.5-anti-LAG-
3, and PE-anti-TIM-3 (BioLegend, San Diego, CA); and Rabbit anti-HA tag antibody (Abcam,
Cambridge, MA). The secondary antibodies used were FITC-conjugated streptavidin
(BioLegend, San Diego, CA) and goat anti-rabbit IgG-HRP (Santa Cruz, San Jose, CA). The
SuperSignal® West Femto Maximum Sensitivity Substrate used for Western blot analysis was
from Thermo Fisher Scientific (Waltham, MA).

Plasmid construction. The retroviral vector encoding anti-CD19 CAR (CAR) was constructed
based on the MP71 retroviral vector kindly provided by Prof. Wolfgang Uckert, as described
previously (Engels et al., 2003). The vector encoding anti-CD19 CAR with anti-PD-1 scFv
(CAR.αPD1) was then generated based on the anti-CD19 CAR. The insert for CAR.αPD1 vector
consisted of the following components in frame 5’ end to 3’ end: the anti-CD19 CAR, an EcoRI
site, a leader sequence derived from human IL-2, the anti-PD-1 scFv light chain variable region,
a GS linker, the anti-PD-1 scFv heavy chain variable region, the HA-tag sequence, and a NotI
site.
The anti-PD-1 scFv portion in the CAR.αPD1 vector was derived from the amino acid sequence
of human monoclonal antibody 5C4 specific against human PD-1 (Alan J. Korman, 2011). The
corresponding DNA sequence of the scFv was codon-optimized for its optimal expression in
human cells using the online codon optimization tool and was synthesized by Integrated DNA
Technologies (Coralville, IA). The anti-PD-1 scFv was then ligated into the CD19 CAR vector
via the EcoRI site through the Gibson assembly method.

54

Retroviral vector production. Retroviral vectors were prepared by transient transfection of
293T cells using a standard calcium phosphate precipitation protocol. 293T cells cultured in 15-
cm tissue culture dishes were transfected with 37.5 µg of the retroviral backbone plasmid, along
with 18.75 µg of the envelope plasmid pGALV and 30 µg of the packaging plasmid encoding
gag-pol. The viral supernatants were harvested 48 h post-transfection and filtered through a 0.45
µm filter (Corning, Corning, NY) before use.

T cell transduction and expansion. Frozen human PBMCs were obtained from AllCells
(Alameda, CA). PBMCs were thawed in TCM and rested overnight. Before retroviral
transduction, PBMCs were activated for 2 days by culturing with 50 ng/ml OKT3, 50 ng/ml anti-
CD28 antibody, and 10 ng/ml recombinant human IL-2 (PeproTech, Rocky Hill, NJ). For
transduction, freshly harvested retroviral supernatant was spin-loaded onto non-tissue culture-
treated 12-well plates coated with 15 µg retronectin (Clontech Laboratories, Mountain View,
CA) per well by centrifuging 2 hours at 2000×g at 32°C. The spin-loading of vector was repeated
once with fresh viral supernatant. Activated PBMCs were resuspended at the concentration of 5
× 10
5
cells/ml with fresh TCM complemented with 10 ng/ml recombinant human IL-2 and added
to the vector-loaded plates. The plates were spun at 1000×g at 32°C for 10 minutes and
incubated overnight at 37°C and 5% CO
2
. The same transduction procedure was repeated on the
following day. During ex vivo expansion, culture medium was replenished, and cell density was
adjusted to 5 × 10
5
/ml every two days.

55

Surface immunostaining and flow cytometry. To detect anti-CD19 CAR expression on the cell
surface, cells were stained with protein L. Before FACS staining, 5 × 10
5
cells

were harvested
and washed three times with FACS buffer (PBS containing 5% bovine serum albumin fraction
V). Cells were then stained with 1 µg of biotinylated protein L at 4°C for 30 minutes. Cells were
washed with FACS buffer three times and then incubated with 0.1 µg of FITC-conjugated
streptavidin in FACS buffer at 4°C for 10 minutes. Cells were washed and fixed with TransFix
cellular antigen stabilizing reagent (Thermo Scientific, Waltham, MA) at 4°C for 10 minutes.
Cells were then washed twice and stained with anti-CD3, anti-CD4, and anti-CD8 at 4°C for 10
minutes. Cells were washed and resuspended in PBS. Fluorescence was assessed using a
MACSquant cytometer (Miltenyi Biotec, San Diego, CA), and all the FACS data were analyzed
using FlowJo software (Tree Star, Ashland, OR).

Intracellular cytokine staining. T cells (1 × 10
6
)

were cultured with target cells at a ratio of 1:1
for 6 hours at 37°C and 5% CO
2
with GolgiPlug (BD Biosciences, San Jose, CA) in 96-well
round bottom plates. PE-Cy5.5-anti-CD3, FITC-anti-CD4, Pacific blue-CD8, PE-anti-IFN-γ and
PE-anti-Ki67 antibodies were used for the intracellular staining. Cytofix/Cytoperm Fixation and
Permeabilization Kit (BD Biosciences) was used to permeabilize the cell membrane and perform
intracellular staining according to the manufacturer’s instruction.

Western blotting analysis. Cell culture supernatant was harvested, and anti-PD-1 scFv was
purified with Pierce
TM
anti-HA magnetic beads (Thermo Scientific, Waltham, MA) according to
the manufacturer’s instruction. The purified antibody was then subjected to SDS-PAGE, and
transferred to a nitrocellulose membrane (Thermo Scientific, Waltham, MA) for Western blot

56

analysis. The Western blot was analyzed with anti-HA tag antibody (Abcam, Cambridge, MA) as
described previously (Xu et al., 2012).

ELISA. IFN-γ was measured using a human IFN-γ ELISA kit (BD Biosciences, San Jose, CA)
according to the manufacturer’s instructions. Briefly, 96-well ELISA plates (Thermo Scientific,
Waltham, MA) were coated with 200 ng/well of capture antibodies against the indicated proteins
at 4°C overnight. On the next day, plates were washed with wash buffer (PBS containing 0.05%
Tween 20) and blocked with assay buffer (PBS containing 10% FBS) for 2 hours at room
temperature. Equal volume of serum, or cell culture supernatant was added to the plate and
incubated for 2 hours at room temperature. Plates were then washed and incubated with detection
antibodies for 1 hour at room temperature. To measure anti-PD-1 antibody and secreted anti-PD-
1 scFv, recombinant human PD-1 (rhPD-1) was used to pre-coat the plate. Goat anti-mouse
IgG1-HRP and anti-HA tag antibodies were used as detection antibodies, respectively.

Competitive blocking assay. The 96-well assay plates (Thermo Scientific, Waltham, MA) were
coated with 3 µg/ml of anti-human CD3 antibody at 4°C overnight. On the second day, the
supernatant of the wells was aspirated and the wells were washed once with 100 µl per well of
PBS. 10 µg/ml of rhPD-L1/Fc (R&D Systems, Minneapolis, MN) in 100 µl of PBS were added.
In each well, 100 µg/ml of goat anti-human IgG Fc antibody in 10 µl of PBS were then added.
The assay plate was incubated for 4 hours at 37°C. Human T cells were harvested, washed once
and then resuspended to 1 × 10
6
cells/ml in TCM. The wells of the assay plate were aspirated.
Then, 100 µl of human T-cell suspension (1 × 10
5
) and 100 µl of supernatant of CAR or
CAR.αPD1 T cell culture 3-day post-transduction, supplemented with GolgiPlug (BD

57

Biosciences), were added to each well. The plate was covered and incubated at 37°C and 5%
CO
2
overnight. After incubation, T cells were harvested and stained with IFN-γ intracellularly.

Specific cell lysis assay. Lysis of target cells (H292-CD19) was measured by comparing the
survival of target cells to the survival of the negative control cells (NCI-H292). This method has
been described previously (Kochenderfer et al., 2009). NCI-H292 cells were labeled by
suspending them in R10 medium with 5 µM CellTracker Orange (5-(and-6)-(((4-
chloromethyl)benzoyl)amino)tetramethylrhodamine) (CMTMR), a fluorescent dye for
monitoring cell movement (Invitrogen, Carlsbad, CA), at a concentration of 1.5 × 10
6
cells/mL.
The cells were incubated at 37°C for 30 minutes and then washed twice and suspended in fresh
R10 medium. H292-CD19 cells were labeled by suspending them in PBS+0.1% BSA with 5 µM
Carboxyfluorescein succinimidyl ester (CFSE) fluorescent dye at a concentration of 1 × 10
6

cells/mL. The cells were incubated for 30 minutes at 37°C. After incubation, the same volume of
FBS was added into the cell suspension and then incubated for 2 minutes at room temperature.
The cells were then washed twice and suspended in fresh R10 medium. Equal amounts of NCI-
H292 and H292-CD19 cells (5 × 10
4
each) were combined in the same well for each culture with
effector CAR-T cells. Cocultures were set up in round bottom 96-well plates in triplicate at the
following effector-to-target ratios: 1:1 and 5:1. The cultures were incubated for 6 hours at 37°C,
followed by 7-AAD labeling, according to the manufacturer’s instructions (BD Biosciences).
Flow cytometric analysis was performed to quantify remaining live (7-AAD-negative) target
cells. For each coculture, the percent survival of H292-CD19 cells was determined by dividing
the percentage of live H292-CD19 cells by the percentage of live NCI-H292 cells. In the wells
containing only target and negative control cells without effector cells, the ratio of the percentage

58

of H292-CD19 cells to the percentage of NCI-H292 cells was calculated and used to correct the
variation in the starting cell numbers and spontaneous cell death. The cytotoxicity was
determined in triplicate and presented in mean ± SEM.

Cell proliferation. 3 × 10
5
H292-CD19 cells were suspended in D10 medium and then seeded in
a 6-well plate. Once the target cells attached, nontransduced T cells, CAR and CAR.aPD1 T cells
were harvested and washed twice with PBS. The cells were then labeled by suspending them in
PBS with 10 µM CFSE at a concentration of 1 × 10
6
cells/mL and incubated for 60 minutes at
37°C. After incubation, the cells were washed twice and suspended in fresh TCM. An equal
number of T cells were added to the target cells for coculture. Cocultures were set up in triplicate
at an effector-to-target ratio of 1:1. The cultures were incubated for 96 hours at 37°C. Flow
cytometric analysis was performed to quantify the intensity of CFSE on T cells. The proliferation
rates were determined in triplicate and presented in mean ± SEM.

Tumor model and adoptive transfer. At 6 to 8 weeks of age, mice were inoculated
subcutaneously with 3 × 10
6
H292-CD19 cells, and 10-13 days later, when the average tumor
size reached 100-120 mm
3
, mice were treated with i.v. adoptive transfer of 1 × 10
6
or 3 × 10
6

CAR transduced T cells in 100 µl PBS. CAR expression was normalized to 20% in both CAR
groups by addition of donor-matched nontransduced T cells. Tumor growth was monitored twice
a week. Tumor size was measured by calipers and calculated by the following formula: W
2
× L /
2. Mice were euthanized when they displayed obvious weight loss, ulceration of tumors, or
tumor size larger than 1000 mm
3
.

59

Statistical analysis. Statistical analysis was performed in GraphPad Prism, version 5.01. One-
way ANOVA with Tukey’s multiple comparison was performed to assess the differences among
different groups in the in vitro assays. Tumor growth curve was analyzed using one-way
ANOVA with repeated measures (Tukey’s multiple comparison method). Mouse survival curve
was evaluated by the Kaplan-Meier analysis (log-rank test with Bonferroni correction). A P
value less than 0.05 was considered statistically significant. Significance of findings was defined
as: ns = not significant, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001.

3.4 Results
3.4.1 Characterization of anti-CD19 CAR-T cells secreting anti-PD-1 antibody
The schematic representation of the retroviral vector constructs used in this study is shown in
Figure 3-1A. The retroviral vector encoding the anti-CD19 CAR composed of anti-CD19 scFv,
CD8 hinge, CD28 transmembrane and intracellular costimulatory domains, as well as
intracellular CD3ζ domain was designated as CAR19. The retroviral vector encoding both anti-
CD19 CAR and secreting anti-PD-1 scFv was designated as CAR19.αPD1. Human PBMCs were
transduced with each construct to test the expression of CAR in primary lymphocytes. As seen in
Figure 3-1B, CAR expression was observed for both constructs in human T cells, although anti-
PD-1-secreting CAR19 T cells expressed slightly lower level of the CAR on the cell surface.
Expression and secretion of anti-PD-1 was assessed by performing Western blotting analysis and
ELISA on the cell supernatant three days post-transduction. We observed that anti-PD-1 could be
successfully expressed and secreted by T cells transduced with CAR19.αPD1 (Figure 3-1C and
3-1D).

60

Figure 3-1 Construction and characterization of CAR19 and CAR19.αPD1.
(A) Schematic representation of parental anti-CD19 CAR (CAR19) and anti-PD-1-secreting anti-
CD19 CAR (CAR19.αPD1) constructs. (B) Expression of both CARs in human T cells. The two
groups of CAR T cells were stained with biotinylated protein L followed by FITC-conjugated
streptavidin to detect CAR expression on the cell surface. A viable CD3
+
lymphocyte gating
strategy was used. NT indicates nontransduced T cells, which were used as a control. (C, D)
Expression of secreted anti-PD-1 antibody in the supernatant from either CAR19 or
CAR19.αPD1 T cell culture was analyzed by Western blot (C) and ELISA (D). (E) The
percentage of CD8
+
T cells expressing IFN-γ over total CD8
+
T cells with the indicated treatment
(n=4, mean ± SEM; **P < 0.01).

61

To evaluate the binding activity and blocking function of anti-PD-1 scFv secreted by
CAR19.αPD1 T cells, a competitive binding and blocking assay was performed. Intracellular
IFN-γ was measured to assess the activity of the T cells. As shown in Figure 3-1E, the
expression of IFN-γ was upregulated when the T cells were stimulated by anti-CD3 antibody,
whereas the presence of recombinant human PD-L1 (rhPD-L1) resulted in significantly lower
IFN-γ expression. However, adding the cell culture supernatant from CAR19.αPD1 T cells
effectively reversed the inhibitory effect of rhPD-L1 on the T cells and significantly increased
IFN-γ production (Figure 3-1E).

3.4.2 Secreting anti-PD-1 antibody enhances the antigen-specific immune responses of
CAR-T cells
To further assess the effector function of anti-PD-1-secreting CAR19 T cells through antigen-
specific stimulation, both CAR19 and CAR19.αPD1 T cells were cocultured for different
durations with H292-CD19 or SKOV3-CD19 target cells, both of which were shown to have
high surface expression of PD-L1 (Supplementary Figure 3-1). T cells at different time points
were then harvested, and the cell function marker IFN-γ in the supernatant was measured by
ELISA. Upon antigen stimulation for 24 hours, we found that both CAR19 and CAR19.αPD1 T
cells, with or without secreting anti-PD-1, had a similar amount of IFN-γ secretion (Figure 3-2A
and Supplementary Figure 3-2). However, after 72 hours, CAR19.αPD1 T cells secreted
significantly higher IFN-γ compared to the parental CAR19 T cells after stimulation with H292-
CD19 cells (Figure 3-2A). Similarly, after 96 hours of antigen stimulation, CAR19 T cells
secreting anti-PD-1 expressed significantly more IFN-γ than that expressed by the parental
CAR19 T cells (Figure 3-2A and Supplementary Figure 3-2).

62

Next, the cytolytic function of engineered T cells was examined by a 6-hour cytotoxicity assay.
The cytotoxic activity of CAR19 and CAR19.αPD1 T cells against H292-CD19 cells was
evaluated at effector/target (E/T) ratios of 1, 5, 10 and 20. We found that both CAR19 and
CAR19.αPD1 T cells mediated significant cell lysis of target cells, especially at higher E/T ratios
in comparison with the nontransduced T cells. However, little difference was found between
CAR19 and CAR19.αPD1 T cells in terms of cytolytic activity (Figure 3-2B).

Supplementary Figure 3-1 The expression of PD-L1 on H292-CD19 and SKOV3-CD19.

63

Figure 3-2 Anti-PD-1 expression enhanced the antigen-specific immune responses of CAR
T cells.
(A) Both CAR19 and CAR19.αPD1 T cells were cocultured with H292-CD19 cells for different
durations. IFN-γ production was measured by ELISA (n=5, mean ± SEM; ns, not significant, P >
0.05; *P < 0.05). (B) Cytotoxicity of both CARs against target cells. The two groups of CAR T
cells were cocultured for 6 hours with H292-CD19 cells at 1:1, 5:1, 10:1, and 20:1 effector-to-
target ratios, and cytotoxicity against H292-CD19 was measured. Nontransduced (NT) T cells
were used as a control. (C) Proliferation of both CARs after antigen-specific stimulation. The
two groups of CAR T cells were pre-stained with CFSE. The stained T cells were then
cocultured for 96 hours with H292-CD19 cells at 1:1 effector-to-target ratio and the intensity of
CFSE was measured. Nontransduced (NT) T cells were used as a control. (D) The summarized
statistics of proliferation rate for nontransduced (NT) T cells, CAR19 T cells, and CAR19.αPD1
T cells in (C) were shown in bar graphs (n=4, mean ± SEM; *P < 0.05).

64

Supplementary Figure 3-2 IFN-γ production of CAR19 and CAR19.αPD1 T cells upon
antigen-specific stimulation.

T cell proliferation was then evaluated by a carboxyfluorescein diacetate succinimidyl ester
(CFSE)-based proliferation assay after 96-hour coculture of engineered T cells with target H292-
CD19 cells. We observed that antigen-specific stimulation of both CAR19 and CAR19.αPD1 T
cells resulted in a markedly higher level of proliferation compared to nontransduced T cells.
Moreover, compared to CAR19 T cells (57.9±10.2 %), the proliferation rate of CAR19.αPD1 T
cells (75.9±5.5 %) was significantly higher (Figure 3-2C and 3-2D).

3.4.3 Secreting anti-PD-1 alleviates CAR T cell exhaustion after antigen stimulation
PD-1 expression on human GD2 and mouse HER2 CAR T cells has been shown to increase
following antigen-specific activation, and PD-1 blockade was found to downregulate PD-1
expression in T cells (Gargett et al., 2016; John et al., 2013). To assess the effect of secreted anti-
PD-1 scFv on protecting human T cells from exhaustion, the engineered CAR T cells were
cocultured with either H292-CD19 or SKOV3-CD19 target cells for 24 hours and then stained
for the T cell exhaustion marker PD-1. We found that the expression of PD-1 was significantly

65

upregulated in both CAR19 and CAR19.αPD1 T cells following antigen-specific stimulation. In
comparison, the upregulated PD-1 expression on CAR19.αPD1 T cells was significantly lower
than that on parental CAR19 T cells (Figure 3-3A and 3-3B).
In addition to PD-1, other cell surface inhibitory molecules, including lymphocyte activation
gene 3 protein (LAG-3), T cell immunoglobulin domain and mucin domain-containing protein 3
(TIM-3; also known as HAVCR2) and cytotoxic T-lymphocyte associated protein 4 (CTLA-4),
also play important roles in inducing T cell exhaustion and limiting the antitumor efficacy of
CAR-T cell therapy (Pardoll, 2012). In order to evaluate whether the expression of other T cell
exhaustion markers is regulated by CAR stimulation, we measured the expression of LAG-3 and
TIM-3 on CAR-engineered T cells. Similar to PD-1, we found that the expression of LAG-3 and
TIM-3 was significantly upregulated on both CAR19 and CAR19.αPD1 T cells following
antigen stimulation, compared with nontransduced T cells. In comparison to CAR19 T cells,
CAR19.αPD1 T cells expressed slightly lower LAG-3 and TIM-3 after stimulation with H292-
CD19 cells. Moreover, upon SKOV3-CD19 stimulation, CAR19.αPD1 T cells had significantly
lower LAG-3 expression than CAR19 T cells, whereas they had similar TIM-3 expression
(Figure 3-3C and 3-3D).
It has been shown that PD-1 blockade could promote the survival of GD2 CAR T cells after
activation with the PD-L1-negative target cells, indicating that the interaction between PD-1-
expressing T cells and T cells expressing PD-1 ligands, such as PD-L1, might contribute to the
suppression of T cell function (Gargett et al., 2016). Thus, in this experiment, we also measured
the expression of PD-L1 in both CAR19 and CAR19.αPD1 T cells and found that it was

66

Figure 3-3 Secreting anti-PD-1 scFv protected CAR T cells from being exhausted.
Both CAR19 and CAR19.αPD1 T cells were cocultured with H292-CD19 cells for 24 hours. (A)
PD-1 expression was measured by flow cytometry. CD8
+
T cells were shown in each panel. PD-
1-expressing CD8 T cells were gated, and their percentage over total CD8
+
T cells was shown in
each scatterplot. (B) The summarized statistics of triplicates were shown in bar graphs (n=3,
mean ± SEM; **P < 0.01; ***P < 0.001). (C) LAG-3 expression was measured by flow
cytometry. The percentage of LAG-3-expressing CD8 T cells over total CD8
+
T cells was shown
in bar graphs (n=3, mean ± SEM; ns, not significant, P > 0.05; **P < 0.01). (D) TIM-3
expression was measured by flow cytometry. The percentage of TIM-3-expressing CD8 T cells
over total CD8
+
T cells was shown in bar graphs (n=3, mean ± SEM; ns, not significant, P >
0.05). (E) Both CAR19 and CAR19.αPD1 T cells were cocultured with either H292-CD19 or
SKOV3-CD19 cells for 24 hours. PD-L1 expression was measured by flow cytometry. The
percentages of PD-L1-expressing CD8 T cells over total CD8
+
T cells and PD-L1-expressing
CD4 T cells over total CD4
+
T cells were shown in bar graphs (n=3, mean ± SEM; *P < 0.05;
**P < 0.01; ***P < 0.001).

67

significantly increased following antigen-specific stimulation. However, the expression of PD-
L1 in CAR19.αPD1 T cells was significantly lower than that in CAR19 T cells (Figure 3-3E).

3.4.4 Anti-PD-1 engineered CAR T cells exhibit enhanced antitumor reactivity
To evaluate the antitumor efficacy of CAR19.αPD1 T cells, we adoptively transferred 1 × 10
6

CAR-engineered T cells into NSG mice bearing established H292-CD19 subcutaneous tumors
(~100 mm
3
). The experimental procedure for animal study is shown in Figure 3-4A. The data in
Figure 3-4B demonstrate that all three anti-CD19 CAR T cell groups showed decreased tumor
sizes compared to nontransduced T cells or nontransduced T cells combined with anti-PD-1
antibody treatment over the course of the experiment. However, in comparison to parental
CAR19 T cells or CAR19 T cells combined with anti-PD-1 antibody treatment, CAR19.αPD1 T
cell treatment significantly enhanced the antitumor effect, which became evident as early as one
week after T cell infusion (Figure 3-4B). Notably, 17 days after adoptive cell transfer, we
observed that the tumors from mice treated with CAR19.αPD1 T cells almost disappeared. In the
parental CAR19 T cell group or combination group, 4 out of 6 mice (~70%) still had either
progressive or stable disease states and only experienced a decrease in tumor size of less than
30% (Figure 3-4C). The overall survival of the tumor-bearing mice was also evaluated. It
showed that CAR19.αPD1 T cell treatment significantly prolonged long-term survival (100%),
compared to either the parental CAR19 T cell treatment alone (17%) or the combined anti-PD-1
antibody and CAR19 T cell treatment (17%) (Figure 3-4D).

68

Figure 3-4 Adoptive transfer of CAR T cells secreting anti-PD-1 scFv enhanced the growth
inhibition of established tumor.
(A) Schematic representation of the experimental procedure for tumor challenge, T cell adoptive
transfer and antibody treatment. NSG mice were s.c. challenged with 3 × 10
6
of H292-CD19
tumor cells. At day 20, when the tumors grew to ~100 mm
3
, 1 × 10
6
of CAR19 or CAR19.αPD1
T cells were adoptively transferred through i.v. injection. One day post-T cell infusion, anti-PD-
L1 antibody treatment was initiated, and the treatment was continued on the indicated dates.
Tumor volume was measured every other day. (B) Tumor growth curve for mice treated with
nontransduced (NT), NT plus anti-PD-1 injection, CAR19, CAR19 plus anti-PD-1 injection, or
CAR19.αPD1. Data were presented as mean tumor volume ± standard error of the mean (SEM)
at indicated time points (n = 8; *P<0.05; ***P < 0.001). (C) Waterfall plot analysis of tumor
reduction on day 17 post-therapy for various treatment groups. (D) Survival of H292-CD19
tumor-bearing NSG mice after indicated treatment. Overall survival curves were plotted using
the Kaplan-Meier method and compared using the log-rank (Mantel-Cox) test (n = 6; ns, not
significant, P > 0.05; *P<0.05; **P < 0.01).

0 5 10 15 20
0
100
200
300
400
500
600
NT
NT+αPD-1
CAR19
CAR19+αPD-1
CAR19.αPD1
***
*
*
Days post ACT
Tumor volume (mm
3
)
A
B
C
Tumor inoculation
H292-CD19, 3×10
6
Day -20 1 5
CAR-T, i.v.
1×10
6
Monitor tumor growth
9
Anti-PD-1 injection
(125 µg/mouse)
0 12
D
-100
-50
0
50
100
NT
NT+αPD-1
CAR
CAR+αPD-1
CAR-αPD-1
Subject
Change from baseline (%)
0 20 40 60 80
0
20
40
60
80
100
NT
NT+αPD-1
CAR
CAR+αPD1
CART-αPD-1
**
*
ns
Days post ACT
Percent survival
Figure 4

69

3.4.5 Anti-PD-1 engineered CAR T cells can expand more in vivo than parental CAR T
cells
Next, the engraftment and expansion of CAR T cells were assessed in vivo. Two days following
T cell infusion, mice were euthanized, and different organs and tissues, including the tumor,
blood, spleen and bone marrow, were harvested for human T cell staining. We found that T cells
in all groups had barely expanded and that less than 2% of T cells could be observed in all
examined tissues. Most T cells (1-2 %) homed to the spleen, while a certain percentage of T cells
(0.1-0.5 %) circulated were in the blood. The infiltration level of transferred T cells was low in
tumor and bone marrow. In addition, the T cell percentage between the nontransduced and CAR-
transduced T cells showed little difference across all examined tissues (Figure 3-5A). However,
one week post-T cell infusion, on day 10, we observed a significant expansion of CAR T cells in
all examined tissues, whereas nontransduced T cells were barely present. Notably, consistent
with our in vitro data, CAR19.αPD1 T cells had a significantly higher expansion rate compared
to parental CAR19 T cells, especially in tumor, spleen and blood (Figure 3-5B and 3-5C).

70

Figure 3-5 CAR T cells secreting anti-PD-1 were expanded more efficiently than parental
CAR T cells in vivo.
The percentage of human CD45
+
T cells in the tumor, blood, spleen and bone marrow of H292-
CD19 tumor-bearing mice that were adoptively transferred with nontransduced (NT), CAR19, or
CAR19.αPD1 T cells was investigated by flow cytometry at day 2 (A) or day 10 (B) post-therapy
(n = 3, mean ± SEM; *P<0.05; ***P < 0.001). (C) A representative FACS scatter plot of the
percentage of human CD45
+
T cells in the tumor, blood, spleen and bone marrow of different
groups.

3.4.6 Anti-PD-1 engineered CAR T cells lead to reversal of T cell exhaustion and higher T
cell effector function at the established tumor site
To further determine if the enhanced antitumor effects observed following CAR19.αPD1 T cell
therapy are correlated with increased function of CAR T cells at the tumor site, mice were
challenged with H292-CD19 tumors before receiving 3 × 10
6
CAR T cells. The experimental

71

design is shown in Figure 3-6A. Eight days after T cell infusion, we euthanized the mice and
analyzed T cells in tumor, blood, spleen and bone marrow, using flow cytometry. Compared to
the CAR T cell treatment, we observed that the injected anti-PD-1 antibody had little effect on
enhancing the expansion of T cells in vivo. However, consistent with our previous observation
(Figure 3-5B), T cells from mice treated with the CAR19.αPD1 regimen expanded at a higher
rate in tumor, blood, and spleen (Figure 3-6B). It has been shown that the population of
cytotoxic CD8
+
T cells among tumor-infiltrating lymphocytes (TILs) is critical in eliciting
antitumor immunity and spontaneous tumor control (Hadrup et al., 2013). Therefore, the ratio of
CD8
+
versus CD4
+
T cells was analyzed among TILs. Compared to the parental CAR19 T cells,
results showed that the CAR19.αPD1 T cells had a significantly higher ratio of CD8
+
versus
CD4
+
T cells, whereas the combined therapy had a similar CD8
+
versus CD4
+
T cell ratio
compared to CAR T cell monotherapy (Figure 3-6C). Further, we assessed PD-1 expression on
tumor-infiltrating CD8
+
T cells and found that both the injected and secreted anti-PD-1
antibodies could significantly decrease the expression of PD-1 (Figure 3-6D). We also
performed the ex vivo culture and activated TILs with either anti-CD3/CD28 antibodies or target
cell H292-CD19. We observed significantly higher expression of IFN-γ in adoptively transferred
CAR19.αPD1 T cells, compared to either parental CAR19 T cells or CAR19 T cells combined
with systemic anti-PD-1 antibody treatment. Little difference was observed in IFN-γ expression
between CAR T cell monotherapy and combined therapy (Figure 3-6E and 3-6F). Additionally,
we measured the expression of IFN-γ and anti-PD-1 antibodies in the sera and found little
difference in IFN-γ expression among all groups (Supplementary Figure 3-3A). Notably,
compared to CAR19 T cell treatment, CAR19.αPD1 T cell therapy had significantly higher anti-

72

PD-1 concentration in the sera, although the concentration was more than 15-fold lower than that
with systemic anti-PD-1 antibody injection (Figure 3-6G).

Figure 3-6 CAR T cells secreting anti-PD-1 were more functional than parental CAR T
cells at local tumor site.
(A) Schematic representation of the experimental procedure for tumor challenge, T cell adoptive
transfer and antibody treatment. NSG mice were s.c. challenged with 3 × 10
6
of H292-CD19
tumor cells. At day 20, 3 × 10
6
of CAR19 or CAR19.αPD1 T cells were adoptively transferred
through i.v. injection. One day post-T cell adoptive transfer, anti-PD-1 antibody treatment was
initiated, and the treatment was continued on the indicated dates. The mice were then euthanized
on day 8 for analysis. (B) The percentage of human CD45
+
T cells in the tumor, blood, spleen
and bone marrow of H292-CD19 tumor-bearing mice that were adoptively transferred with
CAR19 or CAR19.αPD1 T cells, or treated with CAR19 T cells along with injection of anti-PD-
1 antibody, was investigated by flow cytometry. (C) The ratio of CD8
+
versus CD4
+
TILs in the
tumor (n = 3, mean ± SEM; ns, not significant, P>0.05; *P<0.05; ***P < 0.001). (D) The

73

percentage of PD-1-expressing CD8 TILs over total CD8
+
TILs (n=3, mean ± SEM; *P < 0.05).
TILs were harvested and stimulated ex vivo for 6 hours by either anti-CD3/anti-CD28 antibodies
(E) or target cells H292-CD19 (F). The percentage of CAR T cells in the tumor expressing
intracellular IFN-γ was investigated by flow cytometry (n = 3, mean ± SEM; *P<0.05; **P <
0.01). (G) The secreted anti-PD-1 scFvs and injected anti-PD-1 antibodies in the sera were
evaluated using ELISA (n = 3, mean ± SEM; **P<0.01; ***P < 0.001).

Supplementary Figure 3-3 The expression of IFN-γ in the sera was measured by ELISA.

3.5 Discussion
Adoptive T cell therapy has become a promising method of immunotherapy. It has achieved
successful responses in patients with hematopoietic malignancies. However, the outcome has
been less promising in the treatment of solid tumors, partly owing to the immunosuppressive
properties and establishment of an immunosuppressive microenvironment (Vazquez-Cintron et
al., 2010). The PD-1/PD-L1 regulatory pathway has demonstrated particularly antagonistic
effects on the antitumor response of TILs. Solid tumors with poor prognosis showed
upregulation of PD-L1 expression, while TILs were shown to have PD-1 upregulation (Wu et al.,
2015). The combined effect of these two results in tumor escape. However, this can be disrupted
by the use of checkpoint inhibitors (CPIs) targeting the PD-1/PD-L1 pathway (Ansell et al.,

74

2015; Brahmer et al., 2010; Dong et al., 2002). As a result, the ensuing research was designed to
investigate the effects of PD-1/PD-L1 blockade in infused CAR T cells, which showed
upregulation of PD-1 after activation (Moon et al., 2014).
Despite other methods of PD-1/PD-L1 inhibition, such as cell intrinsic PD-1 shRNA and PD-1
dominant negative receptor (Cherkassky et al., 2016), treatment with PD-1 or PD-L1 antibody
has long been a topic of interest and extensively studied in both animal models and clinical trials.
Indeed, both antibodies have resulted in a marked inhibition of tumor growth. However, antibody
treatment has multiple limitations. For example, it requires multiple and continuous antibody
administration to obtain a sustained efficacy (Topalian et al., 2012). Also, the large size of
antibodies prevents them from entering the tumor mass and encountering the infiltrated PD-1-
positive T cells (Beckman et al., 2007; Chames et al., 2009). To account for these inefficiencies,
multiple high-dose treatments with immunomodulatory drugs or antibodies are required, but this
can result in side effects that range from mild diarrhea to autoimmune hepatitis, pneumonitis and
colitis. Moreover, it has been shown that the Fc portion of antibodies may cause immune cell
depletion by activating cytotoxic signals within macrophages and natural killer cells, which
usually express FcαRI and FcγRIIIA/FcγRIIC, respectively (Keler et al., 2000; Maute et al.,
2015; Wang et al., 2015a). Therefore, in this study, we focused our efforts on engineering CAR
T cells to secrete and deliver high concentrations of human scFvs against PD-1, aiming to change
the immunosuppressive tumor microenvironment, prevent tumor-induced hypofunctionality and
enhance the antitumor immunity of infused CAR T cells.
Herein, we engineered human anti-CD19 CAR T cells that secrete human anti-PD-1 scFvs and
demonstrated that anti-PD-1 scFv could be efficiently expressed and secreted by CAR19.αPD1 T
cells. The secreted scFvs successfully bound to PD-1 on the cell surface and reversed the

75

inhibitory effects of PD-1/PD-L1 interaction on T cell function. PD-1 blockade by constitutively
secreted anti-PD-1 scFv decreased T cell exhaustion and significantly enhanced T cell
proliferation and effector function in vitro. Our study using xenograft mouse models also
demonstrated that CAR19.αPD1 T cells, when compared to parental CAR19 T cells, further
enhanced antitumor activity and prolonged overall survival. Mechanistically, we observed that
CAR19.αPD1 T cells had greater in vivo expansion. In addition, at the local tumor site,
CAR19.αPD1 T cells were shown to be less exhausted and more functional than parental CAR19
T cells.
The engagement of PD-1 and its ligand PD-L1 or PD-L2 transduces an inhibitory signal and
suppresses T cell function in the presence of TCR or BCR activation (Chemnitz et al., 2004;
Koyama et al., 2016; Riella et al., 2012). In this study, the presence of recombinant human PD-
L1 protein (rhPD-L1) significantly inhibited T cell activation in an in vitro activation assay. To
examine the binding and blocking activity of anti-PD-1 scFv secreted by CAR19.αPD1 T cells,
we cultured the T cells with cell culture supernatant from either CAR19 T cells or CAR19.αPD-1
T cells in the presence of rhPD-L1 protein. We observed that the supernatant from CAR19.αPD1
T cells rescued T cell function and significantly increased IFN-γ production, indicating that
secreted anti-PD-1 could successfully bind to PD-1 and reverse the inhibitory effects of the PD-
1/PD-L1 interaction on T cell function.
The PD-1/PD-L1 pathway involves the regulation of cytokine production by T cells, inhibiting
production of IFN-γ, TNF-α and IL-2 (Riella et al., 2012). PD-1 expression of human GD2 and
anti-HER2 CAR T cells has been shown to increase following antigen-specific activation, and
PD-1 blockade has been shown to enhance T cell effector function and increase the production of
IFN-γ in the presence of PD-L1
+
target cells (Gargett et al., 2016; John et al., 2013). Therefore,

76

in this study, to compare the functional capacity of CAR19 T and CAR19.αPD1 T cells, we
cocultured T cells with a PD-L1
+
cancer cell line, H292-CD19 or SKOV3-CD19, and found that
the anti-PD-1-secreting CAR19 T cells produced a significantly higher level of IFN-γ than
parental CAR19 T cells. In addition to cytokine production, PD-1 can also inhibit T cell
proliferation (Keir et al., 2008). With CAR-specific stimulation in the presence of PD-L1
+
cancer
cells, we found that CAR19.αPD1 T cells had a significantly higher proliferation rate than the
parental CAR19 T cells. Taken together, these data imply that PD-1/PD-L1 signaling blockade
results in more functional CAR19.αPD1 T cells with higher proliferation capacity compared to
CAR19 T cells alone.
To better understand how secreted anti-PD-1 affects the function of CAR19.αPD1 T cells, we
exposed CAR19 T cells and CAR19.αPD1 T cells to PD-L1
+
target cells and examined the
expression of T cell exhaustion markers, including PD-1, LAG-3 and TIM-3. We observed
significantly lower PD-1 expression on CAR19.αPD1 T cells, as well as lower expression of
other exhaustion markers, such as LAG-3, compared with parental CAR19 T cells. The
decreased expression of PD-1 in CAR19.αPD1 T cells may be caused by the dual effects of
antibody blockade and downregulation of PD-1 surface expression (Gargett et al., 2016; John et
al., 2013). PD-1 upregulation on tumor-infiltrating T cells was reported to be a major contributor
to T cell exhaustion in high PD-L1-expressing tumors. Downregulation of PD-1 may contribute
to reversion of T cell exhaustion and enhanced T cell effector function, which is supported by
increased IFN-γ production of CAR19.αPD1 T cells. In addition, the lower expression level of
other exhaustion makers, such as LAG-3, may also contribute to the higher function of
CAR19.αPD1 T cells upon antigen stimulation. Our observation is consistent with a recent study,
demonstrating that co-expression of multiple inhibitory receptors is a cardinal feature of T cell

77

exhaustion (Thommen et al., 2015; Wherry and Kurachi, 2015). Moreover, we found that PD-L1
expression was significantly increased on CAR T cells with antigen-specific stimulation, which
may also contribute to T cell exhaustion through T cell - T cell interaction. Notably, in
comparison, we observed that the expression level of PD-L1 on CAR19.αPD1 T cells was
significantly lower. These data suggest that the inhibited upregulation of PD-1 and PD-L1
expression on CAR19.αPD1 T cells may contribute to the reduction of tumor cell-induced and/or
T cell-induced exhaustion, thereby further enhancing T cell effector function and its antitumor
immunity.
Our in vivo study showed that the tumor growth could be inhibited by CAR T cell treatment,
irrespective of PD-1/PD-L1 blockade. Compared to CAR19 T cell treatment or combined
CAR19 T cell and systemic anti-PD-1 antibody treatment, in which 67% of the mice still had
either stable or progressive disease, we observed that CAR19.αPD1 T cell treatment achieved
more than 90% tumor eradication in about two weeks. To understand the underlying mechanism
of enhanced antitumor efficacy of CAR19.αPD1 T cells, we analyzed the expansion of
adoptively transferred T cells in vivo. Consistent with our in vitro data, we found that the anti-
PD-1-secreting CAR T cells were expanded significantly more than parental CAR T cells in all
examined tissues, including tumor, blood, spleen and bone marrow. Moreover, the population of
cytotoxic CD8
+
T cells among TILs is critical in eliciting antitumor immunity (Hadrup et al.,
2013). A previous study demonstrated that PD-1 signaling is involved in regulating the
expansion and function of CD8
+
TILs (Chauvin et al., 2015). In this study, the larger population
of CD8
+
TILs expresses IFN-γ

when stimulated ex vivo and the higher ratio of CD8
+
versus CD4
+

TILs in the CAR19.αPD1 T cell group implies that CAR19.αPD1 T cells are more functional and
expandable in vivo compared to parental CAR19 T cells.

78

Interestingly, in this study, we demonstrated that systemic anti-PD-1 antibody injection has little
effect on enhancing the antitumor efficacy of CAR T cell therapy. In a syngeneic HER2
+
self-
antigen tumor model, recent studies have demonstrated that a high-dosage (250 µg/mouse of
anti-PD-1 antibody) PD-1 blockade was capable of enhancing the antitumor activity of anti-
HER2 CAR T cells in the treatment of breast cancer (John et al., 2013). However, a lower
dosage (200 µg/mouse) of anti-PD-1 antibody showed a limited effect on CAR T cell therapy
(Beavis et al., 2017). In the present study, with a low-dose (125 µg/mouse) injection, the anti-
PD-1 antibody failed to inhibit tumor growth or enhance the antitumor efficacy of CAR T cells.
This observation indicates that a large dose of anti-PD-1 antibody, which often causes systemic
toxicity, may be required to achieve substantial antitumor efficacy. We measured the amount of
circulating anti-PD-1 antibodies and found a significant amount of circulating injected antibody
(~0.7 µg/ml) in the combination treatment group and a 15-fold lower amount in the
CAR19.αPD1 T cell treatment group. Although both administered and self-secreting anti-PD-1
antibodies efficiently decreased and blocked the PD-1 expression in CD8
+
T cells in vivo,
systemically injected anti-PD-1 antibody had little effect on increasing the population of
cytolytic CD8
+
TILs or enhancing IFN-γ production of TILs upon ex vivo stimulation. This result
suggests that the injected antibody has little effect on augmenting infused T cell function at the
present dose. It also explains our observed failure of injected PD-1 blockade in enhancing the
antitumor activity of CAR T cell therapy. Given the low concentration of secreted anti-PD-1 and
the augmented effector function at the local tumor tissue, the anti-PD-1 secreted by CAR T cells
may provide a safer and more potent approach in blocking PD-1 signaling and enhancing the
functional capacity of CAR T cells.

79

3.6 Conclusion
In summary, CAR19.αPD1 T cells exhibited alleviated T cell exhaustion, enhanced T cell
expansion, and improved CAR T cell treatment of human solid tumors in a xenograft mouse
model. In an immune competent condition, we speculate that anti-PD-1-engineered CAR T cells
might be more powerful in inducing tumor eradication given the durable effect of PD-1 blockade
on modulating the tumor microenvironment (Pardoll, 2012; Santarpia and Karachaliou, 2015). In
addition, we foresee that engineering the anti-PD-1 scFv into CAR constructs targeting other
tumor-associated antigens, such as mesothelin or HER-2 for the treatment of ovarian cancer or
breast cancer, which usually have high PD-L1 expression, is among the next steps that should be
explored to achieve better antitumor immunotherapy.

80

Chapter 4. Inhibition of Stat3 Enhances Antitumor Activity of PD-
L1 Blockade in a Murine Melanoma Model

81

4.1 Abstract
Melanoma, a common and highly immunogenic malignancy, is difficult to treat despite
advances in such multimodal approaches as surgery, radiation, targeted therapy,
chemotherapy and immunotherapy. Recent clinical trials with anti-PD-1 and anti-PD-L1
monoclonal antibodies have shown notable responses in patients with metastatic
melanoma. However, the overall response rate is still low. Therefore, seeking
combination therapy with enhanced efficacy of tumor eradication mediated by immune
checkpoint blockade remains an urgent task. In this report, we show that PD-L1 blockade
significantly increases Stat3 phosphorylation, a critical factor shown to drive tumor
progression and immune evasion, including modulation of tumor-mediated immune
suppression. By combining BP-1-102, a Stat3 inhibitor, and anti-PD-L1 antibody, we
demonstrate a significant inhibition of tumor growth, compared to either treatment alone.
Mechanistically, the combined therapy enhances cytotoxic CD8
+
T and helper CD4
+
T
cells and decreases regulatory FoxP3
+
T cells infiltrating to tumor tissue. At the
molecular level, the combined therapy results in a significant increase of IFN-γ, TNF-α,
and TNF-related apoptosis-inducing ligand (TRAIL) expression, but a decrease of VEGF
expression, within tumor tissue. Taken together, our data suggest that Stat3 inhibition by
the small-molecule inhibitor BP-1-102 can markedly enhances the antitumor potency of
PD-L1 blockade, improving antitumor immunity through the enhancement of both tumor
microenvironment modulation and local immune responses.

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4.2 Introduction
Melanoma is the fifth leading malignancy in males and the sixth in females worldwide
(Siegel et al., 2012). Current treatment for advanced melanoma consists of surgery,
radiation, targeted therapy (e.g., Imatinib (Hodi et al., 2013) and Nilotinib (Cho et al.,
2012)), chemotherapy (e.g., Dacarbazine (Serrone et al., 2000) and Temozolomide
(Quirbt et al., 2007)), and immunotherapy (e.g., Ipilimumab (Lipson and Drake, 2011)
and IL-2 (Atkins et al., 2000)). Despite recent advances in the treatment of melanoma, the
general survival rate for patients with advanced melanoma is still low. The five-year
survival rate for stage III and IV melanoma ranges from 40% to 78% and 15% to 20%,
respectively (Balch et al., 2009), making melanoma the major cause of death in patients
with skin cancer.
PD-1, one of several important inhibitory receptors characterized in T cells, is
upregulated shortly after T cell activation, promoting T cell dysfunction and exhaustion
via interacting with its cognate ligands, PD-L1 or PD-L2. PD-L1 is constitutively
expressed on T cells, B cells, macrophages, and dendritic cells (DCs) (Yamazaki et al.,
2002). In addition to hematopoietic lineage cells, PD-L1 is also abundant in a wide
variety of solid tumors, including melanoma, ovarian and lung cancer (Brown et al.,
2003; Dong et al., 2002; Konishi et al., 2004). In contrast, the expression of PD-L1 in
normal tissues is undetectable (Dong et al., 2002). Unlike other immune checkpoints, the
major role of PD-1/PD-L1 signaling is to limit the activity of T cells in the process of
inflammatory responses and protect tissues from autoimmune attack. In the case of
cancer, this translates into a major immune resistance mechanism in the tumor
microenvironment (Pardoll, 2012). Additionally, in the tumor microenvironment, PD-L1

83

expression can be induced in most tumor cells in response to T cell activation and
interferons (IFNs) (Kronig et al., 2014; Schoop et al., 2004; Wilke et al., 2011),
implicating that the increased expression of PD-L1 may be one of the major mechanisms
of immune resistance in tumor cells.
As a consequence of its significant role in inducing T cell exhaustion and tumor immune
resistance, the PD-1/PD-L1 pathway has been recognized as an attractive therapeutic
target for cancer immunotherapy. Recent clinical trials with anti-PD-1 and PD-L1
monoclonal antibodies have shown notable responses in patients with various cancers,
including metastatic melanoma (Brahmer et al., 2012; Hamanishi et al., 2015; Rizvi et al.,
2015; Robert et al., 2015). However, only 20-40% of the treated patients respond to the
current immunotherapy treatment (Swaika et al., 2015). Therefore, it is necessary to
elucidate the underlying mechanism of immune resistance in order to find a combination
therapy able to enhance the efficacy of tumor eradication, as mediated by immune
checkpoint blockade.
Signal transducer and activator of transcription 3 (Stat3), first discovered and identified
as a DNA-binding factor in 1993 (Wegenka et al., 1993), mediates responses of cytokines
and growth factors. Ligand binding induces the phosphorylation and activation of Stat3
via intrinsic or receptor-associated tyrosine kinases, including Jak2 and Src (Yu et al.,
2009). Aberrant activation of Stat3 has been observed in most human cancers, including
ovarian, breast, colorectal, brain, lung, melanoma, cervical, and prostate cancers
(Debnath et al., 2012). The mechanisms of Stat3-mediated malignant transformation and
tumorigenesis include dysregulating the expression of its target genes, enhancing tumor
angiogenesis (Niu et al., 2002), and magnifying metastasis (Abdulghani et al., 2008).

84

Interestingly, in addition to its direct tumorigenic properties, Stat3 was found to be a key
regulator of immunosuppression in cancers (Yu et al., 2007). Stat3-deficient CD8
+
T
cells were shown to exhibit faster proliferation and more sensitivity to activation-induced
cell death (Yu et al., 2013). Upon investigating the association between Stat3 and
regulatory T cells (Tregs), it was revealed that Stat3 ablation inhibits FoxP3 expression
and the suppressive function of Tregs (Pallandre et al., 2007), suggesting a direct role of
Stat3 in regulating Treg function. Disrupting the Stat3 signaling pathway enhances local
inflammatory responses by increasing the expression profile of cytokines/chemokines
(Cheng et al., 2003). In addition, Stat3 was found to constitutively occupy the proximal
region of the PD-L1 promoter and regulate PD-L1 expression (Wang et al., 2015b;
Wolfle et al., 2011). Thus, Stat3 seems to be an attractive therapeutic target for tumor
progression and immune evasion.
Accordingly, in this study, we reported that PD-1/PD-L1 blockade with anti-PD-L1
antibody had only limited efficacy in an established murine melanoma model. At the
same time, the expression of Stat3 phosphorylation was significantly increased. However,
when Stat3 phosphorylation and activation were inhibited by BP-1-102, the antitumor
potency of PD-L1 blockade was improved, as well as its antitumor immunity, through
modulating the tumor microenvironment and local immune responses. These results
indicate that the Stat3 signaling pathway may contribute to immune evasion in
checkpoint blockade treatment. Therefore, this study reports the rational design of a
combinational therapy involving immune checkpoint blockade and Stat3 inhibition in the
treatment of melanoma.

85

4.3 Materials and methods
Reagent. BP-1-102 was purchased from Millipore. Stock solutions of 80 mmol/L (67.5
mg/mL) were prepared in dimethyl sulfoxide (DMSO) and stored at −20°C. Further
dilutions were freshly made in cell culture medium or PBS. Anti-p-Stat3, anti-Stat3, and
anti-actin antibodies were purchased from Cell Signaling Technology. Anti-PD-L1
antibody for Western blotting was purchased from Santa Cruz Biotechnology.

Mice. C57BL/6 mice (Charles River Laboratories) were used for in vivo efficacy studies.
Mice were fed ad libitum and kept in air-conditioned rooms at 20°C ± 2°C with a 12-hour
light–dark period. Animal care and manipulation were in conformity with USC
institutional guidelines, which were in accordance with the Guidelines for the Care and
Use of Laboratory Animals.

Cell culture. B16 melanoma cells (B16-F10, ATCC number: CRL-6475) were
maintained in a 5% CO
2
environment with Dulbecco’s modified Eagle’s medium
supplemented with 10% FBS, 2 mM L-glutamine, 100 U/ml penicillin and 100µg/ml
streptomycin. All cell culture media and additives were purchased from Hyclone.

Western blotting. Cells or xenograft tumor tissues were harvested and minced to single
suspension cells, washed with ice-cold PBS, and lysed in cold lysis buffer (20 mmol/L
Tris-HCl, 150 mmol/L NaCl, 1 mmol/L EDTA, 1% Triton X-100, pH 7.5) with 1×
protease and phosphatase inhibitors. Protein concentration was determined by BCA
protein assay (Thermo Scientific). Proteins were resolved on 8% or 10% SDS-PAGE and

86

electrotransferred to Immun-Blot PVDF membrane (Bio-Rad). After blocking with 5%
milk in Tris-buffered Saline with Tween-20 (TBST), membranes were first probed with
the indicated primary antibodies, subsequently probed with horseradish peroxidase–
conjugated secondary antibody, and then developed using Dura Extended Duration
Substrate (Thermo Scientific). Immunoreactive proteins were visualized with the Chemi-
Doc System (Bio-Rad).

Surface immunostaining and flow cytometry. Tumor tissue from treated mice was
harvested, minced to single suspension cells, and filtered through 0.7 µm nylon strainers
(BD Biosciences). The cells from five different mice were then mixed together and
purified by Percoll density-gradient separation. The purified cells were washed twice
with cold PBS and then incubated for 10 minutes at 4 °C with rat anti-mouse
CD16/CD32 mAbs (BD Biosciences) to block nonspecific binding. Next, cells were
stained with monoclonal antibodies conjugated with fluorescent dyes. All staining
antibodies and isotype controls were purchased from eBioscience or BioLegend,
including anti-CD45 (30-F11), anti-CD3 (145-2C11), anti-CD4 (RM4-5), anti-CD8 (53–
6.7), anti-CD25 (PC61), anti-FoxP3 (FJK-16S), anti-CD11c (N418), and anti-PD-L1
(10F.9G2). Tregs were identified by CD45
+
CD3
+
CD4
+
CD25
+
FoxP3
+
markers; CD4 and
CD8 T cells were identified by CD45
+
CD3
+
CD4
+
and CD45
+
CD3
+
CD8
+
markers,
respectively. Dendritic cells were identified by CD45
+
CD11c
+
markers. Data were
acquired on a MACSquant cytometer (Miltenyi Biotec), and the analysis was performed
using FlowJo software (Tree Star).

87

RNA extraction, RT-PCR and real-time qPCR. Total RNA was extracted using the
RNeasy Mini Kit (Qiagen) according to the manufacturer’s instructions. Two µg of total
RNA were used for cDNA synthesis, which was performed using the High-Capacity
cDNA Reverse Transcription Kit (Applied Biosystems) in accordance with the
manufacturer’s instructions. Real-time qPCR was carried out with the appropriate
primers using TaqMan® Real-Time PCR Master Mix (Thermo Scientific). Reactions
were run in triplicate in three independent experiments. The geometric mean of
housekeeping gene GAPDH was used as an internal control to normalize variability in
expression levels. Expression data were normalized to the geometric mean of
housekeeping gene GAPDH to control the variability in expression levels and were
analyzed using the 2
-ΔΔCT
method.

Cell proliferation assay. Cell proliferation was assessed using an XTT assay. Cells were
seeded in 96-well microtiter plates and allowed to attach 24 hours before the addition of
corresponding compounds to the culture medium. After 72 hours, cells were incubated
with 0.3 mg/mL XTT (Amresco) for an additional 3 hours at 37°C. After incubation, the
absorbance was read at 490 nm, with a reference wavelength at 650 nm. All assays were
conducted in triplicate. Percentage of cell growth inhibition was expressed as follows: (1
− A/C) × 100%, where A and C were the absorbance values from experimental and
control cells, respectively. Inhibitory concentration 50% (IC
50
) values were determined
for each drug from a plot of drug concentration versus percentage of cell growth
inhibition.

88

Trypan Blue assay. Cell viability was assessed using trypan blue dye exclusion assay.
B16-F10 cells were seeded in 6-well plates and allowed to attach 24 hours before the
addition of corresponding compounds to the culture medium. After 24 hours incubation,
cells were harvested. Each cell suspension was mixed with an equal volume of 0.4%
trypan blue solution for 5 min. Viable cells were then counted on a hemocytometer under
a light microscope.

Annexin V apoptosis assay. Cell apoptosis was measured using Annexin V dye
exclusion assay according to the manufacturer’s instructions (BD Biosciences). B16-F10
cells were seeded in 6-well plates and allowed to attach 24 hours before the addition of
corresponding compounds to the culture medium. After 24 hours incubation, cells were
harvested, washed once with PBS, and then stained with Annexin V and 7-AAD.

In vivo studies. B10-F10 cells in the logarithmic growth phase from in vitro cultures
were implanted in C57BL/6 mice (3 × 10
6
cells in 100 µL of PBS/mouse) under aseptic
conditions. Tumor growth was assessed by daily measurement of tumor diameters with a
Vernier caliper. Tumor volume was calculated according to the following formula:
Tumor volume (mm
3
) = L × W× H. For intraperitoneal (i.p.) administration, tumors were
allowed to grow to an average volume of 50 mm
3
. Mice were then randomly assigned
into different groups, 5 mice per group, for daily vehicle control, BP-1-102 (6 mg/kg),
anti-PD-L1 antibody or combination treatment. To prepare a BP-1-102 solution for i.v.
administration, 30 mg/mL DMSO stock solution of BP-1-10 were diluted to 1.2 mg/mL
in PBS. Body weight and tumor volume were measured every other day.

89

4.4 Results
4.4.1 PD-L1 blockade enhances Stat3 phosphorylation
To assess the antitumor efficacy of PD-L1 blockade in melanoma therapy, we injected
mice bearing melanoma with anti-PD-L1 antibody intraperitoneally every three days for
four continuous treatments. We found that anti-PD-L1 monotherapy inhibited tumor
growth, but with limited efficacy compared to vehicle treatment (Figure 4-1A). Given
the significant role of Stat3 in inducing immunosuppression and tumor evasion, we
assessed the activation of Stat3 by measuring Stat3 phosphorylation through Western
blotting analysis of the tumor tissue. We observed that PD-L1 blockade significantly
increased the phosphorylation of Stat3 within tumor (Figure 4-1B and 4-1C).

4.4.2 BP-1-102 inhibits Stat3 activation and induces apoptotic cell death in murine
melanoma cells
BP-1-102, the molecular structure of which is shown in Figure 4-2A, is an orally
bioavailable inhibitor of Stat3 able to specifically bind to Stat3 SH2 domain and inhibit
its phosphorylation (Tyr705) and activation (Zhang et al., 2012). Consistent with the
previous observation that BP-1-102 decreased the phosphorylation of Stat3 in breast
cancer and pancreatic cancer cells (Zhang et al., 2012), our results showed that BP-1-102
inhibited Stat3 phosphorylation (Tyr705) in B16-F10 melanoma cells in a dose-
responsive manner (Figure 4-2B). Stat3 has been shown to bind to PD-L1 promoter and
regulate its expression (Wolfle et al., 2011). In our study, we observed that inhibition of
Stat3 by BP-1-102 inhibited PD-L1 expression in B16-F10 cells (Figrue 4-2C). Given

90

the critical roles of Stat3 in cell survival and proliferation (Hirano et al., 2000), MTT
assay and trypan blue assay were performed to evaluate the inhibitory effect on
proliferation and cytotoxicity of BP-1-102 in B16-F10 cells. We found that BP-1-102
inhibited cell proliferation and induced cell death in a dose-responsive manner with IC
50

values in micromolar range (Figrue 4-2D and 4-2E). To further evaluate the effect of

Figure 4-1 PD-L1 blockade increases Stat3 phosphorylation.
(A) Tumor growth curve of transplanted B16-F10 cells in C57BL/6 mice treated with
vehicle or anti-PD-L1 antibody. Data were presented as mean tumor volume ± standard
error of the mean (SEM) at indicated time points (n = 5; *P<0.05). (B) The expression of
Stat3 and p
Tyr705
-Stat3 in tumor tissues from mice treated with vehicle or anti-PD-L1
antibody. (C) Quantification of protein abundance of Stat3 and p-Stat3. Data are
presented as a ratio of phosphorylated-to-total Stat3 and are representative of three
independent experiments. (n = 3, error bars indicate SEM; *P<0.05).

91

Figure 4-2 BP-1-102 inhibits Stat3 phosphorylation and induces apoptotic cell death.
(A) Chemical structure of BP-1-102. (B) The expression of Stat3 and p
Tyr705
-Stat3 in
B16-F10 murine melanoma cells treated with various concentrations of BP-1-102 for 1 h.
Data are representative of three independent experiments. (C) The expression of PD-L1
in B16-F10 cells treated with various concentrations of BP-1-102 for 24 h. β-actin was
measured as a control. Data are representative of three independent experiments. (D)
Analysis of BP-1-102-induced cell proliferation inhibition in B16-F10 cells. After
treatment with various concentrations of BP-1-102 for 72 h, cells were subjected to XTT
assay. (E) BP-1-102 induces cell death in B16-F10 cells. After treatment with the
indicated concentrations of BP-1-102 for 24 h, cells were stained with trypan blue.
Histogram shows the percentage of trypan blue-stained cells after indicated treatment. (F)
BP-1-102 causes cell apoptosis in B16-F10 cells. After treatment with BP-1-102 for 24 h,
cells were stained with Annexin V-PE and 7-AAD and then analyzed by flow cytometry.

92

BP-1-102 on cell apoptosis, Annexin V apoptosis assay was performed on B16-F10 cells
treated with various concentrations of BP-1-102 for 24 hours. We found that BP-1-102
treatment significantly induced apoptotic cell death at 10 µM (Figure 4-2F).

4.4.3 BP-1-102 enhances the infiltration of mature DCs and cytotoxic CD8
+
T cells to
local tumor tissue
Since Stat3 plays a vital role in regulating tumor microenvironment and inducing
immunosuppression (Yu et al., 2007), we suspected that BP-1-102 could modulate
immune responses. To assess the effect of BP-1-102 on the tumor microenvironment,
tumor cell samples from vehicle or BP-1-102-treated mice were collected, stained, and
subjected to flow cytometric analysis. We found that administration of BP-1-102 caused
an increase in the population of CD80
+
/CD86
+
and CD80
+
/MHC II
+
DCs in tumor tissues
(Figure 4-3A). It is well established that both cytotoxic CD8
+
T cells and helper CD4
+
T
cells are involved in antitumor immunity (Fridman et al., 2012; Gu-Trantien et al., 2013).
Therefore, we further analyzed the infiltration of CD8
+
and CD4
+
T cells in tumor tissues
upon BP-1-102 administration. In comparison with vehicle treatment, we observed that
BP-1-102 treatment resulted in markedly increased tumor infiltration of CD8
+
and CD4
+
T cells (Figure 4-3B and 4-3C).

4.4.4 BP-1-102 enhances PD-L1 blockade-mediated tumor regression
Thus far, our results have shown that PD-L1 blockade increased Stat3 phosphorylation
(Figure 4-1B and 4-1C), while, at the same time, Stat3 activation is known to induce
immunosuppression. Therefore, in order to overcome tumor cell resistance and enhance
regression of established tumor, as mediated by PD-L1 blockade, we combined the Stat3

93

Figure 4-3 BP-1-102 modulates the tumor microenvironment.
(A) Representative fluorescence-activated cell sorting plots of CD11c
+
cells to measure
the population of CD80
+
, CD86
+
and CD80
+
MHC II
+
dendritic cells in the tumor tissue
(n = 4). (B) The percentages of CD8
+
tumor-infiltrating T cells in the tumor tissue (n = 4,
error bars indicate SEM; **P < 0.01). (C) The percentages of CD4
+
tumor-infiltrating T
cells in the tumor tissue (n = 4, error bars indicate SEM; **P < 0.01).

inhibitor BP-1-102 and anti-PD-L1 antibody by treating mice bearing melanoma with 6
mg/kg of BP-1-102 every other day and 2.5 mg/kg of anti-PD-L1 antibody every three
days. The treatment regimen is shown in Figure 4-4A. Compared to the control group,
anti-PD-L1 monotherapy inhibited tumor growth, but with limited significance (Figure
4-4B), and it had little effect on survival, which are consistent with our previous
observation (Figure 4-1A). However, we found that BP-1-102 significantly enhanced the

94

Figure 4-4 BP-1-102 enhances the antitumor efficacy of PD-L1 blockade in B16-F10
murine melanoma tumor model.
(A) Schematic representation of the experimental protocol for tumor challenge and drug
treatment. Mice were s.c. challenged with 3 × 10
6
of B16-F10 tumor cells. At day 5,
when the tumors grew to ~30 mm
3
, BP-1-102 and/or anti-PD-L1 antibody treatment was
initiated, and the treatment was continued on the indicated dates. Tumor volume was
measured every other day, and mice were euthanized when the tumor volume in control
groups reached around 1,000 mm
3
. (B) Tumor growth curve of the transplanted B16-F10
cells in C57BL/6 mice treated with BP-1-102 and/or anti-PD-L1 antibody. Data were
presented as mean tumor volume ± standard error of the mean (SEM) at indicated time
points (n = 5; NS, P > 0.05; **P<0.01; ***P < 0.001). (C) Mouse survival was
calculated using the Kaplan-Meier method (n = 5; NS, P > 0.05; **P<0.01; ***P <
0.001). (D) Mouse body weight was measured every other day. Data were presented as
mean body weight ± standard error of the mean (SEM) at indicated time points (n = 5).

95

antitumor activity of PD-L1 blockade and that the combined therapy induced significant
tumor regression compared to anti-PD-L1 monotherapy (Figure 4-4B). Moreover, the
combined therapy enhanced the long-term survival (100%) of mice, which was
statistically significant when compared to that of mice treated with anti-PD-L1 antibody
alone (Figure 4-4C). To assess the systemic toxicity of anti-PD-L1 antibody and BP-1-
102 treatment, the body weight of mice in each group was measured every other day.
Compared to vehicle treatment, we found that either anti-PD-L1 antibody or BP-1-102
alone had little effect on body weight. Notably, the combined therapy also had little effect
on body weight (Figure 4-4D).

4.4.5 Combination therapy modulates the infiltration of immune cells into tumor
tissues
To investigate the underlying mechanism of enhanced antitumor activity of combination
therapy, the subpopulation of tumor-infiltrating lymphocytes (TILs) harvested from mice
treated with anti-PD-L1 antibody and/or BP-1-102 was analyzed by flow cytometry. We
found that monotherapy, either anti-PD-L1 antibody or BP-1-102, increased helper CD4
+

T cell (26.4% versus 16.9% and 21.5% versus 16.9%, respectively) and cytotoxic CD8
+
T
cell (21.6% versus 10.7% and 21.2% versus 10.7%, respectively) and populations within
the tumor. Notably, the combined therapy (25.3% of CD4
+
T cells and 54.1% of CD8
+
T
cells) had an enhanced effect on both T cell populations compared to either treatment
alone (Figure 4-5A and 4-5B). The infiltration of regulatory T cells (Treg) has been
linked to cancer progression (Ha, 2009). By analyzing the Treg population, we observed

96

Figure 4-5 The combined therapy of anti-PD-L1 antibody and BP-1-102 enhanced
tumor-infiltrating T cells and decreased regulatory T cells within tumor.
Single-cell suspension from five different mice were mixed together and then purified by
Percoll density-gradient separation, stained by various markers, and analyzed by flow
cytometry for the composition of various subsets of immune cells. (A) The population of
helper CD4
+
T cells in the tumor tissue within CD45
+
TILs. (B) The population of
cytotoxic CD8
+
T cells in the tumor tissue within CD45
+
TILs. (C) The population of
CD3
+
CD4
+
CD25
+
FoxP3
+
Treg cells in the tumor tissue within CD45
+
TILs. The data
shown were representative of mice that received the indicated therapy.

97

a decrease of Treg population in the tumor tissue treated with either anti-PD-L1 antibody
(41%) or BP-1-102 (25%) compared to that in the control tumor tissue (52.1%).
Interestingly, the combination treatment group had an even lower Treg population
(23.4%) compared to either treatment alone (Figure 4-5C).

4.4.6 Combination therapy improves local immune responses at molecular level
We have shown that BP-1-102 enhanced PD-L1 blockade-mediated tumor
microenvironment modulation. Next, we examined the gene expression of T-cell
activation markers and associated cytolytic cytokines in tumor-bearing mice by real-time
q-PCR. IFN-γ, an important marker for cytotoxic T-cell activation, was significantly
upregulated in tumors after combined therapy, compared to anti-PD-L1 antibody
treatment, BP-1-102 treatment, or untreated tumors (Figure 4-6). Additionally, single
treatment had little effect on the mRNA expression of VEGF, a protein important for
angiogenesis and tumor metastasis. The combination treatment, however, significantly
decreased its expression (Figure 4-6). More importantly, tumors from mice treated with
anti-PD-L1 antibody and BP-1-102 had significantly elevated mRNA levels of TNF-α, a
cytokine that can induce acute and hypoxic death of both cancer and stromal cells, as well
as TRAIL, a stimulator of apoptosis in transformed cells, when compared to those from
control, anti-PD-L1 antibody-, or BP-1-102-treated mice (Figure 4-6).

98

Figure 4-6 The combined therapy altered the tumor immune microenvironment.
The mRNA expression levels of IFN-ɣ, VEGF, TNF-α, and TRAIL from tumor tissues
were analyzed. Five tumors from each group were resected, homogenized and pooled.
Total RNA was extracted, and the mRNA expression levels were determined by real-time
reverse transcription–PCR. Graph depicts relative levels of mRNA after normalizing to
GAPDH mRNA levels (mean ± SEM; *P < 0.05, **P < 0.01).

4.5 Discussion
In this study, we show that PD-L1 blockade increased Stat3 phosphorylation. However,
the introduction of BP-1-102, a Stat3 inhibitor, significantly enhanced PD-L1 blockade-
mediated tumor regression in an established B16-F10 metastatic melanoma model
compared to anti-PD-L1 monotherapy. Specifically, the combined therapy increased the

99

tumor infiltration of cytotoxic CD8
+
and helper CD4
+
T cells and decreased regulatory
FoxP3
+
T cells in the tumor tissues. Finally, gene expression of cytolytic cytokines, such
as IFN-γ, TNF-α, and TRAIL, was significantly upregulated in tumors of mice treated
with BP-1-102 and anti-PD-L1 antibody, when compared to either treatment given alone.
In our tumor model, we observed that B16-F10 cells, which have a high expression of
PD-L1, responded to single PD-L1 blockade poorly. It was initially reported that
blocking PD-L1/PD-1 immune checkpoint for patients with high expression of PD-L1 in
either tumor cells or TILs in primary tumors resulted in poor prognosis compared to PD-
L1-negataive patients (Thompson et al., 2004). More recent studies reporting on the
correlation between overall survival and PD-L1 expression level in metastatic melanoma
still show inconclusive results (Gadiot et al., 2011). Therefore, in an attempt to
understand the underlying mechanism of immune resistance of PD-L1 blockade, we
found that anti-PD-L1 antibody increased the phosphorylation of Stat3 in B16-F10 tumor
cells. Since Stat3 is important for tumor progression and immune evasion, these data
imply that the activation of Stat3 by PD-L1 blockade may, in fact, contribute to acquired
immune resistance, which is consistent with the study conducted by Van Allen et al. who
showed that patients tend to have low responses to immune checkpoint blockade in the
presence of Jak mutation and constitutive downstream signaling activation (Van Allen et
al., 2015). Based on the fact that Stat3 can be phosphorylated and activated by receptor-
associated Jak, mutation of Jak could result in constitutive activation of Stat3 signaling,
which may then lead to immune resistance against immune checkpoint blockade. The
mechanism underlying the regulation of Stat3 signaling by PD-L1 remains unclear.
However, in our study, we found that PD-L1 blockade increased the expression of TNF-α

100

(Figure 4-6) and IL-1β (data not shown). Similarly, previous studies conducted by Erin
West et al. and others show that PD-L1 blockade upregulates TNF-α expression
(Spranger et al., 2014; West et al., 2013). It is well established that TNF-α and IL-1β
initiate Stat3 phosphorylation and activate Jak/Stat3 signaling (Miscia et al., 2002; Mori
et al., 2011), implying that the increased expression of TNF-α and IL-1β induced by PD-
L1 blockade may contribute to the activation of Stat3.
In this study, we aimed to overcome the immune resistance induced by increased Stat3
signaling activity, while, at the same time, taking advantage of PD-L1 blockade-mediated
activation of antitumor immunity. Therefore, we combined an established Stat3 inhibitor,
BP-1-102 (Zhang et al., 2012), with anti-PD-L1 antibody and demonstrated that BP-1-
102 enhanced the antitumor immunity induced by PD-L1 blockade in B16-F10
melanoma. Furthermore, we showed that combined BP-1-102 and anti-PD-L1 antibody
significantly inhibited tumor growth and prolonged survival, even though these
treatments, when administered separately, had limited effects on tumor growth and
survival compared the control group.
To better understand how Stat3 inhibition affected the tumor microenvironment, we
treated mice solely with BP-1-102 intravenously every other day. BP-1-102 injection
alone was shown to enhance the maturation and activation of tumor-infiltrating DCs,
which is consistent with the previous observation that Stat3 deletion increased DC
maturation and activation (Kortylewski et al., 2005; Wang et al., 2004). Furthermore, our
data show that BP-1-102 treatment resulted in a marked increase in the infiltration of
cytotoxic CD8
+
and helper CD4
+
T cells to the local tumor tissues, which is also
consistent with the previous observation that Stat3 deficiency promotes T lymphocyte

101

infiltration to tumor tissues (Kortylewski et al., 2005) and that Stat3-deficient CD8
+
T
cells exhibit faster proliferation (Yu et al., 2013).
In addition to BP-1-102, PD-L1 blockade also increased the infiltration of T cells to the
local tumor tissues. Inhibition of the PD-1/PD-L1 axis by targeting either PD-1 or PD-L1
has been shown to facilitate the accumulation of adoptively transferred T cells to local
tumor tissue (Peng et al., 2012; Pilon-Thomas et al., 2010). Interestingly, compared to
either treatment given alone, it was found that the combined therapy enhanced the
infiltration of cytotoxic CD8
+
T cells and helper CD4
+
T cells to the local tumor tissues.
Additionally, regulatory T cells in the tumor microenvironment were significantly
decreased by the combined therapy. These data indicate that BP-1-102 and PD-L1
blockade act in concert to modulate the tumor microenvironment through regulating
TILs. By analyzing the gene expression of T cell activation markers and associated
cytolytic cytokines in the tumor tissue, we showed that combined BP-1-102 and PD-L1
blockade markedly increased the expression of IFN-γ, TNF-α, and TRAIL, but decreased
the gene expression of VEGF, which is an important marker for angiogenesis. Based on
the biological function of the PD-1 pathway in T cells, the primary role of inhibiting PD-
1/PD-L1 pathway by either PD-1 blockade or PD-L1 blockade is to reinvigorate T cell
function and enhance the antitumor activity of cytotoxic T cells in the tumor
microenvironment. Studies conducted by Tang et al. showed that facilitating T cell
infiltration in the tumor microenvironment increases the responses to PD-L1 blockade,
thereby reducing tumor resistance to checkpoint blockades (Tang et al., 2016). Therefore,
the increased infiltration of cytotoxic CD8
+
and helper CD4
+
T cells to the local tumor

102

microenvironment and the increased T cell activity may, in turn, contribute to enhanced
antitumor immunity and tumor growth inhibition upon combined therapy.

4.6 Conclusion
In summary, this study reveals that activation of Stat3 signaling induced by PD-L1
blockade leads to tumor evasion to immune checkpoint blockade. However, inhibition of
Stat3 by BP-1-102 allows enhanced infiltration of antigen presenting cells and cytotoxic
CD8
+
T cells into local tumor tissue, resulting in stronger antitumor immunity induced by
PD-L1 blockade. Thus, our studies offer evidence to support immunotherapeutic
approaches that combine immune checkpoint blockade with Stat3-targeted, small-
molecule inhibitors to treat melanoma and other cancers.

103

Chapter 5. Conclusion and Future perspectives

104

The adoptive transfer of gene modified T cells is a rapidly evolving modality for cancer
immunotherapy. CAR engineered T cells provide targeted redirection of T cells against
specific TAAs expressed on the tumor cell surface. The efficacy has been demonstrated
in a variety of blood-borne malignancies, prominently, the CD19 CAR T cells in B cell
leukemia (Kochenderfer et al., 2009). Recently, the BCMA CAR T cells has also shown
remarkable success in clinical trials for the treatment of multiple myeloma (Ali et al.,
2016). The clinical success with CD19 CAR T cells in leukemia have boosted the field
and led to the development of a number of different CARs targeting various TAAs for the
treatment of solid tumors, including CD20, HER2, EGFR, CEA, PSMA and so on. Even
though some encouraging clinical data have been reported in certain solid tumors,
including neuroblastoma, and tumors overexpressing mesothelin, HER2, and EGFR
(Newick et al., 2016a), the results of CAR T cell therapy for solid tumors are still much
less inspiring. Therefore, currently, a major focus of CAR T cell therapy is to improve the
efficacy of CAR T cells to be used in cancers beyond leukemia, especially in solid
tumors.
Unlike the liquid tumor environment of blood malignancies, CAR T cells must
successfully traffic to solid tumor sites and infiltrate the stromal elements of solid tumors
in order to elicit TAA-specific cytotoxicity (Zhang et al., 2016). Even after successful
trafficking and infiltration, T cells must overcome the hostile tumor microenvironment
characterized by acidic pH, hypoxia, nutritional depletion, and a number of
immunosuprresive factors, including suppressive cytokines and suppressive immune
cells, for example, Treg, MDSC and TAM. In addition to the hostile tumor
microenvironment, the CAR Ｔcells, themselves, may also be problematic given the T cell

105

intrinsic regulatory mechanisms. With antigen stimulation, T cells will be activated to
proliferate and produce cytokines, thereby launching the immune responses. On the other
hand, during chronic antigen exposure, such as chronic infection or inflammation, in
order to prevent autoimmunity and maintain immunologic tolerance, T cells themselves
will express co-inhibitory molecules on cell surface, such as PD-1 and CTLA-4, to
restrict the extent and strength of the immune response. These inhibitory molecules are
also known as inhibitory immune checkpoints, which normally is very important during
chronic infection and inflammation. However, in the case of cancer, the tumor cells
utilize this mechanism to escape immune surveillance.
So far, a number of approaches have been development to advance the efficacy of CAR T
cell therapy in solid tumors. For example, in order to improve the persistence of CAR T
cells, T cells have been engineered to express two artificial receptors, T1E28z, which is
for TAA recognition, and 4αβ, which provides cytokine-mediated growth stimulation
(Papa et al., 2015). In addition, to protect the CAR T cell from the inhibitory tumor
microenvironment, CAR T cells are further modified to secrete pro-inflammatory
cytokine IL-12, which is also known as armoured CAR T cells (Koneru et al., 2015b).
The efficacy of armoured CAR T cells has been evaluated in early phase I clinical trials
and shown encouraging results (Koneru et al., 2015a). CAR T cells are also being
investigated in clinical trials by combining with ipilumimab, a CTLA-blocking
monoclonal antibody.
In this study, in order to overcome the highly immunosuppressive tumor
microenvironment and improve the potency of CAR T-cell based therapy in the treatment
of solid tumors, a couple of different strategies have been utilized to combine the CAR T

106

cells with immune checkpoint inhibitors (CPIs), aiming to change the
immunosuppressive tumor microenvironment, prevent tumor-induced hypofunctionality
and enhance the antitumor immunity of infused CAR T cells. Moreover, in order to
achieve the best therapeutic efficacy, especially for patients with refractory tumors, other
than combining immunotherapies, we also combined immunotherapy with chemotherapy
and evaluate the antitumor efficacy of combination therapy for the treatment of advanced
melanoma.
First of all, we combined CAR-T cells with anti-PD-1 antibody and evaluated the
antitumor efficacy in a murine lymphoma model. We observed that PD-1 blockade
significantly enhanced CAR T cell expansion and its effector function in vitro. In the
animal study, we demonstrated that the anti-PD-1 antibody significantly enhanced the
antitumor activity of CAR-T cells and prolonged overall survival. Secondly, instead of
combining CAR T cells with systemic injection of anti-PD-1 antibody, we engineered
CAR T cells to secrete and deliver high concentrations of human scFvs against PD-1
(CAR.αPD1). We demonstrated for the first time that PD-1 blockade by continuously
secreted anti-PD-1 prevented T cell exhaustion and significantly enhanced T cell
expansion and effector function both in vitro and in vivo. In the xenograft mouse model,
we found that the secretion of anti-PD-1 enhanced the antitumor activity of CAR-T cells
and prolonged overall survival. Furthermore, for the first time, we showed that in the
treatment of melanoma cells, PD-L1 blockade significantly increased Stat3
phosphorylation, a critical factor shown to drive tumor progression and immune evasion,
including modulation of tumor-mediated immune suppression. In order to overcome the
resistance of melanoma cells to PD-L1 blockade, we combined BP-1-102, a Stat3

107

inhibitor, and anti-PD-L1 antibody, and demonstrated a significant inhibition of tumor
growth, compared to either treatment alone.
Overall, our work supports the immunotherapeutic approaches that combine PD-1
blockade with CAR-T cell therapy. Our findings demonstrates that the blockade of PD-1
immunosuppression, either by systemic injection of anti-PD-1 antibody or by CAR-T
cells’ self-secretion of anti-PD-1 scFv, could significantly enhance the therapeutic
efficacy of CAR-T cell therapy against established lymphoma or solid tumors,
respectively. Taken together, our study provides a distinct strategy for potentially
improving therapeutic outcomes of CAR-T cell therapy in cancer patients with solid
tumors or that do not respond well to first line treatments.
In the current study, we demonstrated that PD-1 blockade by continuously secreted anti-
PD-1 significantly enhanced T cell expansion and effector function, and improved the
therapeutic efficacy of CAR T cell therapy in a xenograft mouse model of human lung
carcinoma. Given the significant role of PD-1 on TILs and PD-1 blockade on modulating
the tumor microenvironment, we think that it would be very interesting to evaluate the
antitumor efficacy of anti-PD-1 scFv secreting CAR-T cells in syngeneic models.
Additionally, instead of artificially overexpressing CD19 in solid tumor cells, targeting
real solid tumor antigens, such as HER2 or mesothelin, should be among the next steps to
further explore the therapeutic efficacy of anti-PD-1 scFv self-secreting CAR-T cells in
the treatment of solid tumors.

108

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Abstract (if available)

Abstract Chimeric antigen receptor (CAR) T-cell based therapy has shown promise as an immunotherapeutic modality for cancers. It has achieved successful responses in patients with certain hematopoietic malignancies. However, the outcome has been less promising in the treatment of lymphoma or solid tumors, partly owing to the immunosuppressive properties and establishment of an immunosuppressive microenvironment. The PD-1/PD-L1 regulatory pathway has demonstrated particularly antagonistic effects on the antitumor response of TILs. As a result, we were trying to investigate the effects of PD-1/PD-L1 blockade on modulating the tumor microenvironment and the function of infused CAR T cells. ❧ In my first study, to overcome the limitation of CAR-T cells in the suppressive tumor microenvironment, we combined CAR-T cells with anti-PD-1 antibody and evaluated the antitumor efficacy in a murine lymphoma model. We observed that PD-1 blockade significantly enhanced CAR T cell expansion and its effector function in vitro. In the animal study, we demonstrated that the anti-PD-1 antibody significantly enhanced the antitumor activity of CAR-T cells and prolonged the overall survival. ❧ Though anti-PD-1 antibody significantly enhanced antitumor immunity of CAR-T cells, antibody treatment has several limitations. For example, it requires multiple and continuous antibody administration to obtain a sustained efficacy. Also, the large size of antibodies prevents them from entering the tumor mass and encountering the infiltrated PD-1-positive T cells. To account for these inefficiencies, multiple high-dose treatments with immunomodulatory drugs or antibodies are required, but this can result in side effects that range from mild diarrhea to autoimmune hepatitis, pneumonitis and colitis. Moreover, it has been shown that the Fc portion of antibodies may cause immune cell depletion by activating cytotoxic signals within macrophages and natural killer cells, which usually express FcαRI and FcγRIIIA/FcγRIIC, respectively. ❧ Therefore, in my second study, we focused our efforts on engineering CAR T cells to secrete and deliver high concentrations of human scFvs against PD-1 (CAR.αPD1-T), aiming to change the immunosuppressive tumor microenvironment, prevent tumor-induced hypofunctionality and enhance the antitumor immunity of infused CAR T cells. We demonstrated for the first time that PD-1 blockade by continuously secreted anti-PD-1 prevented T cell exhaustion and significantly enhanced T cell expansion and effector function both in vitro and in vivo. In the xenograft mouse model, we found that the secretion of anti-PD-1 enhanced the antitumor activity of CAR-T cells and prolonged overall survival. Collectively, our study presents an important and novel strategy that enables CAR-T cells to achieve better antitumor immunity, especially in the treatment of solid tumors. ❧ In order to achieve the best therapeutic efficacy, other than combining immunotherapies, a well-designed combination of chemotherapy and immunotherapy may also be capable of improving the response rates, especially for patients with refractory tumors. For example, clinical trials with anti-PD-1 or anti-PD-L1 monoclonal antibodies alone have shown notable responses in patients with metastatic melanoma. However, the overall response rate is still low. Therefore, seeking combination therapy with enhanced efficacy of tumor eradication mediated by immune checkpoint blockade remains an urgent task. In the present study, we showed that PD-L1 blockade significantly increased Stat3 phosphorylation, a critical factor shown to drive tumor progression and immune evasion, including modulation of tumor-mediated immune suppression. By combining BP-1-102, a Stat3 inhibitor, and anti-PD-L1 antibody, we demonstrated a significant inhibition of tumor growth, compared to either treatment alone. Taken together, the current study suggests that Stat3 inhibition by the small-molecule inhibitor BP-1-102 can markedly enhances the antitumor potency of PD-L1 blockade, improving antitumor immunity through the enhancement of both tumor microenvironment modulation and local immune responses.

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Asset Metadata

Creator Li, Si (author)

Core Title Engineering chimeric antigen receptor (CAR) -modified T cells for enhanced cancer immunotherapy

School School of Pharmacy

Degree Doctor of Philosophy

Degree Program Pharmaceutical Sciences

Publication Date 07/24/2017

Defense Date 05/03/2017

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag adoptive T cell transfer (ACT),cancer immunotherapy,chimeric antigen receptor (CAR),Combination Therapy,immune checkpoints,OAI-PMH Harvest,PD-1

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Okamoto, Curtis (committee chair), Wang, Pin (committee chair), Shen, Wei-Chiang (committee member)

Creator Email linsa99@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c40-410702

Unique identifier UC11214234

Identifier etd-LiSi-5606.pdf (filename),usctheses-c40-410702 (legacy record id)

Legacy Identifier etd-LiSi-5606.pdf

Dmrecord 410702

Document Type Dissertation

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Rights Li, Si

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

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