University of Southern California Dissertations and Theses

Engineered CAR-T cells for treatment of solid cancers

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Engineered CAR-T cells for treatment of solid cancers

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Engineered CAR-T cells for Treatment of Solid Cancers

By

Natnaree Siriwon

A Dissertation Presented to the

FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfullment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMICAL ENGINEERING)

August 2018

Copyright 2018 Natnaree Siriwon

i

Dedication

This thesis is dedicated to my parents, Werapong and Chantana Siriwon, for their
persistent support throughout my entire Ph.D. study.

ii

Acknowledgements

First and foremost, I would like to express my gratitude to my PhD advisor,
Professor Pin Wang. None of this work would have been possible without his
mentorship, patience and support. There were times when we came across obstacles, but
he has mentored me to be an independent thinker and strive to acquire knowledge to
improve. He has taught me that we could always do better than before and keep
improving, which is a lesson I would forever be grateful for. Meanwhile, I would like to
thank my committee members: Professor Stacey Finley and Professor Katherine Shing,
for their insightful comments and encouragement through my PhD journey. I would also
like to thank Professor Noah Malmstadt and Professor Richard Roberts, who offered me
valuable advice regarding my research, critical questions to consider and served on my
qualifying exam committee.
My sincere thanks also goes to all my labmates. All the essential skills for my
projects was demonstrated and trained by Dr. Liang Xiao and Dr. Yarong Liu. I would
like to express deep gratitude to my collaborators, Dr. Yujeong Kim and Dr. Si Li, who
helped made our projects interesting and successful. I would also like to thank Elizabeth
Siegler, Jennifer Rohrs, John Mac, Xianhui Chen, Yun Qu, Dr. Xiaoyang Zhang, Dr.
Kaori Noridomi, Dr. Aaron Nichols, Dr. Lan Huong Lai and Gunce Cinay for their help
and support. I would also like to thank my undergraduate students Diana Kogan,
Thanaporn Liangsupree and Thanida Dangprasert for their assistant in my earlier projects
when I began my PhD studies. I am very fortunate and honored for the teamwork the
Wang lab has provided me.
Last but not least, I would like to express my deepest gratitude to my family and
my dear friend Tanachat Nilanon. Thank you for your love, support and camaraderie.
Your unceasing encouragement has kept me strong.

iii
Table of contents

Dedication i

Acknowledgements ii

List of Figures vi

CHAPTER 1. Introduction 1
1.1. Adoptive cell transfer as immunotherapy for human cancer 2
1.2. Overview of CAR-T cell therapy 4
1.3. CAR-T cell therapy clinical trials 6
1.4. Obstacles of CAR-T cell therapy in solid tumors 8
1.4.1. Tumor recognition and bystander discrimination 8
1.4.2. Trafficking to tumor site 10
1.4.3. Immunosuppressive TME 11
1.5. Overcoming the immunosuppressive TME: adenosine 12
1.6. Immune checkpoint inhibitors 13
1.6.1. CTLA-4 14
1.6.2. PD-1/PD-L1 15
1.7. Summary and thesis work 16

CHAPTER 2. CAR-T cells surface engineered with drug encapsulated
nanoparticles can ameliorate intratumoral T-cell hypofunction 18
2.1. Abstract 19
2.2. Introduction 20
2.3. Materials and methods 23
2.3.1. Mice 23
2.3.2. Cell culture, reagents and antibodies 23
2.3.3. Preparation of T cells for adoptive transfer and viral transduction 24
2.3.4. Plasmid construction 25
2.3.5. Lentiviral and retroviral vector preparation and transduction 25
2.3.6. Detection of receptor expression on T cell surface 26
2.3.7. Synthesis of nanoparticles and drug encapsulation 26
2.3.8. Nanoparticle conjugation with cells and in situ PEGylation 27
2.3.9. In vitro drug encapsulation and release 27
2.3.10. Cytotoxicity assay 27
2.3.11. Transmigration assay 28
2.3.12. In vivo biodistribution assay 28
2.3.13. Quantification of accumulated nanoparticles at tumor sites 29
2.3.14. In vivo xenograft experiments for prevention study 29
2.3.15. In vivo xenograft experiments for rescue study 29
iv
2.3.16. Ex vivo analysis 30
2.3.17. Intratumoral PTX concentration measurements ex vivo 31
2.3.18. Statistics 31
2.4. Results 32
2.4.1. Nanoparticles stably attached to the surface of CAR-T cells 32
2.4.2. CAR-T cells conjugated with nanoparticles maintain T cell
effector functions 34
2.4.3. Conjugation to CAR-T cells increases tumor localization
and systemic circulation of cMLVs 36
2.4.4. Surface-conjugated cMLVs colocalize with CAR-T cells
inside the tumor mass 38
2.4.5. CAR-T cells conjugated with nanoparticles encapsulated with
A2aR antagonist shows improved antitumor responses in vivo 40
2.4.6. CAR-T cells conjugated with A2aR antagonist-encapsulated
nanoparticles are able to rescue hypofunctional tumor residing
T cells in vivo 43
2.5. Discussion 46
2.6. Acknowledgment 51

CHAPTER 3. Enhanced cancer immunotherapy by chimeric antigen receptor
modified T cells engineered to secrete checkpoint inhibitors 52
3.1. Abstract 53
3.2. Introduction 54
3.3. Materials and methods 56
3.3.1. Mice 56
3.3.2. Cell culture and antibodies 56
3.3.3. Plasmid construction 57
3.3.4. Retroviral production 58
3.3.5. T cell transduction and expansion 58
3.3.6. Surface immunostaining and flow cytometry 59
3.3.7. Intracellular cytokine staining 59
3.3.8. Western blot analysis 60
3.3.9. ELISA 60
3.3.10. Competitive blocking assay 61
3.3.11. Specific cell lysis assay 61
3.3.12. Cell proliferation 62
3.3.13. Tumor model and adoptive transfer 63
3.3.14. Statistical analysis 63
3.4. Results 64
3.4.1. Characterization of anti-CD19 CAR-T cells secreting anti-PD-1
antibody 64
3.4.2. Secreting anti-PD-1 antibody enhances the antigen specific
immune response of CAR-T cells 66
3.4.3. Secreting anti-PD-1 alleviates CAR-T cell exhaustion after
v
antigen stimulation 68

3.4.4. Anti-PD-1 engineered CAR-T cells exhibit enhances antitumor
Reactivity 72
3.4.5. Anti-PD-1 engineered CAR T cells can expand more in vivo
than parental CAR-T cells 73
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 75
3.5. Discussion 78
3.6. Acknowledgement 84

REFERENCES 85

vi
List of Figures

Figure 1-1. Schematic representation of the chimeric antigen receptor (CAR) structure. 6

Figure 2-1. Stable conjugation of cross-linked multilamellar liposome (cMLVs). 32

Figure 2-2. Three dimensional (3D) live cell imaging shows conjugation of cMLVs to
the surface of CAR-T cells. 34

Figure 2-3. Conjugation of cMLV does not alter CAR T cell function. 35

Figure 2-4. cMLVs bound to CAR T cells show more efficient infiltration to
antigen-expressing tumors than free cMLVs. 37

Figure 2-5. Surface conjugated cMLVs co-localize with CAR T cells inside the
tumor mass 48 hours post CAR T cell infusion. 39

Figure 2-5. Anti-CD19 CAR T cells conjugated with SCH58261-releasing cMLVs were
prevented from developing hypofunction in SKOV3.CD19 tumors. 41

Figure 2-7. Anti-CD19 CAR T cells conjugated with SCH58261-releasing cMLVs were
able to rescue hypofunctional tumor infiltrated T cells in SKOV3.CD19 tumors. 44

Figure 2-8. In vitro release rates (%) of SCH58261 47

Figure 2-9. Cytotoxicity of cMLVs loaded with SCH58261 against SKOV-3 human
ovarian cancer cells in vitro. 50

Figure 3-1. Construction and characterization of CAR19 and CAR19.αPD1. 66

Figure 3-2. Anti-PD-1 expression enhanced the antigen-specific immune responses of
CAR T cells. 68

Figure 3-3. Secreting anti-PD-1 scFv protected CAR T cells from being exhausted. 71

Figure 3-4. Adoptive transfer of CAR T cells secreting anti-PD-1 scFv enhanced
the growth inhibition of established tumor. 73

Figure 3-5. CAR T cells secreting anti-PD-1 were expanded more efficiently
than parental CAR T cells in vivo. 74

Figure 3-6. CAR T cells secreting anti-PD-1 were more functional than parental
CAR T cells at local tumor site. 77

1

CHAPTER 1: Introduction

2
1.1 Adoptive cell transfer as immunotherapy for human cancer

Adoptive cell therapy (ACT) is a highly personalized method of cancer therapy that
harnesses the anticancer activity of immune cells. ACT is regarded as a “living” treatment due to
the lymphocytes’ ability to proliferate in vivo and sustained antitumor effector functions, which
are considered as major advantages (Rosenberg and Restifo 2015). Cells that have been used for
ACT in clinical trials include tumor-infiltrating lymphocytes (TILs), T cells genetically
engineered with T cell receptors (TCRs) and T cells engineered with chimeric antigen receptors
(CARs). These approaches to ACT have mediated remarkable regression in a variety of cancer
histologies, such as melanoma, cervical cancer, lymphoma, leukemia and neuroblastoma
(Rosenberg and Restifo 2015).
Preclinical studies pioneering the use of ACT for cancer therapy demonstrated the
efficacy of TILs in treatment of animals bearing micrometastases from various types of tumors.
Populations of TILs for ACT are grown from resected tumors, and are generally a mixture of
CD8+ and CD4+ T cells (Rosenberg and Restifo 2015). Studies in murine tumor models showed
that these TILs are capable of recognizing tumor in vitro and could mediate regression of
established lung and liver tumors (Rosenberg, Spiess et al. 1986). However, adoptive transfer of
the TILs alone had little impact. Therefore, additional treatments with cyclophosphamide and IL-
2 were necessary to boost the efficacy of ACT. Studies showed that lymphodepletion with doses
of cyclophosphamide and administration of IL-2 after cell transfer significantly enhanced the
therapeutic potential of infused TILs. Currently, a commonly used lymphodepleting regiment for
humans consists of 60mg/kg of cyclophosphamide given for two days and 25 mg/m
2
fludarabine
administered over 5 days, prior to T cell and IL-2 administration (Dudley, Wunderlich et al.
3
2002). These early studies shed light to the importance of host inhibitory factors and
lymphodepletion using either chemotherapy or radiation prior to treatment could enhance the
efficacy of ACT.
In clinical trials, ACT using TILs was shown to be an effective immunotherapy method
mainly for patients with metastatic melanoma. Resected melanoma specimens are divided into
multiple tumor fragments and individually cultured in IL-2 (Muul, Spiess et al. 1987). The
lymphocytes would expand and lyse the tumor entirely within 2-3 weeks, giving rise to pure
lymphocyte cultures that could be tested for antitumor activity. These individual cultures are
expanded by co-culturing with excess amounts of irradiated feeder cells (Dudley, Wunderlich et
al. 2002). For ACT, patients would receive a lymphodepleting preparative regiment as stated
previously, prior to receiving cell infusion. This method of TIL isolation and culture could be
performed for multiple cancer types. However, only TILs from melanoma specimens
consistently showed antitumor activity. Therefore, genetically engineering lymphocytes to
express specific antitumor receptors has become an ardently pursued method in order to apply
ACT as a method of immunotherapy for a broader range of cancers (Rosenberg and Restifo
2015). The specificity of T cells could be redirected through engineered expression of either
alpha-beta TCRs or CARs.
The first successful clinical trial of genetically engineered lymphocytes was shown in
2006, for treatment of patients with metastatic melanoma. Autologous lymphocytes were
transduced with retrovirus that encodes weakly avid human TCR recognizing the MART-1
melanoma-melanocyte differentiation antigen. This study showed 45% objective response.
Remarkably, two patients were reported to show objective partial regressions of metastatic
4
melanoma and sustained levels of engineered lymphocytes in circulation at 1 year after infusion
(Morgan, Dudley et al. 2006).

1.2 Overview of CAR-T cell Therapy

The development of CARs was pioneered in the late 1980s by Gross and colleagues
(Gross, Waks et al. 1989). CARs specifically target tumor-associated antigens or cancer stromal
antigens with high binding affinity. Activation of a CAR engineered T cell is independent of the
major histocompatibility complex (MHC) and is achieved primarily through the single chain
variable fragment (scFv) that is fused with intracellular signaling domains (Sadelain, Brentjens et
al. 2009). The advantages of CAR over TCR is the broad range of antigens that CAR T cells
could be engineered to target, this includes self antigens that is often eliminated in the
physiological T cell repertoire. Thus, CAR T cells could be engineered to target native host
tissue, T cell subsets, immune cell progenitors, natural killer (NK) cells and etc. Moreover, since
CARs do not require antigen processing by the MHC or human leukocyte antigen (HLA), which
is a gene complex encoding the MHC proteins in humans, it could recognize surface antigens
without regard to the patients’ diverse HLA (Sadelain, Brentjens et al. 2013).
Designing scFv moieties to enhance and optimize CAR T cell activation and cytotoxicity
is extensively studied. ScFv moieties are derived from three general sources: antibodies or Fab’s
selected from libraries or natural ligands that engage their cognate receptor. (Sadelain, Brentjens
et al. 2013) These moieties target tumor-associated antigens (TAA) that are optimally selected
for purposes of CAR T cell targeting. Evidently, an important characteristic of suitable TAAs is
to be highly expressed on tumors, and low to none in normal tissues. A good example is CD19
5
that is expressed on blood-borne tumor surfaces and only on dispensable B cells. CD19 CAR T
cells, thus, showed remarkable clinical success that is yet to be repeated with other CARs.
Suitable TAAs for most solid tumors, especially, are harder to identify due to its general
expression in normal tissues. Currently, approximately 30 solid tumor antigens are being studied
for CAR T cell therapy. These include neoantigens (i.e. mutated sequence occurring on
oncogenic cells), developmental antigens, tumor selective antigens and endogenous tumor
specific antigen. (Gubin, Artyomov et al. 2015, Newick, Moon et al. 2016) EGFR variant 3
(EGFRvIII) is an example of neoantigens that is expressed exclusively on malignant tumor cells,
such as glioblastomas. Unfortunately, most neoantigens are tumor specific and highly
individualized, unlike the mutation of the EGF receptor. Consequently, EGFRvIII CAR became
an attractive model for study of solid cancer treatment with adoptive CAR T therapy. (Johnson,
Scholler et al. 2015, Newick, Moon et al. 2016)
In addition to the scFv targeting moiety, the CAR consists of intracellular signaling
domains that are essential for T cell activation, survival and proliferation in vivo. These
intracellular domains activate other signaling proteins to relay the signal downstream to several
pathways that govern effector T cell functions (Sadelain, Brentjens et al. 2013). The
development of CAR intracellular signaling domains progressed in “generations”. The
intracellular signaling domain of first-generation CARs consists merely of CD3ζ, which is
insufficient to elicit a robust cytokine response and further support T cell clonal expansion
(Gong, Latouche et al. 1999). Second generation CARs incorporates one additional
costimulatory domain, usually either CD28 or 41BB, to CD3ζ, and third-generation CARs
consists of two costimulatory domains and CD3ζ as shown in Figure 1-1 (Long, Haso et al.
2015, Maude, Teachey et al. 2015). Second generation CARs of either CD28 or 4-1BB
6
costimulatory domains were tested in patients and have yielded remarkable outcomes; however,
it has been shown less effective in the case of solid tumors where the tumor microenvironment
has rendered more powerful in suppressing T cell function. (Sadelain, Brentjens et al. 2013)

Figure 1-1. Schematic representation of the chimeric antigen receptor (CAR) structure. CARs target surface
antigens in a major histocompatibility class-independent manner and are comprised of an extracellular portion
typically derived from an antibody and intracellular signaling modules derived from T-cell signaling protiens.

1.3 CAR T cell therapy clinical trials

CAR T cell therapy has been tested in several clinical trials for patients with
hematological malignancies and multiple studies was also performed on patients with solid
tumors. In the case of hematological malignancies, such as acute lymphoblastic leukemia (ALL),
chronic lymphoblastic leukemia (CLL) and multiple myeloma, CD19 has been a successful
target for CAR T cell therapy.(Kalos, Levine et al. 2011, Tasian and Gardner 2015) Other targets
7
such as CD20 in lymphoma patients(Zhang, Wang et al. 2016) and Lewis Y antigen in acute
myeloid leukemia were also studied.(Ritchie, Neeson et al. 2013) However, CD19 remains the
most investigated target to date due to its uniform expression by most B cell malignancies, and
not on normal tissues other than those derived from the B cell lineage. CAR-T cell therapy, on
the other hand, is still in its early stages of development for treatment of solid cancers. To date,
only a few clinical trials have been completed on solid cancers: glioblastoma (against IL13Rα2
antigen), HER2-positive sarcoma (against HER2 antigen), neuroblastoma (against GD2 antigen),
liver metastases (against carcinoembryonic (CEA) antigen) and pleural mesothelioma (against
MSLN antigen) (Mirzaei, Rodriguez et al. 2017). Modifications to the CAR design, T cell
culture and treatment strategies had to be made to yield success in treatment of solid cancers.
The first clinical trial utilizing CAR-T cells was conducted in 2006 to assess the safety
and efficacy of adoptive immunotherapy for treatment of metastatic ovarian cancer (Kershaw,
Westwood et al. 2006). Unlike melanoma reactive TILs, endogenous ovarian reactive T cell are
difficult to isolate. Therefore, to facilitate production of therapeutic cells, T cells were
genetically engineered to recognize the ovarian cancer associated antigen α-folate receptor
(Kershaw, Westwood et al. 2006). The surface chimeric receptor linking single-chain (scFv) was
linked to the transmembrane and cytoplasmic domains of the Fc receptor. None of the patients
from this clinical trial showed evidence of antitumor response. Some patients from cohort 1 of
the study showed grade 3 and 4 toxicities, likely due to the use of high-dose IL-2. Moreover, the
study reported that there was poor trafficking of T cells to the tumor, though the reasons were
unclear. The study suggested several factors that could have contributed to poor tumor
localization. First, the T cells were retained in the lung, liver and spleen due to the intravenous
route of delivery. Thus, less T cells trafficked to the tumor. Second, the transferred T cells
8
showed low persistence in patients. This may be caused by long-term culture conditions that
hinder in vivo adaptation and “exhaust” T cells (Kershaw, Westwood et al. 2006).
The first objective response observed for CAR-T cell therapy against solid cancers was
reported in 2011, for treatment of neuroblastoma (Louis, Savoldo et al. 2011). First generation
CARs targeting GD2 was expressed either on Epstein Barr-virus (EBV) specific cytotoxic T
lymphocytes (CAR-CTLs) or activated T cells (CAR-ATCs) (Pule, Savoldo et al. 2008). The
advantages of utilizing CAR-CTLs is its ability to be costimulated by EBV antigens expressed
on antigen presenting cells (APCs), and consequently survive in circulation at a higher level than
ATCs. In this study, three out of 11 patients showed complete remission (27%). For several
weeks after treatment, CAR-CTLs had higher number in circulation compared to CAR-ATC.
However, there was no discernable difference after six weeks. The cells were found to persist at
low levels in a long-term study: 96 weeks for CAR-CTLs and 192 weeks for CAR-ATCs (Louis,
Savoldo et al. 2011). The conclusion of this study suggests that long term low –level presence of
CAR expressing T cells is associated with clinical benefits, as shown in the three patients that
had complete remission and long-term CAR-T cell persistence. This study did not report
complications and toxicities, which has been one of the major challenges facing CAR-T cell
therapy.

1.4 Obstacles of CAR-T cell therapy in solid tumors
For solid tumors, positive results have been acquired, although to a lesser degree
compared to blood borne cancers, from a clinical trial in patients with metastatic or recurrent
HER2 positive sarcoma. HER2.CD28ζ CAR T cells were able to stabilize the disease for 12 to
14 months (Ahmed, Brawley et al. 2015). Another clinical trial was performed on neuroblastoma
9
patients using CAR T cells to target GD2 ganglioside (Louis, Savoldo et al. 2011). Three out of
19 patients showed complete remission in this clinical trial. The limitation of adoptive CAR T
cell therapy in solid tumors is attributed to multiple factors, such as the absence of unique tumor
associated antigens, inefficient homing of T cells and the ability to persist in an
immunosuppressive tumor microenvironment (TME) (Dai, Wang et al. 2016).

1.4.1 Tumor recognition and bystander discrimination
There are two forms of bystander cross-reaction that are possible: OFF-target and ON-
target OFF-tissue cross-reaction. OFF-target cross-reaction occurs when the engineered receptor
cross-reacts with an untargeted stereochemically related antigen that is present on an essential
tissue. ON–target, OFF–tumor cross-reaction occurs with a targeted tumor antigen expressed on
other tissues at a level that is detectable by CAR-T cells (Lim and June 2017). The absence of
unique TAA raises the risk of ON–target, OFF–tumor toxicity, which prompts an immune-
mediated destruction of normal tissues. One particular case was reported in a patient who
received a third-generation CAR T cell therapy targeting HER2, which is expressed at a low
level in several normal tissues, such as the heart and pulmonary vasculature (Morgan, Yang et al.
2010). Another case of ON–target OFF–tumor toxicity was reported in a clinical trial that used T
cells specific for carbonic anhydrase IX, which incurred cholestasis in a patient (Lamers, Sleijfer
et al. 2006). However, CAR against other solid tumor targets has yet been reported to result in
ON–target, OFF–tumor toxicity. These include CD171-, GD2-, CEA- and IL13Ra2 directed
CAR T cell therapies (Dai, Wang et al. 2016). In order to avoid adverse side effects, it is critical
to select T cell targets that are most specific to tumor tissues in addition to improving the
receptor affinity and specificity of the CAR. Other approaches to reduce the risk of side affects
10
are to select an appropriate cell dosage and condition regiments prior to T cell infusion (Dai,
Wang et al. 2016).

1.4.2 Trafficking to tumor site
The second problem concerning effective CAR T cell therapy against solid cancers is
efficient trafficking of effector T cells to tumor sites (Newick, Moon et al. 2016). Infused T cells
travel throughout the body and home to sites where a target antigen is expressed. T cell
trafficking is largely facilitated by chemokines. However, a problem of a chemokine mismatch
between the tumor and CAR T cells arises (Yong, Dardalhom et al. 2017). T cells are frequently
found to lack appropriate chemokine receptors that directs it towards the tumor, and thus, are
hindered from migrating to its target. Engineering CAR T cells to express chemokine receptors
specific to tumors could thereby help increase their localization to tumor sites. Di Stasi et al
(2009), for example, performed a study where CD30-specific CAR T cells were engineered to
express CCR4, a receptor for CC chemokine ligand 17 (CCL17) and macrophage derived
chemokine (MDC)/CCL22 that is secreted from Hodgkin lymphoma tumors. Generally, CCR4 is
expressed on regulatory T cells (Tregs) and helper T cells (Th2), which generates an
immunosuppressive environment once they home to the tumor (Di Stasi, De Angelis et al. 2009)
Once the CAR T cells have homed to the tumor, they have to survive and proliferate in
order to effectively eradicate the tumor mass and sustain protection from tumor recurrence. This
gives rise to the third problem that CAR T cells become exhausted and progressively reduce in
number (Newick, Moon et al. 2016). In attempt to improve the persistence of CAR T cells,
subsets of CAR T cells that may proliferate and survive for longer periods of time, such as naïve
or central memory T cells are selected for adoptive transfer. This was proposed in Berger et al.
11
(2007) that demonstrated CD8+ T cells derived from the central memory are districted from
those derived from effector memory T cells and possess an intrinsic property that enables them
to survive after adoptive transfer and revert back to the memory cell pool (Berger, Jensen et al.
2008).

1.4.3 Immunosuppressive tumor microenvironment (TME)
The immunosuppressive TME is one of the ultimate obstacles for immunosurveillance
and especially for CAR T cell therapy, due to multiple cellular and molecular components that
reduce the antitumor immune functions. The combined effect of immunosuppressive factors can
severely impair CAR-T cell functions, although the effector functions of these cells could be
restored upon removal from the TME (Riese, Wang et al. 2013). These findings suggest that
appropriate interference of tumor immunosuppression could prevent CAR-T cells from being
inhibited in the TME and the reduction of T cells effector functions could be reversed with
appropriate treatments.
The TME consists of tumor cells and also multiple regulatory cells –regulatory T cells,
NKT cells, and subsets of immature and mature dendritic cells–that could inhibit CAR-T cell
effector functions either through inhibitory cytokines or activation of a negative inhibitory
pathway. Regulatory T cells, NKT cells and tumor cells secrete tumor-promoting cytokines such
as TGF-β and IL-10. Tumor cells up-regulate checkpoint inhibitory proteins such as PD-L1,
which suppresses T cells through the PD-1 receptor that is expressed on the surface of T cells.
These regulatory cells in combination with tumors themselves foster a tolerant
microenvironment that hinders any successful attempt of an effective immune response to
eradicate the tumor mass (Newick, Moon et al. 2016).
12
In addition to regulatory cells, the tumor microenvironment is characterized by hypoxia
and immunosuppressive soluble factors that inhibit T cell function such as adenosine (Stagg and
Smyth 2010). Adenosine is converted from extracellular ATP through the enzymatic activity of
ectonucleases CD39 and CD73. A large number of tumors were reported to have highly
upregulated CD39 and CD73 expression, these include melanoma, leukemia, pancreatic, liver,
gastric, colon and breast cancers. The ectonucleases are also expressed on a variety of tissues,
such as the heart, placenta, lung, liver, colon, brain and kidney. Notably, CD39 and CD73
ectonucleases are also expressed by immune cells, and particularly highly expressed on CD4+
Foxp3+ regulatory T cells (Ohta and Sitkovsky 2014). CD39 and CD73 expression are
upregulated under hypoxic conditions, the presence of IGF-B1, IL-6, type-1 IFNs and Wnt
signaling pathway agonists. Adenosine allows the tumor to escape immunosurveillance through
multiple mechanisms: (i) inhibition of activation, expansion and cytokine production by T-cells,
(ii) the abrogation of T-cell and NK-cell cytotoxicity, (iii) the induction of regulatory T cells, (iv)
the prevention of dendritic cell maturation and activation, for instance (Stagg and Smyth 2010,
Ohta and Sitkovsky 2014).

1.5 Overcoming the immunosuppressive TME: adenosine
Interfering the adenosine pathway was demonstrated to increase T cell and NK cell
cytotoxicity against human ovarian cancer cells. Adenosine in the TME suppresses cytotoxic
immunity by triggering the A2A receptor expressed on the cell surface of T cells and NK cells.
Multiple strategies were tested to inhibit immunosuppression by adenosine in the TME, these
include using specific anti-CD39 and anti-CD73 antibodies, small molecule antagonist to the
A2A receptor and siRNA knockdown of the A2A receptor. These strategies showed positive
13
outcome, which supports the hypothesis that the A2A receptor pathway is a critical
immunosuppressive mechanism (Ohta, Gorelik et al. 2006).
Ohta et al. (2006) hypothesized that the A2a receptor pathway protects the cancerous
tissue from incoming immune cells and the use of A2a receptor antagonists or siRNA
knockdown of the receptor would prevent the inhibition of tumor infiltrating immune cells (Ohta,
Gorelik et al. 2006). Binding of adenosine to the A2a receptor up regulates intracellular cAMP,
which suppresses TCR activation and inhibits antitumor effects of T cells. Ohta et al.
demonstrated that this inhibition could be “rescued” through genetically targeting the A2a
receptor or abrogating adenosine activation by treatment with an A2a receptor antagonist. A
study in C57BL/6 mice model showed that A2AR gene deficiency leads to complete tumor
rejection, even in mice with large established tumors. In wild type mouse models, the A2a
receptor antagonists –ZM241,385 and caffeine- were also found to enhance the antitumor effects
of CD8
+
T cells, which express functional A2 adenosine receptors. Notably, tumor rejection
ensuing both strategies of blocking A2a receptor signaling is the result of CD8
+
T cell antitumor
functions. These findings provide evidence of the critical immune regulatory role in which the
A2a receptor pathway suppresses effector functions of antigen-specific CD8
+
T cells in vivo and,
particularly, in the TME (Ohta, Gorelik et al. 2006).

1.6 Immune Checkpoint Inhibitors
To deliver potent antitumor immunity, T cells must evade negative regulatory signals
present in the TME that induces tolerance programs such as anergy or exhaustion. It is of note
that these immune checkpoints regulate a diversity of functions, and each inhibitor has different
mechanisms of action. Recent development of checkpoint inhibitors, such as ipilimumab
14
targeting CTLA-4 or a human IgG4 anti-PD-L1 mAB, provided new opportunities to enhance
antitumor immunity with the potential to produce durable clinical response (John, Devaud et al.
2013). These two inhibitors, for example, regulate the immune response at different levels and
by different mechanisms (Pardoll 2012). The positive clinical response to either of these
inhibitors suggests that antitumor immunity could be augmented at multiple levels and that
combinatorial treatments could be designed.
1.6.1 CTLA-4
The first clinically targeted immune checkpoint inhibitor is CTLA-4, which is expressed
exclusively on T cells (Pardoll 2012). CTLA-4 counteracts the activity of the T cell co-
stimulatory receptor, CD28, by engagement with ligands: CD80 (also known as B7.1) and CD86
(also known as B7.2) (Freeman, Gribben et al. 1993). The exact mechanisms of CTLA-4 are
under considerable debate, though it has been proposed that its expression on the T cell surface
dampens the activation of T cells by outcompeting CD28 in binding the shared target ligands,
CD80 and CD86 (Schwartz 1992). The specific signaling pathway that CTLA-4 engages is also
under investigation, although a number of studies suggested the activation of protein
phosphatases, SHP2 and PP2A, are crucial for counteracting kinase signals that are induced by
TCR and CD28 (Schwartz 1992). Both CD4 and CD8 T cells express CTLA4, although the
major physiological role of CTLA4 appears to be through distinct effects on the two major
subsets of CD4
+
T cells: downregulation of helper T cell activity and enhancement of regulatory
T cell immunosuppressive activity (Pardoll 2012).
The general strategy of CTLA-4 blockade was initially controversial because there is no
tumor specificity to the expression of the CTLA4 ligands –except for some myeloid and
lymphoid tumors. CD80 and CD86 are majorly expressed on antigen presenting cells (APCs)
15
such as dendritic cells, and full blockade of CTLA-4 signaling may lead to immune toxicities due
to hyperactive immunity (Pardoll 2012). However, preclinical models in mice demonstrated that
a therapeutic window could be achieved with partial CTLA-4 blockade by antibodies. Two
CTLA-4 antibodies, Ipilimumab and Trmelimumab, have been tested in clinical trials (Pardoll
2012). Ipilimumab is FDA approved for melanoma, while phase II and phase III are ongoing for
multiple cancers. A notable feature of clinical response to oncogene-targeted small molecule
drugs is their kinetics (Pardoll 2012). Unlike responses to chemotherapies and tyrosine kinase
inhibitors, responses to immune checkpoint blockers is slower and, in some patients, could delay
up to six months after treatment. There are instances where metastatic lesions increase in size
before regressing, which seems to occur due to increased immune cell infiltration into the lesions
(Phan, Yang et al. 2003).
1.6.2 PD-1/PD-L1
The PD-1 signaling pathway has emerged as another promising immune checkpoint to
modulate for inducing antitumor responses. Unlike CTLA-4, the PD-1 signaling pathway
suppresses T cell activity in peripheral tissues at the time of an inflammatory response to
infection and to limit autoimmunity. Its target ligand, PD-L1 and PD-L2, are expressed on APCs,
peripheral tissues and also tumor cells (Zou, Wolchok et al. 2016). PD-1 expression is up
regulated after T cells have become activated and inhibits kinases that are involved in T cell
activation through the phosphatase SHP2, when engaged with its ligand. Consequently, PD-1
mainly regulates T cell activity, while CTLA-4 predominately regulates T cell activation.
While CTLA-4 is exclusively expressed on T cells, PD-1 is expressed on a variety of
immune cells in addition to T lymphocytes, such as B cells and NK cells(Zou, Wolchok et al.
2016). Therefore, treatments with PD-1 blockade may enhance antibody production from PD1
+

16
B cells and NK cell activity in tumors and tissues, in addition to effector T cell functions (Pardoll
2012). Chronic antigen exposure in the TME could result in persistent PD-1 expression, which
induces a T cell anergy. However, this state is reversible through interference with the PD-1
signaling pathway by PD-1 antibodies (John, Devaud et al. 2013).
Antibodies targeting PD-1 and PD-L1 have been tested in clinical trials for treatment of
several cancer histologies: colon, renal and lung cancers, melanoma and follicular lymphoma
(Brahmer, Drake et al. 2010, Westin, Chu et al. 2014, Rizvi, Mazières et al. 2015). In the first
Phase I clinical trial with fully human PD-1 antibody, the goal was to evaluate the safety and
tolerability of anti-PD-1 blockade, and to preliminary assess antitumor activity. Patients with
treatment-refractory solid tumors –advanced metastatic melanoma, colorectal cancer, castrate-
resistant prostate cancer, non-small cell lung cancer, or renal cell carcinoma- received an
intravenous infusion of anti-PD-1. The study showed that blocking the PD-1 immune checkpoint
is well tolerated and has antitumor activities, particularly in tumor tissues with PD-L1
expression. This favorable outcome propelled further clinical trials to test the efficacy of PD1
antibodies, in the USA and internationally. Currently, multiple clinical studies are ongoing in
Phase III clinical trials for several PD-1 antibodies (Liu and Wu 2017). In 2017, FDA approved
the anti-PD-1 antibody, nivolumab (OPDIVO, Bristol-Myers Squibb Company) for treatment of
patients with melanoma (Weber, Mandala et al. 2017).

1.7 Summary of thesis work

This thesis work is divided into two major studies that aim to overcome two different
immunosuppressive mechanisms in the TME: adenosine up-regulation and PD-1/PD-L1 immune
inhibition. In chapter 2, we surface-engineered CAR-T cells to deliver an A2a receptor small
17
molecule inhibitor in the TME. The small molecule inhibitor is encapsulated in cross-linked
multilamellar liposomes, which are covalently conjugated to the surface of CAR-T cells. We
hypothesized that this co-delivery strategy would be able to enhance CAR-T cell functions in the
TME and, consequently, augment CAR-T cell antitumor activities. We demonstrated in this
study that our drug delivery strategy was able to prevent CAR-T cells from becoming
hypofunctional in the TME and improved CAR-T cell antitumor response. Moreover, we showed
that the antitumor function of tumor infiltrated CAR-T cells could be rescued upon interference
of the adenosine pathway using A2a receptor small molecule inhibitors. This study signifies the
effects adenosine has on tumor infiltrating immune cells and demonstrated a strategy in which it
could be overcome, in order to improve the efficacy of CAR-T cell therapy.
The second obstacle we aim to overcome is the PD-1/PD-L1 immune regulatory
pathway. In chapter 3, we engineered CAR-T cells to secrete the PD-1/PD-L1 checkpoint
inhibitor and evaluated the antitumor efficacy in vivo. This strategy was proved to significantly
enhance CAR-T cell effector functions in vivo and was able to completely eradicate established
tumors. We evaluated the effector functions in an ex vivo assay and demonstrated that the
intrinsically secreted PD-1/PD-L1 checkpoint inhibitor was able to prevent CAR-T cell
exhaustion and maintained IFNγ secretion upon restimulation.

18

CHAPTER 2:

CAR-T Cells Surface-Engineered With Drug-Encapsulated Nanoparticles Can Ameliorate
Intratumoral T-Cell Hypofunction

Portions of this chapter are adapted from: Natnaree Siriwon*, Yujeong Kim*, Elizabeth Siegler,
Xianhui Chen, Jennifer Rohrs and Pin Wang. Cancer Immunology Research (Under review)

*These authors contributed equally to this work

19

2.1. ABSTRACT

The chimeric antigen receptor T (CAR-T) cell therapy has become a promising cancer
immunotherapeutic method, particularly in treating B cell malignancies; however, this therapy is
still in the early stages of development for the treatment of solid tumors. One limiting factor of
CAR-T cell therapy is the suppressive tumor microenvironment, which inactivates the function
of tumor infiltrating lymphocyte (TIL) through the production of immunosuppressive molecules
such as adenosine. Adenosine inhibits the function of CD4 and CD8 T cells by binding to and
activating the A2a adenosine receptor (A2aR) expressed on their surface. This suppression
pathway can be blocked using the A2aR-specific small molecule antagonist SCH-58261 (SCH),
but its applications have been limited owing to difficulties delivering this drug to immune cells
within the tumor microenvironment (TME). To overcome this limitation, we have used CAR-
engineered T cells as active chaperones to deliver SCH-loaded cross-linked multilamellar
liposomal vesicles (cMLVs) to tumor-infiltrating T cells deep within the immune suppressive
TME. Through in vitro and in vivo studies, we have demonstrated that this system can be used to
deliver SCH to the TME. This treatment can prevent or rescue the emergence of hypofunctional
CAR-T cells within the TME.

20
2.2. INTRODUCTION

Chimeric antigen receptor (CAR)-engineered T (CAR-T) cell therapy has demonstrated
success in treating hematological cancers, such as leukemia and B cell lymphoma, in preclinical
and clinical trials (Brentjens, Davila et al. 2013, Davila, Riviere et al. 2014, Lee, Kochenderfer et
al. 2015). This success has not been translated to the treatment of solid tumors (Morgan, Dudley
et al. 2006, Morgan, Yang et al. 2010, Fesnak, June et al. 2016). Unlike hematological cancers,
which circulate throughout the body in the blood stream, solid tumors have their own complex
tumor microenvironment (TME), which provides a unique barrier to immunotherapy (Quail and
Joyce 2013, Newick, Moon et al. 2016). To be effective, immune cells must efficiently infiltrate
the solid tumor mass and have extended persistence of in vivo expanded cells (Dudley,
Wunderlich et al. 2002, Louis, Savoldo et al. 2011). The TME contains a variety of pro-
tumorigenic factors that work to both prevent cancer-killing immune cells from entering the
tumor area and dampen the activation of tumor infiltrating lymphocytes (TILs) (Mellman,
Coukos et al. 2011, Quail and Joyce 2013, Jiang, Li et al. 2015). Many of these immune
suppressive mechanisms can also negatively impact adoptively transferred CAR-engineered T
cells.
One of the underlying mechanisms responsible for the progressive loss of TIL function
in the TME is an inhibitory pathway involving the A2a adenosine receptor (A2aR) expressed on
the surface of activated T cells (Ohta, Gorelik et al. 2006, Cekic and Linden 2014, Beavis,
Henderson et al. 2017). The A2aR pathway is triggered by abnormally high concentrations of the
extracellular immunosuppressive molecule adenosine, which has been reported to suppress T cell
proliferation and IFN-γ secretion (Lappas, Rieger et al. 2005, Jin, Fan et al. 2010). In the TME,
extracellular adenosine triphosphate (ATP) is released in response to tissue damage and cellular
21
stress. ATP in the extracellular environment is converted into adenosine by ecto-nucleases CD39
and CD73, which are upregulated in the hypoxic TME (Stagg and Smyth 2010). Overexpression
of CD73 has been observed in multiple aggressive cancers, conferring resistance to antitumor
agents (Bono, Fernández et al. 2015). Binding of adenosine to A2aR leads to increased
intracellular cyclic AMP (cAMP) production in the TILs. Elevation of intracellular cAMP
induces activation of protein kinase A (PKA) and phosphorylation of the cAMP response
element binding protein (CREB), which, in turn, abrogates T cell receptor (TCR) signaling and
IFN-γ production by reducing the activity of the Akt pathway and inhibiting NF-kB-mediated
immune activation (Cekic and Linden 2014, Bono, Fernández et al. 2015).
Studies have demonstrated that blocking A2aR signaling through either pharmacological
inhibition or genetic deletion can significantly improve antitumor immunity by enhancing the
cytotoxic function of T and natural killer (NK) cells (Ohta, Gorelik et al. 2006, Jin, Fan et al.
2010, Beavis, Divisekera et al. 2013, Beavis, Henderson et al. 2017). SCH-58261 (SCH) is one
of the most selective and potent antagonists of A2aR (Fredholm, IJzerman et al. 2001). Despite
its therapeutic potential, the clinical application of SCH has been hindered by the drug’s poor
solubility and suboptimal in vivo pharmacokinetic profile (Cacciari, Pastorin et al. 2003). Small-
molecule drugs like SCH, which act directly on CAR-T cells, need to be maintained at high
concentrations near their site of action in order to be effective. Thus, a carrier capable of
regulating drug circulation time in vivo and efficiently delivering the drug to CAR-T cells in
tumors, while minimizing delivery to other tissue/organ sites, is desired.
Recently, advances in drug delivery nanotechnology have enhanced the therapeutic
efficacy of several anticancer drugs. Compared to free drugs, drug-loaded nanoparticles can
improve targeted delivery by prolonging blood circulation time, controlling and sustaining drug
22
release profiles, altering tissue distribution to reduce systemic toxicities, and increasing drug
concentration in the tumor through the enhanced permeability and retention (EPR) effect (Bae
and Park 2011). The EPR effect is highly dependent on adequate vascularization of tumors.
Vascularization, however, may be completely lacking in some tumors that exhibit poor blood
supply and hypoxia (Nishida, Yano et al. 2006). Additionally, high interstitial fluid pressures
within the tumor can act to transport therapeutics back into the bloodstream. Most administered
nanoparticles (up to 95%) are reported to accumulate in organs other than tumor, including, for
example, liver, spleen and lungs (Bae and Park 2011). Hence, efficient delivery and distribution
of nanoparticles within the tumor mass remains challenging.
Numerous studies have shown that the addition of targeting moieties on nanoparticles
can significantly improve their tumor specificity and accumulation, but these targeting strategies
still rely on passive distribution of the nanoparticles through the bloodstream (Byrne, Betancourt
et al. 2008). Recently, more advanced strategies have been explored, such as the use of immune
cells as nanoparticle “chaperones” for targeted delivery and active transport into tumors
(Stephan, Moon et al. 2010, Stephan, Stephan et al. 2012, Huang, Abraham et al. 2015). Studies
have shown that these immune delivery cells can provide effective treatment of tumors by
facilitating drug delivery through active cellular migration and extravasation in response to
chemoattractant gradients around the tumor and inflammatory sites (Mrass, Takano et al. 2006,
Stephan, Moon et al. 2010, Huang, Abraham et al. 2015). We believe that this delivery approach
can be applied to alter interactions between the TME and endogenous or infused immune cells in
vivo.
Therefore, we herein report an approach that enhances the efficacy of CAR-T therapy by
chemical conjugation of adjuvant drug-loaded maleimide-functionalized cross-linked
23
multilamellar liposomes (cMLVs) to the surface of CAR-T cells ex vivo prior to systemic
administration (Fig. 2-1A). We demonstrated that cMLV nanoparticles could be covalently
attached to CAR-T cells without affecting the cells’ viability and effector functions. We further
revealed that the therapeutic potential of CAR-T cells could be improved upon surface
engineering with SCH-encapsulated cMLVs.

2.3. MATERIALS AND METHODS

2.3.1. Mice
Female NOD.Cg-Prkdc
scid
IL2Rγ
tm1Wj1
/SZ (NSG) mice (6 to 10 weeks old) were purchased
from the Jackson Laboratory (Bar Harbor, ME). 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 USC.
2.3.2 Cell culture, reagents and antibodies
The human ovarian cancer cell line SKOV3 (ATCC
®
HTB-77™) and the human chronic
myelogenous leukemia cell line K562 (ATCC
®
CCL-243™) were maintained in RPMI-1640
with 10% FBS, 2 mM L-glutamine, 10 mM HEPES, 100 U/ml penicillin, 100µg/ml streptomycin
and 100µg/ml Normocin™ (InvivoGen, San Diego, CA). K562.CD19 and SKOV3.CD19 cells
were generated by transducing parental K562 and SKOV3 cells with lentiviral vectors encoding
the cDNA of human CD19. All cells were routinely tested for potential mycoplasma
contamination using the MycoSensor qPCR assay kit (Agilent Technologies).
SCH-58261 was purchased from Sigma-Aldrich (St. Louis, MO). All lipids were purchased
from NOF Corporation (Japan): 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-
dioleoyl-sn-glycero-3-phospho-(10-rac-glycerol) (DOPG), and 1,2-dioleoyl-sn-glycero-3-
24
phosphoethanolamine-N-[4-(p-maleimidophenyl)but-yramide (maleimide-headgroup lipid,
MPB-PE).
Primary antibodies used in this study include 1) anti-HA tag antibody from Abcam
(Cambridge, MA); 2) Alexa488-anti-phospho-CREB antibody (clone 87G3) from Cell Signaling;
3) PE-anti-CD45 (clone HI30), APC-anti-CD45 (clone HI30), PE-Cy5.5-anti-CD3 (HIT3a),
FITC-anti-CD4 (clone RPAT4), Pacific Blue
TM
-anti-CD8 (clone SK1), FITC-anti-CD8 (clone
SK1), PE-anti-IFN-γ (clone B27) and APC-anti-EGFR (clone AY13), all from BioLegend (San
Diego, CA); and 4) AlexaFluor-647 goat anti-rabbit antibody from Life Technologies.

2.3.3 Preparation of T cells for adoptive transfer and viral transduction
Thawed peripheral blood mononuclear cells (PBMCs) from healthy donors were cultured in
T cell medium (TCM) containing X-VIVO™ 15 serum-free medium (Lonza, Allendale NJ), 5%
(vol/vol) GemCell human serum antibody AB (Gemini Bio-Products, West Sacramento CA), 1%
(vol/vol) Glutamax-100× (Gibco Life Technologies), 10mM HEPES buffer (Corning), 1%
(vol/vol) penicillin/streptomycin (Corning) and 12.25 mM N-Acetyl-L-cysteine (Sigma). The
culture was supplemented with 10 ng/mL human IL-2. The PBMCs were activated and expanded
using Dynabeads
®
human T-expander CD3/CD28 (Invitrogen) at a bead:PBMC ratio of 3:1.
Activated PBMCs were transduced with viral vectors 48 hours after activation. During ex vivo
expansion, culture medium was replenished, and the T cell density was maintained between 0.5-
1 × 10
6
cells/mL.

25

2.3.4 Plasmid construction
The lentiviral vector encoding the HA-tagged CD19scFv-28-ζ CAR was constructed
based on the CD19 CAR previously reported (Milone, Fish et al. 2009). The CD19 single-chain
fragment variable (scFv) sequence, followed by the human CD8 hinge region (aa 138-184), was
codon optimized and synthesized by Integrated DNA Technologies (Coralville, IA). The
CD19/CD8 hinge gene block was combined with the transmembrane and intracellular domains
of human CD28 (aa 153-220) and the intracellular domain of human CD3ζ (aa 52-164) using
PCR. The CD8 leader sequence and HA-tag were inserted upstream of the CD19 scFv to allow
for labeling and detection of CAR-expressing cells. To make the lentiviral vector, this sequence
was inserted downstream of the human ubiquitin-C promoter in the lentiviral plasmid pFUW
using Gibson assembly, as previously described (Dai, Xiao et al. 2012).
The retroviral vector encoding the same CD19 CAR sequence was constructed based on
the MP71 retroviral vector kindly provided by Prof. Wolfgang Uckert, as described previously
(Engels, Cam et al. 2003). The cDNAs of CD19 CAR and truncated EGFP (tEGFR) linked by a
2A sequence were inserted into the MP71 vector to yield the retroviral vectors for making
CART.tEGFR cell.

2.3.5 Lentiviral and retroviral vector preparation and transduction
Lentiviral vectors were prepared by transient transfection of 293T cells using a standard
calcium phosphate precipitation protocol as described previously (Dai, Xiao et al. 2012). The
viral supernatants were harvested 48h post-transfection and filtered through a 0.45 µg filter
26
(Corning, Corning, NY). Virus supernatants were loaded into a 24-well plate with 5 × 10
5

activated human PBMCs and spun for 90 minutes at 1050g at 25°C.
Retroviral vectors were prepared by transient transfection of 293T cells using a standard
calcium phosphate precipitation protocol. Activated human PBMCs were transduced and
expanded as described previously (Han, Bryson et al. 2017).

2.3.6 Detection of receptor expression on T cell surface
HA-tagged CD19scFv-28-ζ CAR-T cells were washed with PBS and stained with rabbit
anti-HA followed by Alex647-conjugated anti-rabbit antibodies for CAR detection. Retrovirus-
transduced cells were stained with APC-conjugated anti-human EGFR for tEGFR detection.
Receptor expression was analyzed using the MACS Quant flow cytometry analyzer (Miltenyi
Biotec, Inc., San Diego, CA).

2.3.7 Synthesis of nanoparticles and drug encapsulation
Liposomes were prepared based on the established dehydration-rehydration method
previously reported (Joo, Xiao et al. 2013). To encapsulate SCH-58261 into cMLVs, 1 mg of
SCH in organic solvent was mixed with the lipid mixture to form dried thin lipid films. To label
liposome particles with DiD lipophilic dyes, DiD dyes were added to the lipid mixture in
chloroform at a ratio of 0.01:1 (DiD:lipids). Morphology of multilamellar structure of the
vesicles was analyzed and confirmed by cryo-electron microscopy, as in previous studies (Joo,
Xiao et al. 2013, Liu, Fang et al. 2014). The hydrodynamic size of cMLVs was measured by
dynamic light scattering (DLS) (Wyatt Technology, Santa Barbara, CA). The particles were
suspended in filtered water, vortexed and sonicated prior to analysis.
27

2.3.8 Nanoparticle conjugation with cells and in situ PEGylation
Chemical conjugation of nanoparticles to the T cells was performed based on a method
reported by Stephan et al (Stephan, Moon et al. 2010). For quantification of cell-bound particles,
nanoparticles were fluorescently labeled with the lipid-like fluorescent dye 1,1′-dioctadecyl-
3,3,3′,3′-tetramethylindodicarbocyanine (DiD) before conjugation, and fluorescence was
detected by flow cytometry and fluorescent microplate reader. The surface conjugation of DiD-
labeled cMLVs and CFSE-labeled CAR-T cells was further visualized using confocal
microscopy.

2.3.9 In vitro drug encapsulation and release
The amount of encapsulated SCH-58261 in the cMLV(SCH) was evaluated by C-18
reverse-phase high-performance liquid chromatography (RPHPLC) (Beckman Coulter, Brea,
CA), as previously reported (Liu, Fang et al. 2014, Kim, Liu et al. 2015). To obtain the release
kinetics of SCH from cMLVs before and after cell conjugation, cMLV(SCH) and
CART.cMLV(SCH) were incubated in 10% FBS-containing media at 37 °C and were spun down
and resuspended with fresh media daily. SCH was quantified from the harvested media every
day by HPLC.

2.3.10 Cytotoxicity assay
A modified version of a cytotoxicity assay was performed, as previously described in the
study by Kochenderfer et al., to assess the cytotoxicity of CAR-modified T cells (Kochenderfer,
Feldman et al. 2009). Non-target cells, SKOV3 cells and K562 cells, were stained with the
28
fluorescent dye 5-(and-6)-(((4-chloromethyl)benzoyl)amino) tetramethylrhodamine (CMTMR)
(Invitrogen). Target cells (SKOV3.CD19 and K562.CD19) were stained with carboxyfluorescein
diacetate succinimide ester (CFSE) (Invitrogen). The cultures were set up in triplicate in a sterile
96-well round bottom plate (Corning) at effector:target (E:T) cell ratios of 20:1, 10:1, 5:1 and
1:1. Immediately after the incubation, 7-amino-actinomycin D (7AAD, BD Pharminogen) was
added as recommended by the manufacturer. The fluorescence was analyzed by flow cytometry.
Cell cytotoxicity was calculated as [CFSE
+
7AAD
+
cells / (CFSE
+
7AAD
−
+ CFSE
+
7AAD
+
)] cells.

2.3.11 Transmigration assay
T cell transmigration assays were performed in 24-mm diameter, 3-µm pore size
transwell plates (Costar). cMLV-conjugated and unconjugated CAR-T cells (0.5 × 10
6
/well)
were plated on the upper wells, and TCM was added to the lower wells. The T cell
chemoattractant CXCL-9 (100ng/ml, Peprotech) was added to the lower wells. After incubation
at 37°C for 6 hours, T cells that migrated into the lower chamber were counted.

2.3.12 In vivo biodistribution study
For the in vivo nanoparticle biodistribution study, a xenograft tumor model was used. To
establish the tumor, SKOV3.CD19

cells in PBS solution were injected subcutaneously into the
flanks of NOD/scid/IL2Rγ
-/-
(NSG) mice. DiD-labeled cMLVs (cMLV), CD19 CAR-T cells (5 ×
10
6
) mixed with DiD-labeled cMLVs (CART+cMLV), CD19 CAR-T cells (5 × 10
6
) surface-
conjugated with DiD-labeled cMLVs (CART.cMLV), or PBS were intravenously injected into
the tumor-bearing mice. After 24 and 48 hours, indicated tissues were removed, weighed, and
macerated with scissors. DiD-specific tissue fluorescence (Abs 644 nm, Em 665 nm) was
29
quantified for each organ using the Xenogen IVIS spectrum imaging system by the USC Imaging
Core scientists blinded to the groups, and the percentage of injected dose per gram of tissue
(%ID/g) was calculated.

2.3.13 Quantification of accumulated nanoparticles at tumor sites
SKOV3.CD19

tumors were implanted into NSG mice, as described above, and CFSE-
labeled CAR-T cells and DiD-labeled cMLVs were injected into tumor-bearing mice. At the
indicated times, tumors were excised, fixed, frozen, cryo-sectioned, and mounted onto glass
slides. Fluorescence of CFSE-labeled CAR-T cells and DiD-labeled cMLVs was visualized
using a Zeiss 700 Confocal Laser Scanning Microscope (Inverted) (Carl Zeiss, Germany).
Quantification analysis was performed using Zeiss Zen microscope software

2.3.14 In vivo xenograft experiments for prevention study
SKOV3.CD19

tumors were implanted into NSG mice, as described above. After tumors
were established, mice were randomly assigned to each treatment group. Tumor growth was
measured using calipers and calculated using the formula (width
2
× length)/2. Three mice from
each group were sacrificed on day two and day 14 post-treatment. The tumor and spleen from
each mouse were harvested for further ex vivo analysis.

2.3.15 In vivo xenograft experiments for rescue study
SKOV3.CD19

tumors were implanted into NSG mice, as described above. After tumors
were established, all the mice were injected with 3 × 10
6
CART.tEGFR cells. Ten days after
initial adoptive CAR-T cell transfer, the mice were randomly assigned to receive the following
30
treatments: (1) PBS; (2) CAR-T cells; (3) CAR-T cells conjugated to empty cMLVs
(CART.cMLV); (4) a mix of CAR-T cells and cMLV(SCH) (CART+cMLV(SCH)); and (5)
CAR-T cells conjugated to SCH-loaded cMLVs (CART.cMLV(SCH)). Each mouse was injected
with 2.5 × 10
6
CAR-positive T cells. For mice treated with unconjugated cMLVs, 10
9
drug-
loaded cMLVs were co-infused with CAR-T cells. Forty-eight hours after the second adoptive T
cell transfer, the mice were sacrificed. The spleen and tumor were harvested for ex vivo assays.

2.3.16 Ex vivo analysis
Three analyses were performed: (1) anti-CD3/anti-CD28-induced intracellular IFN-γ
cytokine staining, (2) phospho-CREB and (3) Ki-67 expression in CAR-T cells. For intracellular
IFN-γ staining, a total of 0.5 × 10
6
cells were stimulated with human CD3/CD28 antibodies and
10 ng/mL Brefeldin A. The culture was incubated for 6 hours at 37°C in 96-well round bottom
plates. Fluorophore-conjugated human CD3, CD45, CD4 and CD8 antibodies were used for
immunostaining. Cytofix/Cytoperm solution (BD Bioscience) was used to permeabilize cell
membrane and perform intracellular staining according to the manufacturer’s instruction.
For intracellular phospho-CREB staining, cells were fixed with 4%
paraformaldehyde (PFA), followed by permeabilization in methanol for 30 minutes on ice. Cells
were then stained with Alexa488-conjugated anti-human phospho-CREB for 30 minutes at 4°C.
Flow cytometry analysis was carried out using the MACSQuant® Instrument from Miltenyi
Biotec (Auburn, CA).
For Ki-67 staining, cells stained with fluorophore-conjugated human CD3, CD45 and
EGFR were fixed with 80% ethanol and incubated at -20°C for 48h. Cells were washed twice
with staining buffer (PBS with 1% FBS, 0.09% NaN
3
), centrifuged for 10 minutes at 200xg and
31
resuspended to a concentration of 10
7
cells/mL. Cells were then stained with anti-Ki-67
antibody for 30 minutes at room temperature in the dark, washed twice with staining buffer,
centrifuged at 200xg for 5 minutes, and analyzed by flow cytometry.

2.3.17 Intratumoral PTX concentration measurements ex vivo
Using high performance liquid chromatography (HPLC), the PTX concentration in the
frozen tumor tissues was quantified as previously detailed (Liu, Fang et al. 2014). Briefly,
thawed tumor tissues were homogenized in ethyl acetate, with a known concentration of control
drug added to each sample as an internal standard. The samples were centrifuged and the organic
layer was transferred to a clean tube. The organic layer was evaporated under a stream of argon
and rehydrated in diluted acetonitrile. After running the samples on HPLC, the peak heights were
analyzed to determine intratumoral SCH concentration.

2.3.18 Statistics
The differences between two groups were determined with Student’s t test. The
differences among three or more groups were determined with a one-way analysis of variance
(ANOVA). Prism (GraphPad) was used to calculate statistical significance of the difference in
mean values and P values.

32
2.4. RESULTS

2.4.1. Nanoparticles stably attached to the surface of CAR-T cells

To improve the efficacy of CAR-engineered T cell therapy, we have used CAR-T cells as
chaperones to carry nanoparticles loaded with SCH-58261, a drug that can inhibit an immune-
suppressive mechanism in the TME. To express CARs on T cells, activated human PBMCs were
transduced with a lentiviral vector to deliver anti-CD19 CAR consisting of CD28 and CD3ζ
intracellular signaling domains. FACS analysis of surface CAR expression showed 50%
transduction efficiency.

Figure 2-1. Stable conjugation of cross-linked multilamellar liposome (cMLVs). (A) Schematic diagram of
adoptively transferred cMLV-conjugated CAR T cells in the presence of recipient cells in vivo. Maleimide
functionalized cMLVs loaded with A2AR small molecule inhibitors are conjugated to CAR-T cells via cell surface
thiols. (B) Quantification graph of conjugated cMLVs per T cell at different conjugation ratios. cMLVs labeled with
the fluorescent dye DiD were co-incubated with T cells over a range of ratios. The DiD fluorescence was analyzed
to calculate the number of cMLVs on the surface of each cell. (C) Percent of CD3
+
T cells conjugated with cMLVs
at 1000:1 (cMLV:T cell) conjugation ratio. (D) Representative flow cytometry analysis of percent cMLV conjugated
CD3
+
T cells. (E) Single cell confocal microscopy image of DiD-loaded cMLV-conjugated CAR T cells. These CAR
33
T cells were labeled with 1 µM CFSE and washed with PBS prior to conjugation to cMLV(DiD). Confocal
microscopy was used to visualize the cMLVs on the CAR T cell surface. Scale bar represents 10 µm. (Red: DiD
labeled cMLVs, Green: CFSE labeled CAR T cell) (n = 3, mean ± SD; ns, not significant; *p < 0.05; **p < 0.01;
***p < 0.001).

Next, crosslinked multilamellar liposomal vesicles (cMLVs) were conjugated to the surface
of CAR-T cells. According to previous reports, high levels of free thiols have been detected on
the surfaces of T, B, and hematopoietic stem cells (Stephan, Moon et al. 2010); therefore, we
used thiol-reactive maleimide headgroups present on the lipid bilayer surface of the cMLVs to
stably couple the nanoparticles to the cell surface. The conjugation was performed in two steps.
First, CAR-T cells and cMLVs containing maleimide-functionalized lipids were coincubated to
permit coupling of the liposomes to free thiols on the cell surface. After the initial coupling
reaction, the conjugated cells underwent in situ PEGylation to quench residual reactive groups on
the cMLVs. To determine the maximum number of particles that could be conjugated per T cell,
we performed nanoparticle conjugation reaction at different cMLV-to-T cell ratios (5000:1,
1000:1, 500:1, 250:1 and 100:1). At a ratio of 1000:1, the conjugation of cMLVs reached a
saturation point that resulted in an average of 287±49 surface-bound nanoparticles per cell (Fig.
2-1B). The average conjugation efficiency of the nanoparticles on the T cell population was
55.9% (Fig. 2-1C and 2-1D). Moreover, single-cell imaging and three-dimensional
reconstruction of CART.cMLVs demonstrated that the nanoparticles were distributed in several
clusters on the cell surface (Fig. 2-1E and Fig. 2-2).
34

Figure 2-2. Three dimensional (3D) live cell imaging shows conjugation of cMLVs to the surface of CAR-T cells.
Red: DiD-labeled cMLVs; Green: CFSE-labeled CAR-T cell.

2.4.2. CAR-T cells conjugated with nanoparticles maintain T cell effector functions

We next sought to test whether surface-bound cMLVs could impact key cellular
functions of CAR-T cells, such as cell cytokine secretion, cytotoxicity, and migration. CAR-T
cells with and without cMLV conjugation were co-cultured with either SKOV3.CD19 or
K562.CD19 target cells for 4 hours. CART and CART.cMLV stimulated with SKOV3.CD19
target cells induced 17.05±0.07% and 19.15±1.63% IFN-γ
+
T cell populations, respectively,
indicating that both CART and CART.cMLV were able to secrete IFN-γ with similar efficiency
(Fig. 2-3A and 2-3B). When cMLVs were labeled with DiD dye, IFN-γ was secreted from both
cells with and without surface-conjugated cMLVs. Moreover, surface conjugation of cMLVs did
not reduce CAR-T cell cytotoxicity against SKOV3.CD19 or K562.CD19 cells (Fig. 2-3C).
Supplementary Figure S1
35
Lastly, we assessed CAR-T cell transmigration capabilities in vitro. Comparable percentages of
conjugated and unconjugated cells migrated to the lower chamber of the transwell co-culture
system, indicating that CART.cMLV cells maintain their transmigration capabilities (Fig. 2-3D).
Thus, the cell surface conjugation of cMLVs does not hinder recognition of target cells, IFN-γ

secretion, cell cytotoxicity, or migration.

Figure 2-3. Conjugation of cMLV does not alter CAR T cell function. (A) IFNγ staining assays. Representative
FACS analysis of CAR-T cells either unconjugated (CART) or CAR-T cells conjugated with empty cMLVs
(CART.cMLV) stimulated with SKOV3.CD19 cells for 6 hours to detect IFN-γ release. Untransduced CAR- T cells
served as a negative control. IFN-γ was measured with intracellular staining. (B) The summarized statistics of
triplicates were shown in bar graphs. (C) In vitro cell cytotoxicity assay. CAR T cells either unconjugated (CART)
or conjugated to empty cMLVs (CART.cMLV) were co-cultured with SKOV3.CD19 cells for 6 hours and
cytotoxicity was measured. Untransduced CAR T cells served as a negative control. (n = 3, mean ± SD; ns, not
significant; *p < 0.05; **p < 0.01; ***p < 0.001)

36
2.4.3. Conjugation to CAR-T cells increases tumor localization and systemic circulation of
cMLVs

To determine whether conjugation of cMLVs to CAR-T cells could improve the
accumulation of nanoparticles to the tumor site, we performed an in vivo biodistribution study.
DiD-labeled cMLVs alone (cMLV(DiD)), mixed with CAR-T cells (CART+cMLV(DiD)), or
conjugated to CAR-T cells (CART.cMLV(DiD)) were intravenously injected into NSG mice
bearing SKOV3.CD19 tumors, and DiD-tagged cMLV accumulation was monitored in different
organs. At 24 hours, significantly higher cMLV accumulation was detected from
CART.cMLV(DiD), compared to cMLV and CART + cMLV groups, in the tumor (p<0.001),
spleen (p<0.001), lymph node (p<0.01), and lung (p<0.01). No significant difference in cMLV
accumulation was noted between cMLV(DiD) and CART+cMLV(DiD) groups in any tissues.
Additionally, no significant difference in DiD signal was detected at 24 h in circulating blood in
any group (Fig. 2-4A and 2-4B). At 48 hours, CART.cMLV(DiD) demonstrated higher cMLV
accumulation in the blood (p<0.05), tumor (p<0.05), spleen (p<0.05), and lung (p<0.05)
compared to both cMLV(DiD) and CART+cMLV(DiD) groups. Notably, CAR-T cell
conjugation to cMLVs resulted in significantly lower cMLV accumulation in the liver compared
to both cMLV(DiD) and CART+cMLV(DiD) groups at 24 (p<0.05) and 48 (p<0.001) hours
(Fig. 2-4C and 2-4D).
37

Figure 2-4. cMLVs bound to CAR T cells show more efficient infiltration to antigen-expressing tumors than
free cMLVs. Group of 3 NOD/scid/IL2rγ-/- (NSG) mice bearing subcutaneous SKOV3.CD19 tumors were
intravenously injected with 1 × 10
7
CAR-T cells conjugated (CART.cMLV), co-infused with DiD-labeled cMLVs
(CART+cMLV) or an equivalent number of DiD-labeled cMLVs alone (cMLV). After 24 hours (A) and 48 hours
(C), indicated tissues were removed, weighed, and macerated with scissors. Specific DiD tissue fluorescence for
each organ was quantified using the IVIS spectrum imaging system and calculated the mean percentage of injected
dose per gram of tissue (% ID/g) as final readout. (B, D) Representative images of DiD fluorescence from the tumor
tissues. Summarized statistics are displayed in the graphs. Data shown are pooled from two independent
experiments. (n = 3, mean ± SD; ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001)

38

2.4.4. Surface-conjugated cMLVs colocalize with CAR-T cells inside the tumor mass

We next evaluated the tumor infiltration properties of carrier CAR-T cells by confocal
imaging of histological SKOV3.CD19 tumor sections that had been treated with cMLV-
conjugated, or unconjugated, fluorescently labeled CAR-T cells. Representative confocal images
demonstrate that the surface-conjugation of cMLVs does not impede intratumoral CAR-T cell
migration (Fig. 2-5A and 2-5B). Both CART.cMLV and CART+cMLV had comparable
infiltration of T cells (Fig. 2-5C). The density of CAR-T cells in the CART.cMLV- and
CART+cMLV-treated tumor was 0.26±0.1 cells/mm
2
and 0.35± 0.2 cells/mm
2
, respectively.
However, the colocalization of CAR-T cells and cMLVs was only observed inside tumors treated
with CART.cMLV. The percentage of colocalization was 78.57±26.7% in the CART.cMLV-
treated tumors compared to no detection in CART+cMLV-treated tumors (Fig. 2-5D). These
results indicate that cMLV conjugation to the CAR-T cells is able to increase the amount of
cMLVs delivered to the tumor without impeding the migration of T cells.
39

Figure 2-5. Surface conjugated cMLVs co-localize with CAR T cells inside the tumor mass 48 hours post
CAR T cell infusion. (A) Representative confocal images of DiD- labeled cMLVs (red) conjugated CAR-T cells
(green) infiltrating SKOV3.CD19 tumor 48 hours post injection (CART.cMLVs). (B) Representative confocal
image of co-administered DiD-labeled cMLVs (red) and CAR-T cells (green) without conjugation infiltrating
SKOV3.CD19 tumor 48 hours post injection (CART+cMLVs). Scale bar, 30 µm. (C) Density of SKOV3.CD19
tumor infiltrating CAR T cells from either CART.cMLV or CART+cMLVs group. (D) Percentage of detecting
tumor infiltrated CAR T cell that are co-localized with cMLVs inside the tumor tissues. (n = 6, mean ± SD; NS, not
significant; *p < 0.05; **p < 0.01; ***p < 0.001)

40
2.4.5. CAR-T cells conjugated with nanoparticles encapsulated with A2aR antagonist
shows improved antitumor responses in vivo

To test whether the pharmacological inhibition of A2aR could prevent CAR-T cell
hypofunction in the adenosine-rich TME, we monitored tumor growth and intratumoral CAR-T
cell infiltration in vivo. SKOV3.CD19 tumor-bearing mice were assigned into six different
groups as shown in Figure 2-6A. Animals in all treatment groups showed tumor progression.
CART, CART.cMLV, CART+cMLV(SCH) and CART.cMLV(SCH) treatment groups showed
statistically significant tumor growth control, from day 2 until day 17 post ACT, when compared
to the PBS treatment group (Fig. 2-6B). Compared to other treatment groups,
CART.cMLV(SCH) treatment demonstrated the most distinguished tumor growth inhibition
resulting in significantly smaller tumors for all days measured post ACT. (Fig. 2-6B)
Consequently, CART.cMLV(SCH) treatment markedly improved the survival of the mice
compared to CART (p<0.0001, log-rank test), CART.cMLV (p<0.0001, log-rank test), and
CART+cMLV(SCH) (p = 0.0008, log-rank test) treatment groups. CART, CART.cMLV,
CART+cMLV(SCH) and CART.cMLV(SCH) had a median survival of 22, 22, 24 and 36 days
after treatment, respectively. Both PBS and cMLV treatment had median survival of 22 days.
Only CART.cMLV(SCH) treatment significantly improved the median survival compared to
PBS and cMLV groups (p=0.0003 for PBS and p=0.0002 for cMLV, log-rank test).
41

Figure 2-5. Anti-CD19 CAR T cells conjugated with SCH58261-releasing cMLVs were prevented from
developing hypofunction in SKOV3.CD19 tumors. SKOV3.CD19 cells were injected subcutaneously into the
right flank of NSG mice. Mice were randomized into six groups and treated with indicated treatments via i.v.
injections. (A) Schematic illustration of targeted in vivo delivery of different treatments. (B) Tumor size was
measured with a digital caliper (n = 8, mean ± SD; n/s, not significant; *p < 0.05; **p < 0.01; ***p < 0.001). (C)
Mouse survival curve was calculated using the Kaplan-Meier method. After indicated treatments, flank tumors were
harvested and digested for ex vivo analyses. The quantity and function of tumor infiltrated CAR-T cells were
evaluated. (D) The percentage of CD3
+
CD45
+
T cells in the tumor at 48h post treatment. (E) Ex vivo IFNγ secretion
of tumor infiltrated T cells upon stimulation with anti-hCD3 and anti-hCD28, 2 days post treatments. (F) Detection
of phosphorylated CREB expression levels in tumor infiltrated T cells 2 days post treatments. (n = 3, mean ± SD;
n/s, not significant; *p < 0.05; **p < 0.01; ***p < 0.001).

To explore how SCH affected tumor-infiltrating T cells, we analyzed T cell engraftment
and functionality. CD3
+
and CD45
+
T cell engraftment in the tumor was evaluated on day 2 post-
treatment. CART.cMLV(SCH) had higher T cell engraftment (52.96 ± 15.5%) compared to
CART (15.06 ± 1.2%, p = 0.0134), CART.cMLV (15.36 ±1.9%, p = 0.0139) and CART +
42
cMLV(SCH) (16.93 ±0.6%, p = 0.0157) treatment groups (Fig. 2-6D). Furthermore, we
evaluated the functionality of tumor-infiltrating T cells that were exposed to the adenosine-rich
immunosuppressive TME in vivo. CART.cMLV(SCH) treatment group showed significantly
higher intracellular IFN-γ expression (MFI = 744.24 ± 45.3) in CD45
+
T cells compared to
CART (MFI = 335.8 ± 91.2, p = 0.0023), CART.cMLV (MFI = 307.57 ± 53.6, p = 0.0004) and
CART + cMLV(SCH) (611.52 ± 26.21, p = 0.0118) treatment groups. The free cMLV(SCH)
treatment group, CART + cMLV(SCH) also resulted in an increased IFN-γ expression compared
to CART (p = 0.0073) and CART.cMLV (p = 0.0009) treatments, although this level is less than
observed in CART.cMLV(SCH) treatment (Fig. 2-6E).
To determine if the functional preservation of tumor-infiltrating T cells is, at least in part,
the result of A2a receptor blockade, we tested the level of phosphorylated-CREB downstream of
A2aR on isolated T cells. Our data showed that CD45
+
T cells from the CART- and
CART.cMLV-treated groups had significantly higher phosphorylated CREB compared to T cells
harvested from the CART + cMLV(SCH) and CART.cMLV(SCH)-treated groups, indicating
that SCH released from surface-engineered CAR-T cells could block A2a receptor signaling
mediated by adenosine in TME. Notably, CART.cMLV(SCH) resulted in lower phosphorylated
CREB compared to CART + cMLV(SCH) (p < 0.0001) (Fig. 2-6F). Overall, compared with free
cMLV(SCH) treatment, conjugation of cMLV(SCH) to CAR-T cells significantly prolonged
tumor growth inhibition, indicating higher efficiency in blocking the A2a receptor pathway and
preventing CAR-T cell hypofunction.

43

2.4.6. CAR-T cells conjugated with A2aR antagonist-encapsulated nanoparticles are able
to rescue hypofunctional tumor residing T cells in vivo

Although tumor-infiltrated CAR-T cells can migrate into the tumor mass, they tend to
gradually lose tumor killing and inflammatory cytokine secretion abilities after entering the
adenosine-rich tumor microenvironment (Stagg and Smyth 2010, Moon, Wang et al. 2014, Jiang,
Li et al. 2015). We hypothesized that hypofunctional tumor-residing T cells could regain their
effector functions upon the blocking of A2aR signaling with SCH. To demonstrate the potential
of our conjugated system in this application, we established an in vivo model with
hypofunctional tumor-residing CAR-T cells in the TME by an initial intravenous infusion of
CD19 CAR-T cells that express a truncated epidermal growth factor receptor (tEGFR) to the
tumor bearing mice; these CAR-T cells are designated as CART.tEGFR. The EGFR surface
marker was used to trace the initial population of hypofunctional tumor-residing CAR-T cells,
enabling us to distinguish them from the subsequent treatment dose of surface-engineered CAR-
T cells lacking EGFR. Ten days after the initial CART.tEGFR cell transfer, the rescue treatment
was infused to mice in five different groups (Fig. 2-7A).
44

Figure 2-7. Anti-CD19 CAR T cells conjugated with SCH58261-releasing cMLVs were able to rescue
hypofunctional tumor infiltrated T cells in SKOV3.CD19 tumors. (A) Schematic illustration of targeted in vivo
delivery of CAR T cells conjugated with cMLVs releasing SCH to inhibit SKOV3.CD19 tumors by rescuing tumor
infiltrated CART.tEGFR cells. (B) The waterfall plot displaying the percent change in the tumor size from baseline
at Day 35 post i.v. injections. (n = 6, mean ± SD; ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001) (C) The
percentage of CD45
+
T cells in the tumor 2 days post indicated treatments. (D) Representative FACS plots of the
percentage of CART.tEGFR cells in the tumor 2 days post indicated treatment. (E) Quantitative graph showing the
percentage of CART.tEGFR cells in the tumor 2 days post indicated treatments. (F) Detection of Ki-67 expression
in CART.tEGFR cells 2 days post indicated treatments. (G) Ex vivo IFNγ secretion of tumor infiltrated
CART.tEGFR cells upon stimulation with anti-hCD3 and anti-hCD28, 2 days post indicated treatments. IFN-γ
release was measured with intracellular staining. (H) Detection of phosphorylated CREB expression levels in
CART.tEGFR cells 2 days post indicated treatments. (n = 3, mean ± SD; ns, not significant; *p < 0.05; **p < 0.01;
***p < 0.001). All data are representative of at least two independent experiments.

45
Two days after the treatments, 5 out of 6 tumor-bearing mice that received
CART.cMLV(SCH) treatment showed over 50% reduction in tumor size, with one mouse
showing 44% reduction. The combination treatment group of CART+cMLV(SCH) showed more
than 25% reduction in tumor size in 2 out of 6 mice. Tumor-bearing mice that received either
CART or CART.cMLV had no significant reduction in tumor size, and the tumor-bearing mice
that received PBS treatment showed an overall increase in tumor size (Fig. 2-7B). Tumor-
infiltrating T cells, including CAR-positive cells, were isolated from tumors for further ex vivo
analysis. As shown in Figure 2-7C, CART.cMLV(SCH) treatment resulted in 10.79±0.3% total
T cell population, which is significantly higher than all other treatment groups (CART,
CART.cMLV, and CART+cMLV(SCH), p<0.001) (Fig. 2-7C).
We further investigated the effect of these treatments on the initial hypofunctional
CART.tEGFR cells. Tumors treated with CART, CART.cMLV, and CART+cMLV(SCH) had
25.65±2.8%, 25.35±0.5% and 28.90±3.1% intratumoral CART.tEGFR, respectively, while
CART.cMLV(SCH)-treated tumors had 63.08±5.8% CART.tEGFR cells, significantly higher
than all other groups (p<0.001) (Fig. 2-7D and 2-7E). The proliferation of CAR-T cells was
assessed by the expression of Ki-67. CART.cMLV(SCH) showed significantly higher Ki-67
expression level (MFI = 3442.4±272.3) compared to other treatment groups: CART
(MFI=1066.1±253.5, p=0.0004), CART.cMLV (MFI=1162.5±129.5, p=0.0002) and
CART+cMLV(SCH) (MFI=1044.32±224.8, p=0.0003) (Fig. 2-7F).
Next, we evaluated the ability of this CART-chaperoned drug to restore inflammatory
function of the hypofunctional CART.tEGFR population. The CART.cMLV(SCH) treatment
group showed significantly higher IFN-γ secretion in CART.tEGFR cells than that of CART,
CART.cMLV, or CART+cMLV(SCH) groups (p<0.001) (Fig. 2-7G). Evaluation of pCREB
46
expression levels in CART.tEGFR cells showed that the CART.cMLV(SCH) treatment
significantly reduced pCREB level in CART.tEGFR cell populations compared to the CART,
CART.cMLV, and CART+cMLV(SCH) treatment groups (p<0.001) (Fig. 2-7H). Taken
together, this collective evidence suggests that the surface-engineered CART system can
effectively deliver a small-molecule inhibitor of A2aR to TME, thereby rescuing hypofunctional
tumor-residing T cells in vivo.

2.5. DISCUSSION

Our strategy to enhance CAR-T cell efficacy in solid tumors was to conjugate
nanoparticles loaded with a small-molecule inhibitor of the A2a receptor pathway onto the
surface of CAR-T cells. Ohta et al. demonstrated that a variety of adenosine receptor blockades,
including antagonists such as caffeine and ZM241385, which selectively target both A2aR and
A2bR improved tumor growth suppression, reduced metastasis, and prevented
neovascularization by antitumor T cells (Ohta, Gorelik et al. 2006). This comprehensive study
strongly suggests that the A2aR pathway is a promising immunotherapeutic target to prevent
inhibition of T cell function in the TME of aggressive cancer that produces adenosine. While
there are other methods to block the A2aR pathway, such as genetically engineering CAR-T cells
with the CRISPR/Cas system or receptor siRNA knock down, the major advantage of this
strategy is the ability of the drug to affect endogenous T cells and circulating CAR-T cells, as
well as the carrier CAR-T cells themselves. In vitro, we demonstrated that cMLV nanoparticles
could be stably conjugated to the CAR-T cell surface while maintaining its ability to release
loaded drug in a sustained manner (Fig 2-8) and did not disrupt CAR-T cell effector functions.
These findings corroborate previous results reported by Stephan et al. and Huang et al. (Stephan,
47
Moon et al. 2010, Huang, Abraham et al. 2015), which demonstrated that liposomal
nanoparticles with thiol-reactive maleimide headgroups could be successfully conjugated to the
thiol-rich surface of T cells and that this could be done without altering effector functions and
transmigration capabilities of the T cells.

Figure 2-8. In vitro release rates (%) of SCH58261 in cMLVs either unconjugated (cMLV) or conjugated to CAR-T
cells (CART.cMLV). Error bars represent the standard deviation of the mean of triplicate experiments.

Our biodistribution study further shows that CAR-T cells enhance the efficacy of
therapeutic drugs by actively directing drug-loaded nanoparticles to the tumor site in vivo
(Kennedy, Bear et al. 2011, Irvine, Hanson et al. 2015, Mitchell and King 2015 ), an event driven
by the ability of CAR-T cells to migrate into the tumor mass through tumor-associated
chemokine attraction. Overall, CART.cMLVS(DiD) had the highest particle accumulation at the
tumor site at both 24 and 48 hours, reemphasizing the importance of cell-mediated delivery.
Moreover, both cMLV(DiD) and CART+cMLV(DiD) resulted in significantly higher cMLV
accumulation in the liver, which is where liposomal nanoparticles are typically cleared from the
system by Kupffer and endothelial cells (Ishida, Harashima et al. 2002, Longmire, Choyke et al.
2008). However, while CART.cMLV(DiD) showed significantly lower cMLV accumulation in
48
the liver, increased levels were observed in lymphoid tissues, such as the lymph node, spleen and
lungs. These data provide evidence that CAR-T cell-bound nanoparticles may be retained in
circulation for a longer period of time than free nanoparticles owing to reduced nanoparticle
clearance by the liver.
In order to achieve maximal drug action on hypofuctional T cells within the TME, the
drug-loaded nanoparticles must be able to reach the immune cells deep within the tumor mass. In
this regard, the CART.cMLV drug delivery system promotes the colocalization of nanoparticles
and CAR-T cells inside the tumor mass due to the innate mobility of T cells within the tumor to
deliver drugs inside the TME (Mrass, Takano et al. 2006, Boissonnas, Fetler et al. 2007).
Confocal microscopic images showed that cMLVs from the CART.cMLV group were able to
penetrate deep inside the tumor and colocalize with CAR-T cells. This maximum intratumoral
localization of cMLV could be a major factor contributing to the higher potency of
CART.cMLV(SCH) therapy.
The tumor-targeted CART.cMLV(SCH) therapeutic system was effective at preventing
hypofunction of nanoparticle-conjugated CAR-T cells. Groups treated with CART.cMLV(SCH)
demonstrated significant tumor growth suppression compared to groups without the conjugated
drug. Prophylactic CART.cMLV(SCH) treatment showed high tumor engraftment of T cells with
low CREB phosphorylation, indicating the mechanistic importance of A2aR blockade that leads
to increased T cell proliferation (Jin, Fan et al. 2010). This is supported by our observation that
CART.cMLV(SCH) had a higher percentage of tumor-infiltrated T cells and increased IFN-γ
production compared to the other groups (Ohta, Gorelik et al. 2006, Beavis, Henderson et al.
2017). (Ohta, Gorelik et al. 2006, Beavis, Henderson et al. 2017). It should be noted, however,
that the maximal levels of IFN-γ in our most successful treatment group was still significantly
49
lower than the control T cells isolated from tumor-free mice, suggesting that in addition to A2aR
signaling, CAR-T cells could be exposed to other mechanisms for immunosuppression in this
SKOV3 tumor model.
We further aimed to restore activity to hypofunctional CAR-T cells. This rescue
treatment mirrors a clinical setting where patients have pre-existing TILs or have previously
received CAR-T cell therapy. CART.cMLV(SCH) treatment resulted in significantly higher IFN-
γ secretion in the initial hypofunctional CART.tEGFR cell population upon ex vivo restimulation
compared to other treatment groups. In the CART.cMLV(SCH) treatment group, the
CART.tEGFR population also showed lower phosphorylated CREB and significantly higher cell
number compared to other groups, presumably due to the release of A2aR-mediated inhibition of
T cell proliferation (Jin, Fan et al. 2010). We also confirmed that SKOV3.CD19 cells were not
directly affected by SCH (Fig. 2-9), indicating that SKOV3.CD19 tumor reduction was mainly
achieved by the effect of SCH on the tumor-infiltrated CAR-T cells. Moreover, the immediate
reduction of tumor burden after CART.cMLV(SCH) treatment is most likely caused by the
recovery of cytotoxicity induced by the CART.tEGFR cell population. This is supported by the
fact 1) CART.cMLV(SCH) treatment alone at the same dosage in our prophylactic study could
only suppress, but not reduce, tumor growth; 2) tumor size reduction mediated by immediately
infused T cells take 5-6 days (Boissonnas, Fetler et al. 2007), which is longer than 2 days we
observed in this study.
50

Figure 2-9. Cytotoxicity of cMLVs loaded with SCH58261 against SKOV-3 human ovarian cancer cells in vitro.
Error bars represent the standard deviation of the mean of triplicate experiments.

Although CART.cMLV(SCH) has shown promising results, further improvements and
modifications could be made to expand this treatment to different clinical settings. This delivery
platform is highly flexible, and it can be applied to other drugs, cytokines, antibodies, or any
combination thereof. For example, a previous study by Stephan et al. successfully demonstrated
that the use of different therapeutic cells, such as tumor-specific T lymphocytes and
hematopoietic stem cells (HSC) as targeted delivery vehicles, markedly increased the therapeutic
efficacy of cytokines and a small-molecule inhibitor (Stephan, Moon et al. 2010). Moreover,
immune regulatory drugs could be delivered in combination with immune checkpoint blockade,
such as anti-PD-1, to further promote antitumor immunity(Beavis, Divisekera et al. 2013,
Beavis, Henderson et al. 2017). In a previous review by Iwai et al., multiple clinical studies
involving αPD1 and αPD-L1, when combined with small-molecule inhibitors of VEGF and
EGFR, for example, have shown potential for the treatment of ovarian and lung cancers (Iwai,
Hamanishi et al. 2017). Furthermore, to overcome the limitations of CAR-T cell therapy, such as
the lengthy manufacturing process and toxicities due to cytokine release syndrome or on
10
-4
10
-2
10
0
10
2
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
SCH concentraiton (µM)
Cell viability (100%)
51
target/off tumor recognition (Bonifant, Jackson et al. 2016), CAR-T cells could also be
exchanged for other “chaperone” cells, such as natural killer (NK) cells, which can be used more
universally (Siegler, Kim et al. 2017).
Cell-mediated drug delivery by surface engineering of CAR-T cells with nanoparticles
not only enables controlled drug effect on the carrier cells, but also allows active targeting to the
tissue of interest. By using CAR-T cells as chaperones, we were able to efficiently localize
nanoparticles in specific tissues favorable for T cell homing, including tumor, spleen, lungs and
lymph nodes (Masopust and Schenkel 2013). This method of combining CAR-T cell
immunotherapy with A2aR small molecule antagonists could also be applied to various types of
cancers –such as breast, prostate, brain cancers and leukemia. These cancers have been reported
to express CD73, which is associated with poor prognosis (Loi, Pommey et al. 2013, Ohta 2016).
Overall, this is a promising platform that can potentially improve the efficacy and specificity of
solid tumor therapies.

2.6 Acknowledgments
We thank Dr. Wolfgang Uckert at Humboldt University Berlin in Germany for providing
retroviral plasmid MP71. This work was supported by National Institutes of Health grants
(R01AI068978, R01CA170820, R01EB017206 and P01CA132681) and a translational
acceleration grant from the Joint Center for Translational Medicine. We also want to thank the
University of Southern California Flow Cytometry Core for the assistance

52

CHAPTER 3:

Enhanced Cancer Immunotherapy by Chimeric Antigen Receptor Modified T Cells
Engineered to Secrete Checkpoint Inhibitors

Portions of this chapter are adapted from: Natnaree Siriwon*, Si Li*, Xiaoyang Zhang, Shuai
Yang, Tao Jin, Feng He, Yujeong Kim, John Mac, Zhengfei Lu, Sijie Wang, Xiaolu Han and Pin
Wang. Clinical Cancer Research (2017), 10.1158/1078-0432.CCR-17-0867

* These authors contributed equally to this work

53
3.1. ABSTRACT

Despite favorable responses of 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. 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 was investigated by analyzing the expansion and functional
capacity of TILs. 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 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. 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.

54

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 (Mackensen,
Meidenbauer et al. 2006, Khammari, Labarriere et al. 2009, Grupp, Kalos et al. 2013, Maus,
Grupp et al. 2014, Park, Geyer 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 (Louis, Savoldo et al.
2011, Davila, Riviere et al. 2014, Gui, Han et al. 2016).
Unlike naturally occurring T cell receptors (TCRs), CARs are artificial receptors 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, Westwood 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, Jackson et al. 2016, Gill, Maus 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
55
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 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, Akiba et al. 2002). PD-L1 is also shown to
be abundantly expressed in a wide variety of solid tumors (Dong, Strome et al. 2002, Brown,
Dorfman et al. 2003, Konishi, Yamazaki et al. 2004). In contrast, the expression of PD-L1 in
normal tissues is undetectable (Dong, Strome 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, Wang et al. 2014), thereby suggesting a potential strategy
whereby CAR T therapy could be combined with PD-1 blockade in cancer treatment (Chong,
Melenhorst et al. 2017). 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
constitutively 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
56
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.

3.3. MATERIALS AND METHODS

3.3.1. 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.

3.3.2. 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
57
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).
Primary antibodies used in this study include biotinylated Protein L (GenScript, 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).

3.3.3. 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,
Cam 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
58
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.

3.3.4. 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.

3.3.5. 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 Dynabeads human T-activator CD3/CD28 (Thermo Fisher Scientific,
Waltham, MA) with cell/bead 1:1 ratio, and 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
59
10 minutes and incubated overnight at 37°C and 5% CO
2
. During ex vivo expansion, culture
medium was replenished, and cell density was adjusted to 1-2 × 10
6
/ml every two days.

3.3.6. 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).

3.3.7. 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, and PE-anti-IFN-γ 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.

60

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

3.3.9. 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) (GenScript, Piscataway, NJ) was used to pre-coat the
plate. Goat anti-mouse IgG1-HRP and anti-HA tag antibodies were used as detection antibodies,
respectively.

61

3.3.10. 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 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.

3.3.11. 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, Feldman 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
62
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
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.

3.3.12. 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
63
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.

3.3.13. 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
. For
PD-1 blockade, tumor-bearing mice were injected i.p. with 125 µg anti-human PD-1 monoclonal
antibody (mAb) (J116; Bio X Cell, West Lebanon, NH) twice a week for two weeks.

3.3.14. 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; ****, P < 0.001.
64

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 (Fig. 3-1C and 3-
1D). Furthermore, in order to better quantify the amount of anti-PD-1 secreted by CAR19.αPD1
T cells, we seeded 1×10
6
of CAR19.αPD1 T cells and cultured for 24 hours with or without
Brefeldin A (BFA). The anti-PD-1 scFv in the cell culture supernatant was then measured by
ELISA. Approximately 0.1 µg/ml of anti-PD-1 was present in the supernatant. The expression of
anti-PD-1 scFv was also monitored over the course of CAR19.αPD1 T cell expansion. We found
that the production of anti-PD-1 was maintained at a relatively stable level (0.2-0.5 µg/ml).
To evaluate the binding activity of anti-PD-1 scFv secreted by CAR19.αPD1 T cells, we
incubated the activated T cells with CAR19.αPD1 T cell culture supernatant for 30 min. The T
cells were then stained with anti-HA antibody to detect the bound anti-PD-1 on the T cell
65
surface. Compared to the control medium incubation, the secreted anti-PD-1 was able to bind to
the PD-1 on the activated T cell surface and then detected by the anti-HA antibody. To further
investigate the 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, and indeed, 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 (Fig. 3-
1E).

66

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).

3.4.2. Secreting anti-PD-1 antibody enhances the antigen specific immune response 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. 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
67
for 24 hours, we found that both CAR19 and CAR19.αPD1 T cells had a similar amount of IFN-
γ secretion (Fig. 3-2A). However, after 72 hours of stimulation with H292-CD19 cells,
CAR19.αPD1 T cells secreted significantly higher IFN-γ compared to the parental CAR19 T
cells (Fig. 3-2A). Combination of CAR19 T cells with anti-PD-1 antibody (0.6 µg/ml) resulted in
IFN-γ expression comparable to the parental CAR19 T cells after stimulation. 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 (Fig. 3-2A).
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 target cell lysis, 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 (Fig. 3-2B).
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 rate 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 (Fig 3-2C and 3-2D). The cell proliferation potential
was further assessed by cell expansion. With antigen-specific stimulation, it was shown that both
CAR19 and CAR19.αPD1 T cells significantly expanded compared to the nontransduced T cells.
Remarkably, in comparison with parental CAR19 T cells (2.4±0.2), the number of cell doublings
was significantly higher in CAR19.αPD1 T cells (3.2±0.3).
68

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).

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 (John, Devaud et al. 2013, Gargett, Yu et al. 2016). 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
69
stained for the T cell exhaustion marker PD-1. We found that the expression of PD-1 was
significantly 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 (Fig. 3-3A and 3-3B). However, without
antigen-specific stimulation, the expression of PD-1 in both CAR19 and CAR19.αPD1 T cells
maintained at a similar and stable level over the course of T cell expansion.
To further determine whether the lower expression of PD-1 in CAR19.αPD1 T cells is due to
the blocking function of secreted anti-PD-1 scFv on the binding of PD-1 detection antibody or
the downregulation of PD-1, we incubated the activated T cells with either the control medium or
CAR19.αPD1 T cell culture supernatant for 30 min before staining them with anti-PD-1
antibody. We found that the secreted anti-PD-1 scFv was able to block approximately 20% of the
binding of the PD-1 detection antibody. In tandem, we cocultured either the CAR19 or
CAR19.αPD1 T cells with target cells H292-CD19 for 24 hours. Both T cells were then
harvested and the transcriptional expression of PD-1 was measured by q-PCR. We observed that
PD-1 expression in CAR19.αPD1 T cells was significantly lower than that in parental CAR19 T
cells. This indeed confirms that CAR19.αPD1 T cells have downregulated PD-1 expression.
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
70
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 (Fig.
3-3C and 3-3D). In comparison, without antigen-specific stimulation, LAG-3 in CAR19 and
CAR19.αPD1 T cells was expressed at a similar level and remained stable over the course of T
cell expansion.

71

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 either H292-CD19 or SKOV3-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).

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
72
suppression of T cell function (Gargett, Yu 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 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 (Fig. 3-3E).

3.4.4. Anti-PD-1 engineered CAR-T cells exhibit enhances 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 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 (Fig. 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% (Fig. 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%) (Fig. 3-4D).
73

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).

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
74
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 (Fig. 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 (Fig. 3-5B and 3-C).

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.

75
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
design is shown in Figure 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
(Fig. 3-5B), T cells from mice treated with the CAR19.αPD1 regimen expanded at a higher rate
in tumor, blood, and spleen (Fig. 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, Donia 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 combination therapy had a similar CD8
+
versus CD4
+
T cell ratio compared to
CAR T cell monotherapy (Fig. 3-6C). Similarly, in the blood and spleen, the ratio of CD8
+

versus CD4
+
in CAR19.αPD1 T cell treatment was also significantly higher than that in parental
CAR19 T cell monotherapy and combination treatment groups (Figure 3-6C), though there was
little difference between the CD8
+
versus CD4
+
T cell ratio between CAR19 and CAR19.αPD1 T
cells before T cell infusion. 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 (Fig. 3-6D). We also performed the ex vivo culture and
76
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 (Fig. 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. Notably, compared to CAR19 T cell treatment, CAR19.αPD1 T
cell therapy had significantly higher anti-PD-1 concentration in the sera, although the
concentration was more than 15-fold lower than that with systemic anti-PD-1 antibody injection
(Fig. 3-6G).

77

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
+
T cells in the tumor, blood and spleen (n = 3, mean ± SEM; ns,
not significant, P>0.05; *P<0.05; ***P < 0.001). (D) The 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).

78
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, Monu 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, 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 (Dong, Strome et al. 2002, Brahmer, Tykodi et al. 2012, Ansell, Lesokhin et al.
2015). 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,
Wang 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, Morello 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. Also, the large size of
antibodies prevents them from entering the tumor mass and encountering the infiltrated PD-1-
positive T cells (Beckman, Weiner et al. 2007, Chames, Van Regenmortel et al. 2009). To
account for these inefficiencies, multiple high-dose treatments with immunomodulatory drugs or
79
antibodies are required, but this can result in side effects that range from mild diarrhea to
autoimmune hepatitis, pneumonitis and colitis (Topalian, Hodi et al. 2012). 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, Wallace et al. 2000, Maute, Gordon et al. 2015, Wang,
Erbe et al. 2015). 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 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, Parry et al.
2004, Riella, Paterson et al. 2012, Koyama, Akbay et al. 2016). In this study, the presence of
80
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, Paterson 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 (John, Devaud et al.
2013, Gargett, Yu et al. 2016). Therefore, 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, Butte 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 potential 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
81
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 is caused by the dual effects of antibody
blockade and downregulation of PD-1 surface expression (John, Devaud et al. 2013, Gargett, Yu
et al. 2016). 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 exhaustion (Thommen, Schreiner 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
82
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, Donia et
al. 2013). A previous study demonstrated that PD-1 signaling is involved in regulating the
expansion and function of CD8
+
TILs (Chauvin, Pagliano 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.
Our data show that for the CAR19.αPD1 T cell group, the ratio of CD8
+
versus CD4
+
T
cells was significantly lower in tumor than that in blood and spleen. Carter et al. reported that
CD8
+
T cells are more sensitive to PD-1-mediated inhibition than CD4
+
T cells (Carter, Fouser et
al. 2002). In tumor microenvironment, which is enriched by the PD-L1
+
tumor cells, even with
the secretion of anti-PD-1 antibody, CD8
+
T cells remain more likely to be inhibited than those in
blood or spleen, thereby causing lower ratio of CD8
+
versus CD4
+
T cells. In addition, the active
recruitment of CD4
+
T cells, especially Treg cells by the tumor, which has been shown by
Schabowsky et al., may also contribute to the lower CD8
+
versus CD4
+
T cell ratio (Schabowsky,
Madireddi et al. 2007).
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
83
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, Devaud 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, Henderson et al. 2017). In the present xenograft tumor model that is
treated with a low dose (125 µg/mouse) of anti-PD1, the antibody failed to inhibit tumor growth
or enhance the antitumor efficacy of CAR T cells even though the amount of circulating
antibody (~0.7 µg/ml) was 15-fold higher than the amount detected in the CAR19.αPD1 T cell
treatment group. This observation indicates that administration of a modest dose PD-1 antibody
blockade may not be sufficient to improve the therapeutic outcome of CAR T cell therapy.
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, which may contribute to
the observed failure of injected PD-1 blockade in enhancing the antitumor activity of CAR T cell
therapy. Indeed, according to our combination treatment regime, the PD-1 antibody was
administered 24 hours after CAR T cell infusion, and one may argue that this delay may have
also been a contributing factor to the subpar effect on enhancing the antitumor efficacy of CAR
T cell. However, our in vivo results demonstrated that CAR T cell homing to the tumor is low
within the first 48 hours, which suggest that the delay of PD-1 antibody injection may not be a
substantial factor limiting the efficacy of the combination treatment. Thus, given the low
concentration of secreted anti-PD-1 and the augmented effector function at the local tumor
84
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.
In conclusion, 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. Even though CD19 may not be an ideal antigen for the study of solid tumors, our data
indeed implies that self-secreting anti-PD-1 CAR T cells could be another promising approach to
improve the capacity of CAR T cell therapy in the treatment of solid tumors. For future studies,
other solid tumor antigens, such as mesothelin or HER2, should be investigated in order to better
evaluate the antitumor efficacy of CAR.αPD1 T cells for solid tumors. In addition, it is unclear
from our current study how the self-secreted anti-PD-1 affects immune cells other than infused
CAR T cells in tumor. Given the durable effect of PD-1 blockade on modulating the tumor
microenvironment (Pardoll 2012, Santarpia and Karachaliou 2015), it could be beneficial to
explore the capacity of CAR.αPD1 T cells to eradicate solid tumor in an immune-competent
condition, such as syngeneic mouse models. We anticipate that in such a condition, anti-PD-1-
engineered CAR T cells may be more effective in inducing tumor eradication.

3.6 Acknowledgments
We thank Dr. Wolfgang Uckert at Humboldt University Berlin in Germany for providing
retroviral plasmid MP71. This work was supported by National Institutes of Health grants
(R01AI068978, R01CA170820, R01EB017206 and P01CA132681) and a translational
acceleration grant from the Joint Center for Translational Medicine. We also want to thank the
University of Southern California Flow Cytometry Core for the assistance

85

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Abstract (if available)

Abstract The chimeric antigen receptor T (CAR-T) cell therapy has become a promising cancer immunotherapeutic method, particularly in treating B cell malignancies

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Asset Metadata

Creator Siriwon, Natnaree (author)

Core Title Engineered CAR-T cells for treatment of solid cancers

School Viterbi School of Engineering

Degree Doctor of Philosophy

Degree Program Chemical Engineering

Publication Date 08/07/2018

Defense Date 03/26/2018

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

Tag cancer,CAR-T cells,immunotheray,OAI-PMH Harvest

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Wang, Pin (committee chair), Finley, Stacey (committee member), Shing, Katherine (committee member)

Creator Email nsiriwon@gmail.com,siriwon@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c89-58781

Unique identifier UC11672466

Identifier etd-SiriwonNat-6685.pdf (filename),usctheses-c89-58781 (legacy record id)

Legacy Identifier etd-SiriwonNat-6685.pdf

Dmrecord 58781

Document Type Dissertation

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Rights Siriwon, Natnaree

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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