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Combination therapy for solid tumor
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Content
COMBINATORIAL THERAPY FOR SOLID TUMOR
by
Xiaoyang Zhang
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMICAL ENGINEERING)
December 2017
Copyright 2017 Xiaoyang Zhang
ii
DEDICATION
This thesis is dedicated to my family and my friends.
iii
ACKNOWLEDGEMENT
Firstly, I am grateful to my advisor Prof. Pin Wang for accepting me as part of his research
group. His ideas, attitude and motivation towards work and science influence me deeply and that
would benefit me for the rest of my life. Thank you very much Prof. Wang for your guidance and
support in the past four years. Meanwhile, I would like to thank the rest of my thesis committee:
Prof. Nicholas Graham, and Prof. Stacey Deleria Finley, for their insightful comments and
encouragement. I would also like to thank Prof. Katherine Shing and Prof. Noah Malmstadt, who
offered me valuable advice regarding my research and served on my qualifying exam committee.
My sincere thanks also goes to all my talented fellow labmates. I would like to show my special
thanks to Dr. Xiaolu Han, Dr. Yarong Liu and Dr. Jinxu Fang for tutoring me when I join the lab
and Dr. Si Li and John Mac for collaboration on research. I would also like to thank Dr. Paul
Bryson, Dr. Biliang Hu, and Dr. Chupei Zhang, Natnaree Siriwon, Yu-Jeong Kim, Jennifer
Rohrs, Elizabeth Siegler, Xianhui Chen, Guance Cinay and Yun Qu for their help and support. I
would also like to thank Baiyang Liu and Guanmeng Wang for their technical assistance. I feel
very fortunate and honored to have worked with them.
Last but not the least, I’d like to express my deepest gratitude to my family. Thank you all for
your unconditional love and support. I could not reach this far without your unceasing
encouragement.
iv
TABLE OF CONTENT
Dedication ........................................................................................................................... ii
Acknowledgement ............................................................................................................. iii
List of Tables .................................................................................................................... vii
List of Figures .................................................................................................................. viii
CHAPTER 1. INTRODUCTION..................................................................................................... 1
1.1 Tumor and tumor microenvironment ..................................................................................... 2
1.2 Chemotherapy ........................................................................................................................ 3
1.2.1 Liposome-assisted chemotherapy ....................................................................................... 6
1.3 Immunotherapy .................................................................................................................... 10
1.3.1 Immunotoxin ................................................................................................................. 12
1.3.2 PD-1 blockade ............................................................................................................... 14
1.3.3 CAR T cell therapy ....................................................................................................... 17
1.4 Combination therapy ............................................................................................................ 19
1.4.1 Combination chemotherapy .......................................................................................... 19
1.4.2 Combination immunotherapy ....................................................................................... 20
CHAPTER 2. CO-DELIVERY OF CARBOPLATIN AND PACLITAXEL VIA CROSS-LINKED
MULTILAMELLAR LIPOSOME FOR OVARIAN CANCER TREATMENT .......................... 21
2.1 Abstract ................................................................................................................................ 22
2.2 Introduction .......................................................................................................................... 23
2.3 Materials and Methods ......................................................................................................... 25
2.3.1 Materials ....................................................................................................................... 25
2.3.2 Preparation of cMLVs ................................................................................................... 26
2.3.3 Characterization of cMLVs ........................................................................................... 27
2.3.4 In vitro drug encapsulation and release ......................................................................... 27
2.3.5 In vitro cytotoxicity and data analysis .......................................................................... 28
2.3.6 Flow cytometry and fluorescence-activated cell sorting ............................................... 29
v
2.3.7 Evaluation of the acute toxicity .................................................................................... 29
2.3.8 In vivo antitumor study ................................................................................................. 30
2.3.9 Statistical analysis ......................................................................................................... 31
2.4 Results and discussion ......................................................................................................... 31
2.4.1 Drug encapsulation and release from cMLVs ............................................................... 31
2.4.2 In vitro cytotoxicity of drug-loaded cMLVs ................................................................. 33
2.4.3 In vivo toxicity .............................................................................................................. 38
2.4.4 In vivo anticancer efficacy of drug-loaded cMLVs ...................................................... 40
2.5 Discussion ............................................................................................................................ 43
2.6 Conclusion ........................................................................................................................... 44
CHAPTER 3. ANTI-PD-1 ANTIBODY ENHANCES ANTITUMOR IMMUNITY OF CAR-MODIFIED
T CELLS IN B-CELL LYMPHOMA ........................................................................................... 45
3.1 ABSTRACT ......................................................................................................................... 46
3.2 INTRODUCTION ............................................................................................................... 47
3.3 MATERIALS AND METHODS ......................................................................................... 49
3.3.1 Plasmid .......................................................................................................................... 49
3.3.2 Mice .............................................................................................................................. 49
3.3.3 Cell culture .................................................................................................................... 49
3.3.4 Retroviral T-cell Transduction ...................................................................................... 50
3.3.5 Anti-CD19 CAR staining .............................................................................................. 50
3.3.6 Surface immunostaining and flow cytometry ............................................................... 51
3.3.7 Intracellular staining ..................................................................................................... 51
3.3.8 Specific cell lysis assay ................................................................................................. 51
3.3.9 ELISA ........................................................................................................................... 52
3.3.10 In vivo study ............................................................................................................... 52
3.3.11 Statistical analysis ....................................................................................................... 53
3.4 RESULTS ............................................................................................................................ 53
3.4.1 Expression and activation of 1D3-28Z.1-3 anti-CD19 CAR in mouse splenic T cells . 53
3.4.2 PD-1 expression is upregulated on anti-CD19-CAR-transduced T cells following antigen-
specific stimulation ................................................................................................................ 55
3.4.3 PD-1 blockade enhances the antigen-specific immune responses of 1D3-28Z.1-3 CAR T cells
............................................................................................................................................... 56
3.4.4 PD-1 blockade enhances 1D3-28Z.1-3 CAR T cell-mediated tumor regression in established
tumor model ........................................................................................................................... 58
3.4.5 The antitumor efficacy of combined therapy is associated with tumor microenvironment
modulation ............................................................................................................................. 60
vi
3.4.6 Combination therapy specifically regulates local immune responses within tumor ..... 61
3.5 DISCUSSION ...................................................................................................................... 62
CHAPTER 4. TARGETED DEPLETION OF TUMOR STROMAL CELLS CAN ENHANCE PD-1
IMMUNOTHERAPY AGAINST MELANOMA IN MICE ......................................................... 67
4.1 ABSTRACT ......................................................................................................................... 68
4.2 INTRODUCTION ............................................................................................................... 69
4.3 MATERIALS and METHODS ............................................................................................ 72
4.3.1 Mice, cell line construction and cell culture ................................................................. 72
4.3.2 Plasmid construction and protein purification .............................................................. 72
4.3.3 Dye labeling of αFAP-PE38 ......................................................................................... 73
4.3.4 In vitro cytotoxicity of αFAP-PE38 .............................................................................. 73
4.3.5 Tumor challenge and treatment ..................................................................................... 73
4.3.6 Pharmacokinetics .......................................................................................................... 74
4.3.7 Flow cytometry analysis ............................................................................................... 74
4.3.8 Immunofluorescence imaging and Immunohistochemical analysis .............................. 75
4.3.9 RNA isolation and transcripts analysis by RT-qPCR ................................................... 75
4.3.10 Statistical analysis ....................................................................................................... 76
4.4 RESULTS ............................................................................................................................ 76
4.4.1 Construction, purification and in vitro cytotoxicity of original and mutant αFAP-PE3876
4.4.2 Both mutant and original αFAP-PE38 restrain tumor growth in vivo ........................... 79
4.4.3 Pharmacokinetics showed a faster metabolic rate of mutant αFAP-PE38 .................... 81
4.4.4 Combinatorial therapy of PD-1 blockade and mutant αFAP-PE38 shows improved antitumor
activity.................................................................................................................................... 82
4.4.5 Combinatorial therapy of PD-1 blockade and mutant αFAP-PE38 alters immunosuppressive
TMEs to promote immune stimulatory TMEs. ...................................................................... 83
4.5 DISCUSSION ...................................................................................................................... 86
References ...................................................................................................................................... 90
vii
LIST OF TABLES
Table 1-1 Marketed liposome-based therapeutics and products in clinical development. ..8
Table 1-2 Recently completed and ongoing immunotoxin clinical trials ..........................14
Table 2-1 Drug encapsulation properties and mean diameter of cMLVs ..........................32
Table 2-2 IC50 and CI values of free drug and cMLVs therapeutics against OVCAR8 and
NCI/ADR-RES ..................................................................................................................33
viii
LIST OF FIGURES
Figure 1-1 The defective vascular structure in solid tumor .................................................5
Figure 1-2 Mechanism of action of PE and PE-based immunotoxins. ..............................13
Figure 1-3 Programmed death-1 (PD-1) signaling. ...........................................................16
Figure 1-4 Chimeric antigen receptor design and evolution. .............................................17
Figure 2-1 Schematic Illustration of the Codelivery of Carboplatin and Paclitaxel via cMLVs.
............................................................................................................................................31
Figure 2-2 Release behavior of Carboplatin and Paclitaxel from cMLVs. ........................32
Figure 2-3 In vitro cytotoxicity of cMLVs (CPT/PTX) and CPT/PTX against ovarian cancer
cells. ...................................................................................................................................35
Figure 2-4 In vitro cytotoxicity of cMLVs with various CPT/PTX ratios against ovarian cancer
cells. ...................................................................................................................................36
Figure 2-5 In vitro cytotoxicity against CSCs ...................................................................37
Figure 2-6 In vivo toxicity (body weight change). ............................................................39
Figure 2-7 In vivo toxicity (Histopathological change) .....................................................40
Figure 2-8 In vivo tumor growth inhibition. ......................................................................41
Figure 2-9 TUNEL staining of apoptotic positive cells in the OVCAR8 tumor. ..............42
Figure 3-1 Expression and activation of 1D3-28Z.1-3 anti-CD19 CAR in mouse splenic T cells.
............................................................................................................................................54
Figure 3-2 Upregulation of PD-1 expression on CAR T cells following antigen-specific
stimulation..........................................................................................................................56
Figure 3-3 Anti-PD-1 enhanced the antigen-specific immune responses of CAR T cells.57
ix
Figure 3-4 PD-1 blockade enhanced CAR19 T cells mediated tumor regression of established
tumor. .................................................................................................................................59
Figure 3-5 The enhanced antitumor efficacy of combined therapy is correlated with tumor
microenvironment modulation. ..........................................................................................61
Figure 3-6 Combined therapy specifically regulates immune responses at local tumor site.62
Figure 4-1 Schematic Illustration of the Combination Therapy of αFAP-PE38 and PD-1
Blockade ............................................................................................................................71
Figure 4-2 Construction, purification and characterization of mutant αFAP-PE38. .........78
Figure 4-3 Antitumor efficacy of original and mutant αFAP-PE38 in B16-F10 tumor-bearing
mice. ...................................................................................................................................79
Figure 4-4 Pharmacokinetics of original and mutant αFAP-PE38. ...................................81
Figure 4-5 Combinatorial antitumor efficacy of mutant αFAP-PE38 and αPD-1. ............82
Figure 4-6 Combinatorial therapy of mutant αFAP-PE38 and αPD-1 increased tumor-infiltrating
T cells, the ratio of effector T cells versus Treg cells and versus myeloid-derived suppressor cells
and PD-1 expression in effector T cells within the tumor. ................................................84
Figure 4-7 Combinatorial therapy of mutant αFAP-PE38 and αPD-1 altered immunosuppressive
tumor microenvironment. ..................................................................................................85
1
CHAPTER 1. INTRODUCTION
2
1.1 Tumor and tumor microenvironment
In 2005, there were 7.6 million people died of cancer among 58 million deaths worldwide. And,
it has been predicted that cancer deaths will increase to an estimated 11.4 million dying by 2030.
Solid tumors are abnormal mass of tissue characterized by resistance to cell death and
uncontrolled cell division.
Tumor development are mainly caused by mutation or damage of proto-oncogenes that code for
proteins could induct cell differentiation and proliferation, and tumor suppressor genes that code
for proteins that produce inhibit cell growth and/or generate stimulation of apoptosis. Mutations
in both oncogenes and tumor suppressor genes are essential for tumor formation and
development and are facilitated by alterations in the tumor susceptibility genes, which encode for
a family of proteins involved with repairing DNA damage. (Pérez-Herrero and Fernández-Medarde
2015, 52-79). Initially, a mutant cell surrounded by healthy tissue would keep replicating in an
uncontrolled way which will lead to the formation of a small tumor mass. Until tumor grows to a
diffusion-limited maximal size, tumor cells absorb nutrients and remove toxic metabolites
through diffusion. Eventually, so as to grow beyond the diffusion-limited maximal size, tumor
must recruit the angiogenesis to supply the nutrients essential to fuel its continued growth
(Brannon-Peppas and Blanchette 2004, 1649-1659). It has been reported that the angiogenesis is
mainly triggered by the interaction between tumor and its microenvironment. In addition to
angiogenesis, tumor microenvironment are also involved importantly in other tumor proliferation
and progressing.
The tumor microenvironment was lately recognized as the product of a developing crosstalk
between different cells types. Except the proliferating tumor cells and blood vessels, it also
comprises infiltrating inflammatory cells and a variety of associated tissue cells. Additionally,
3
various immune effector cells are also recruited to the tumor site, however, their anti-tumor
functions are downregulated. Infiltration of inflammatory cells present in human tumors are
chronic and are dominated in regulatory T cells (Treg) as well as myeloid derived suppressor
cells (MDSC). Treg would suppress the proliferation of both CD4
+
CD25
-
and CD8
+
CD25
-
T
cells through secreting cytokines such as IL-10 and TGF-b (Bacchetta, Gambineri, and
Roncarolo 2007, 227-235). Meanwhile, MDSC could also induce the downregulation of immune
cell functions by producing an enzyme abundantly relating to L-arginine metabolism, arginase 1,
which could increase superoxide and NO production in combination with iNOS, inactivating
lymphocyte responses (Ochoa et al. 2007, 721s-726s). In addition to the failure to execute anti-
tumor functions, the immune cells in the tumor microenvironment even switch their role to
promote tumor growth. Consequently, tumor could escape from surveillance of the host immune
system. Several molecular mechanisms may be involved in tumor escape simultaneously to
impede immune cell functions or promote apoptosis of anti-tumor effector cells. In order to
eliminate tumor escape, a better understanding of cellular and molecular pathways operating in
the tumor microenvironment and creative therapeutic strategies that change the immune-
suppressive tumor microenvironment to one facilitating acute responses and efficacious anti-
tumor activity are highly needed.
1.2 Chemotherapy
Most chemotherapies target tumor cells that are constantly replicating. Many chemotherapeutic
agents interfere with cell division and activating programmed cell death by directly damaging
DNA. Other chemotherapeutic agents act indirectly by hampering mitosis or by impeding the
utilization of nucleotides that are essential for DNA synthesis in replicating tumor cells. Since
the first drugs approved by the Food and Drug Administration (FDA) for the treatment of solid
4
tumors and hematological cancers back in the forties and fifties (nitrogen mustards, anti-folate
drugs, methotrexate, etc.), chemotherapy drugs have been developed toward increasingly
effective treatments. However, drug resistance is an intractable problem limit the efficacy of
chemotherapy. The tumor cells tend to acquire mutations that endow them the resistance to the
effects of chemotherapeutic drugs. Tumors are comprise of chemo sensitive and chemo resistant
cells, which characterizes its heterogeneity and make it appear to respond to treatment initially,
but then recur when the chemo sensitive cells are eliminated and the drug resistant cells become
predominant. To overcome this limitation, modern chemotherapy almost always recruits
combinations of multiple antitumor drugs that possess different mechanisms of killing cells.
Almost all aggressive malignancies that are sensitive to chemotherapy are best treated with drug
combinations rather than single agents.
In addition to drug resistance, toxicity is another limitation for chemotherapy. Chemotherapy
they can also inhibit the rapid growth of normal cells needed for the maintenance of hair
follicles, bone marrow and gastrointestinal tract, which could cause side effects including hair
loss, myelosuppression nausea and vomiting. However, the development of targeted
chemotherapies overcome the limitation of side effects dramatically. Targeted therapies could be
obtained by direct approaches that alter specific cell signaling events by small molecules
inhibitors or monoclonal antibodies (Wu, Chang, and Huang 2006, 57-66), or by indirect
approaches using tumor-overexpressed molecular targets, to deliver chemotherapeutic agents
such as cytokines or toxins that can be conjugated to monoclonal antibodies of molecular targets,
this approach could achieve higher concentrations of chemotherapeutic agents in tumor sites and
decreasing the peripheral toxicity.
5
Figure 1-1. The defective vascular structure in solid tumor.
(Stockhofe et al. 2014)
Another approach to avoid the non-specificity of the conventional chemotherapy, is
encapsulating the chemotherapeutic agents in nanocarriers. Especially, a vast of studies about the
application of nanocarriers in chemotherapy have been done in last two decades, due to its
unique advantages. Firstly, nanocarriers can protect the drug from degradation and, reduce the
renal clearance and increase its half-life in the bloodstream, augment the payload of cytotoxic
drugs, allow the control of the release kinetics of the antitumor drugs, and improve the solubility
of those insoluble agents (Peer et al. 2007, 751-760; Davis and Shin 2008, 771-782; Danhier, Feron,
and Préat 2010, 135-146). Secondly, as shown in Figure 1-1 enhanced permeability and retention
effect combining with the poor lymphatic drainage of tumors allow the passive accumulation of
the nanocarriers in tumor tissues, releasing the chemotherapeutic agents in the vicinity of the
tumor.
6
1.2.1 Liposome-assisted chemotherapy
Liposomes are the most well-known and well-investigated nanoparticles for drug administration.
They have the ability to enhance the effectiveness of therapies for a plenty of biomedical
applications from different aspects including stabilizing therapeutic compounds, overcoming
obstacles to cellular and tissue uptake, and improving biodistribution of compounds to target
sites in vivo (Hua and Wu 2013, 143; Sercombe et al. 2015, 286). Liposome is a type of
phospholipid vesicle consisting of one or more concentric lipid bilayers enclosing discrete
aqueous spaces. Both lipophilic and hydrophilic compounds could be encapsulated in the
liposomes. Hydrophobic therapeutic agents are inserted into the bilayer membrane, and
hydrophilic therapeutic agents can be encapsulated in the aqueous layer simultaneously (Hua and
Wu 2013, 143; Sercombe et al. 2015, 286).
Additionally, different macromolecules, such as DNA, proteins and imaging agents are allowed
to be delivered due to the relatively big aqueous center and biocompatible lipid exterior.
Liposomes provide several attractive advantages containing biocompatibility, ability to achieve
self-assembling, ability to encapsulate large drug payloads, and a variety of physicochemical and
biophysical properties that could be modified to adjust their biological characteristics(Hua and
Wu 2013, 143; Sercombe et al. 2015, 286). Their formulations are characterized by properties
such as particle size, charge, number of lamellae, lipid composition, and surface modification
with polymers and ligands-these all govern their stability in vitro and in vivo (Hua and Wu 2013,
143). Liposomes act as a shield to protect compounds from early inactivation, degradation and
dilution in the circulation. It is well known that liposomes are pharmacologically inactive with
minimal toxicity because of their natural phospholipids composition (Hua and Wu 2013, 143). In
recent years, their clinical translation has progressed incrementally (Sercombe et al. 2015, 286).
7
The first generation of liposomes to be developed were conventional liposomes. Their
composition is a lipid bilayer that consist of cationic, anionic, or neutral (phospho) lipids and
cholesterol, which encloses an aqueous core. The reduced the toxicity of compounds in vivo and
the modified pharmacokinetics and biodistribution make the conventional liposomes could
enhance drug delivery to diseased tissue compared to free drug. However, the short circulation
time in the bloodstream limit its therapeutic efficacy. This rapid clearance caused by
opsonization of plasma components and uptake by fixed macrophages from the
reticuloendothelial system (RES), mainly in the liver and spleen (Hua and Wu 2013, 143;
Sercombe et al. 2015, 286).
In order to enhance liposome stability and prolong their circulation times in the bloodstream,
sterically-stabilized liposomes were introduced. Polyethylene glycol (PEG), which a type of
hydrophilic polymer, has been shown to be the optimal choice for obtaining sterically-stabilized
liposomes. With the protection of PEG, in vivo opsonization with serum components, and the
rapid recognition and uptake by the RES could be decrease significantly. As a result, the efficacy
of encapsulated therapeutics would be improved. In addition to the reduction of the elimination
of drugs by prolonging blood circulation and providing accumulation at pathological sites, side
effects could also be attenuated(Sercombe et al. 2015, 286; Ishida et al. 2006, 15-25).
8
Table 1-1. Marketed liposome-based therapeutics and products in clinical development.
Many liposomal-based drugs that are FDA approved or currently in clinical trials are
summarized in Table 1-1(Sercombe et al. 2015, 286). Some of the most successful delivery
methods rely on PEG conjugated lipids. Actually, the first FDA approved nano-drug,
doxorubicin, is delivered by PEGylated liposomes. It usually combined with other therapeutics,
Doxil has been applied to treat various types of cancer including AIDS-related Kaposi’s
sarcoma, leukemia, and ovarian, breast, bone, lung, and brain cancers (Sercombe et al. 2015,
9
286). Doxil is also deemed as an effective alternative to conventional doxorubicin in patients
with pre-existing cardiac dysfunction. When doxorubicin is incorporated in PEGylated
liposomes, the minimized uptake and clearance by the RES and prolonged the serum and plasma
half-life allows the Doxil to accumulate in the tumor tissue, rather than in non-target healthy
tissues. Furthermore, the usage of Doxil could avoid the release of doxorubicin when it pass
through the myocardium and then reduce the side effects to cardiac muscle cell (Rahman, Yusuf,
and Ewer 2007, 567-583). Finally, Doxil also could avoid the high plasma peak levels of free
drug, which induces cardiotoxicity.
However, there are still some issues with Doxil. For instance, slow and incomplete drug release
could still lead to low drug bioavailability within tumor tissue, limiting, in turn, therapeutic
activity. Furthermore, a lack of controlled-release properties of encapsulated drug may lead to
toxic side effects, such as palmar-plantar erythrodysesthesia that is thought to result from
unwanted drug distribution to skin during prolonged circulation of Doxil.
In our previous study, we investigated a cross-linked multilamellar liposome as anticancer
therapeutic nanocarrier and it has been demonstrated that this cross-linker multilamellar
liposome could achieve a improve drug release compared to Doxil and lower the systemic
toxicity and enhance therapeutic efficacy (Joo et al. 2013, 3098-3109). Results from our another
previous study indicated that the superior applicability of the cross-linked multilamellar
liposome in combination therapy due to both of the extended exposure of drugs to cancer cells
and codelivery of drug combination at synergistic dose ratios to the site of action without
inducing significant systemic toxicity (Liu et al. 2014a).
10
1.3 Immunotherapy
Immunotherapy is an approach which treat cancer by inducing or boosting an immune response
against it. This approach has been investigated, mostly outside of mainstream cancer research,
for over a century (Coley 1910, 1-48; Khalil et al. 2016a, 273-290). However, in the past decade,
cancer immunotherapy start to show the capacity of improving the overall survival of patients
with advanced-stage cancer in phase III clinical trials (Kantoff et al. 2010, 411-422; Hodi et al.
2010, 711-723; Robert et al. 2015, 2521-2532; Robert et al. 2015, 320-330), and bring
unparalleled interest to this field.
Generally, the human natural immune system seems unable to recognize and identify cancer as a
foreign invader. First reason is that cancer cells are derived from human body and aren't like
external invaders such as viruses and bacteria. That means cancer cells are altered versions
(mutations) of normal cells and don't produce a unique feature like an antigen that will trigger an
immune response. Secondly, cancer cells may also suppress immunity, which may contribute to
the immune system's failure to recognize mutant portion of cancer cells.
Immunotherapy is based on the concept that immune cells or antibodies that can recognize and
kill cancer cells can be produced in the laboratory and then given to patients to treat cancer. Two
types of immunotherapy have shown the particularly efficacy over the past decade: monoclonal
antibody (mAb) therapy and adoptive cellular therapy (ACT).
Monoclonal antibodies are a type of proteins that can specifically recognize and bind to antigens
expressed on the surface of tumor cells or surface of immune cells. Tumor-specific monoclonal
antibodies are capable to bind to tumor associated antigens and elicit direct or indirect immune
responses to destroy the tumor cells (Scott, Allison, and Wolchok 2012, 14). Monoclonal
11
antibody therapies can cause side effects, but they're generally milder than those of
chemotherapy. Because they're designed to target and attack specific substances, they tend to
leave normal cells unharmed (Scott, Allison, and Wolchok 2012, 14). In clinical setting, the
naked antibodies and conjugated antibodies are widely used by doctors. Naked antibodies could
attack cancer cell directly once it recognizes the tumor associated antigen on cell’s surface. In
contrast, the conjugated antibodies, which have toxins attached to them (immunotoxin), could
deliver the toxin to the cancer cells and kill them.
Instead of killing tumor cells directly, immunomodulatory mAbs try to stimulate T-cell function
by blocking or activating regulatory receptors is sufficient to elicit the regression of some
malignancies. Importantly, checkpoint blockade is a method by which T-cell function is restored
with mAbs that could block their inhibitory receptors, which received lots of attention in past
decade, due to it led to unprecedented response in patients with advanced-stage tumor. Indeed,
programmed cell-death protein 1 (PD-1), has resulted in impressive antitumor activity and is now
approved by the FDA to treat patients with non-small-cell lung cancer (NSCLC) and melanoma.
Adoptive immunotherapy for cancer has a long history. And the important finding that
hematopoietic stem cell transplantation from sibling donors was more effective at preventing
relapse of leukemia compared with syngeneic donors provided the initial rationale for adoptive
T-cell therapy (Dudley and Rosenberg 2003, 666-675). Furthermore, the direct isolation and ex
vivo activation of the tumor-infiltrating lymphocytes (TILs) was tested in multiple early-phase
studies and resulted in durable responses in melanoma (Rosenberg et al. 2011, 4550-4557).
Additionally, studies of chimeric antigen receptor (CAR) T-cells conducted at several academic
institutions have acquired tremendous attention for their clinical development. Transgenic
chimeric antigen receptors expressed T-cells could be redirected to target tumor antigens via
12
human leukocyte antigen (HLA)-independent recognition. Importantly, in clinical trials, T-cells
expressing CAR were infused into adult and pediatric patients with B-cell malignancies,
sarcoma, and neuroblastoma and these results have demonstrated the potential of this approach
(Porter et al. 2011, 725-733; Brentjens et al. 2013, 177ra38).
1.3.1 Immunotoxin
Immunotoxins derive their potency from the toxin and their specificity from the antibody or
antibody fragment to which they are attached. Two types of bacterial toxins derived immunotoxin are
usually applied in clinical trials. Both diphtheria toxin (DT, 580 amino acids) and Pseudomonas
aeruginosa exotoxin A (PE, 613 amino acids) are produced as single polypeptide chains, each of
which has three functional domains. Domain Ia is the cell-binding domain of PE located at the N
terminus, domain II has translocation activity and domain III, which is located at the C terminus,
catalyses the adenosine diphosphate (ADP)-ribosylation and inactivates elongation factor 2
(EF2), which could induce the inhibition of protein synthesis and cell apoptosis (
2). The third generation PE-based immunotoxins, which is most currently used, only contains Fv
portion of an antibody required to recognize target cell and the minimum translocation and cell-
killing domains to kill cells (Becker and Benhar 2012).
13
Figure 1-2. Mechanism of action of PE and PE-based immunotoxins.
(Becker and Benhar 2012)
Our approach to cancer treatment has been revolutionized by the improvement of immunotoxin
therapy. A variety of immunotoxin now approved for clinical use and are very efficacious
against several types of cancer (Table 1-2) (Pastan et al. 2006, 559+). However, most success
come from treating hematologic tumors. Because the poor penetration into tumor masses and the
immune response to the toxin component of the immunotoxin pose the obstacles to successful
treatment of solid tumors, also limit the number of cycles that can be given. New approaches are
being pursued to overcome these limitations (Pastan et al. 2007, 221-237).
14
Table 1-2. Recently completed and ongoing immunotoxin clinical trials.
1.3.2 PD-1 blockade
PD-1 is a negative regulator of T-cell activity expressed on many immunologic cells, including T
cell, B cells and natural killer cells, which can interact with its two ligands PD-L1 and PD-L2 to
inhibit the function of T cells at a variety of stages of the immune response. When PD-1 interact
with its ligand, kinase signaling pathways that normally lead to T-cell activation would be
15
downregulated through phosphatase activity (Fig1-3). Mice with deficiency of PD-1 have a
distinct autoimmune phenotype from mice with deficiency of CTLA-4. That because CTLA-4 is
primarily believed to regulate immune responses early in T-cell activation, whereas PD-1 is
primarily believed to inhibit effector T-cell activity in the effector phase within tissue and
tumors. Therapeutic blockade of the PD-1 pathway have shown the ability to affect the function
of T cell (Postow, Callahan, and Wolchok 2015, 1974-1982)(Dong et al. 2002c, 793-800; Chinai
et al. 2015, 587-595)(Postow, Callahan, and Wolchok 2015, 1974-1982).
A variety of antibodies that block the PD-1 mediated signaling have been applied in clinical
development. Even though these antibodies have different structures, they can be mainly
categorized into two main types: PD-1 targeting antibodies (nivolumab) and PD-L1 targeting
(MPDL3280A, MEDI4736, BMS-936559, MSB0010718C) (Chinai et al. 2015, 587-595). AMP-
224 is a PD-L2 recombinant protein that does not directly target PD-1 or PD-L1, but it could
deplete PD-1-positive T cells (Infante et al. 2013, 3044-3044).
Both nivolumab and pembrolizumab have shown highly durable response rates without
significant toxicity in large phase I studies involving patients with advanced melanoma, non–
small-cell lung cancer, renal cell carcinoma, and other solid tumors (Postow, Callahan, and
Wolchok 2015, 1974-1982).
16
Figure 1-3. Programmed death-1 (PD-1) signaling.
(Chinai et al. 2015, 587-595)
Although success of PD-1 blockade in large phase I studies have received lots of attention,
randomized phase III data in patients with melanoma are now emerging. In one study of patients
with melanoma who had experienced progression on ipilimumab, nivolumab resulted in a 32%
overall response rate compared with 11% for chemotherapy (dacarbazine or
carboplatin/paclitaxel) with less frequent high-grade treatment-related adverse events (Weber et
al. , 375-384). A separate phase III study was stopped early by an independent data monitoring
committee because patients with melanoma treated by nivolumab had improved overall survival
compared with patients treated with dacarbazine chemotherapy (Postow, Callahan, and Wolchok
2015, 1974-1982).
Pembrolizumab has also been demonstrated to lead to attractive tumor responses and was
recently approved by FDA for patients with melanoma previously treated with BRAF inhibitor.
Although nivolumab and pembrolizumab have predominantly demonstrated activity in solid
17
tumors, pidilizumab has mostly been clinically evaluated in hematologic malignancies, with
responses shown as monotherapy and in combination therapy with rituximab (Postow, Callahan,
and Wolchok 2015, 1974-1982).
Targeting PD-L1 is an alternative promising approach to targeting PD-1. However targeting PD-
L1 would lead to different biologic effects compared with targeting PD-1. PD-L1 also is believed
to have the ability to not only bind PD-1 but also to exert negative signals on T cells through
interacting with B7 (Butte et al. 2017, 111-22). PD-L1-blocking antibodies inhibit this
interaction, whereas PD-1 blocking antibodies do not. One more difference is that PD-L1
antibodies just block the interaction between PD-1 and PD-L1, not for the interaction between
PD-1 and PD-L2, the effect of this interaction still remains unknown (Postow, Callahan, and
Wolchok 2015, 1974-1982).
PD-L1 antibodies also show objective responses in early-phase clinical trials in a number of
malignancies including tumor types such as bladder cancer, head and neck cancer, and GI
malignancies (Powles et al. 2014, 5011-5011).
1.3.3 CAR T cell therapy
Figure 1-4. Chimeric antigen receptor design and evolution.
18
CAR T-cell therapy could specifically target an extracellular antigen independent of the peptide-
HLA complex through a single-chain variable fragment (scFv) derived from the variable heavy
and variable light chains of an antibody. The first-generation CAR is compsed of a scFv, a
transmembrane region, and the CD3ζ chain (the signaling domain of the TCR complex), which
could provide only activation signal 1 to T cells, and has been shown to lead to T-cell anergy
upon repeated antigen specific stimulation. Second-generation CARs incorporate an additional
co-stimulatory domain that provides activation signal 2 upon the scFv engaging the target
antigen. The signaling domains of CD28 or 4-1BB are the most widely used co-stimulatory
molecules (Khalil et al. 2016b, 273-290). Importantly, almost all clinical trials performed to date,
have used second-generation CARs. In third generation CARs, two co-stimulatory domains are
added to the above design, although direct comparisons with second-generation CARs have not
yet been conducted in the clinical setting. The design and evolution of CARs are shown in Figure
1-4 (Fesnak, June, and Levine 2016, 566-581).
CAR-T-cell therapy showed remarkable clinical efficacy in trials in hematological malignancies,
but this modality have not received impressive successes in solid tumors yet. Solid tumors pose
several challenges not seen in B-ALL. First one is the microenvironment in solid tumor can be
considerably more immuno-suppressive compared with B-ALL. Furthermore, the high antigen
heterogeneity across the same malignancy in solid tumors makes the antigen selection more
difficult. Additionally, potential target antigens in solid tumors are more likely to be expressed in
other essential organs, thus ‘on-target, off-tumor’ toxicity is more problematic (Khalil et al.
2016b, 273-290). Several targets for solid tumors have already been investigated in clinical
studies include mesothelin for the treatment of mesothelioma, pancreatic and ovarian cancer;
19
disialoganglioside GD2 and EGFRvIII189 for CNS malignancies; and mucin-16 for the
treatment of ovarian cancer (Khalil et al. 2016a, 273-290).
1.4 Combination therapy
Combination therapy, a treatment modality that combines two or more therapeutic agents, is a
cornerstone of cancer therapy. The amalgamation of anti-cancer therapies enhances efficacy
compared to the mono-therapy approach because it targets key pathways in a characteristically
synergistic or an additive manner. This approach potentially reduces drug resistance, while
simultaneously providing therapeutic anti-cancer benefits, such as reducing tumor growth and
metastatic potential, arresting mitotically active cells, boosting immune response, reducing
cancer stem cell populations, and inducing apoptosis (Mokhtari et al. 2017, 38022-38043).
1.4.1 Combination chemotherapy
The use of combination chemotherapy to treat cancer was inspired in the 1960s when scientists
wondered whether the approach to treating tuberculosis-using a combination of antibiotics to reduce
the risk of resistance – would work for the treatment of cancer as well. Using this approach, cancers
that had previously been almost universally fatal such as acute lymphocytic leukemia and Hodgkin’s
lymphoma became largely curable. Since that time, combination chemotherapy has been adopted for
the treatment of many other cancers as well.
Combination chemotherapy offers several advantages comparing to using single agents alone: (1)
Decreasing the chance that a tumor will be resistant to the treatment. (2) Being able to address
several targets in the cancer process at the same time. (3) More efficient to tumor heterogeneity. (4)
Reduce anti-cancer drugs dosage to reduce toxic effects.
20
1.4.2 Combination immunotherapy
The low response rate is the critical challenge for single immunotherapy, even though it has
shown efficient anti-tumor activity across a wide spectrum of cancers. It has been supposed that
the reason induce the low response could be the numerous molecules and pathways are involved
in the escape of tumors from immune destruction and the suppressive tumor microenvironment.
Combination immunotherapy could provide the rationale to solve this problem by introducing
different arms of the immune response and then lead to additive or synergistic effect (Khalil et al.
2016a, 273-290).
In order to produce a durable antitumor response in patients who would not benefit from mono-
immunotherapy, the safest and most efficacious combinations of immunomodulatory antibodies
and/or adoptive cellular therapeutics is highly anticipated.
21
CHAPTER 2. CO-DELIVERY OF CARBOPLATIN AND PACLITAXEL VIA
CROSS-LINKED MULTILAMELLAR LIPOSOME FOR OVARIAN CANCER
TREATMENT
22
2.1 Abstract
Carboplatin (CPT) and paclitaxel (PTX) used in combination is one of the most effective
treatments for ovarian cancer. However, due to their distinct pharmacokinetics, it is very difficult
to use traditional administration methods to co-deliver CPT and PTX in a controlled manner.
Here, we used cross-linked multilamellar liposomal vesicles (cMLVs) to encapsulate and deliver
this drug combination to ovarian cancer cells at a controlled ratio. In vitro cytotoxic assays
determined the strongest anti-tumor synergism to be when the drug combination was delivered at
a 1:1 CPT/PTX molar ratio. Moreover, we demonstrated that our co-encapsulation strategy
reduced cytotoxity and resulted in a stronger anti-tumor effect when compared to free drug
combinations and individual drug-loaded cMLVs in an OVCAR8 ovarian cancer xenograft
mouse model.
23
2.2 Introduction
Ovarian cancer is the sixth most common cancer and the seventh most common cause of cancer
deaths in women (Pujade-Lauraine et al. 2010, 3323-3329).Each year, approximately 2.4 million
women are diagnosed with ovarian cancer worldwide. Currently, the relative five-year survival
rate for ovarian cancer is only 44.2 percent which is much lower than that of other cancers that
affect women. Chemotherapy based on platinum drugs (carboplatin or cisplatin) in combination
with paclitaxel is a standard treatment for patients with ovarian cancer (Ohta et al. 2009, 4639-
4647). Platinum drugs damage and bind to DNA, which induces the inhibition of DNA
replication (Basu and Krishnamurthy 2010, 10.4061/2010/201367), while paclitaxel induces cell
cycle arrest at the G2/M phase via binding to the beta subunit of micro-tubulin, followed by the
induction of apoptosis. In addition, platinum drugs and paclitaxel trigger different apoptosis
signaling pathways in ovarian cancer cells (Villedieu et al. 2006, 507-519; Villedieu et al. 2007,
373-384; Xiao et al. 2006, 10166-10173). It has been shown that the co-administration of
platinum drugs with paclitaxel results in a synergistic improvement in tumor killing due to their
distinct mechanisms of action (Ramalingam et al. 2010, 56-62). One notable platinum drug is
carboplatin, which shows no renal toxicity unlike the widely used cisplatin. Moreover, for a
comparable level of treatment effectiveness, it has been determined that the combination of
carboplatin and paclitaxel has better tolerability and quality of life than the combination of
cisplatin and paclitaxel. Therefore, the combination of carboplatin and paclitaxel should be
regarded as a very important regimen for treating patients with ovarian cancer (Katsumata et al.
2009, 1331-1338) and even platinum-sensitive recurrent ovarian cancer (Pujade-Lauraine et al.
2010, 3323-3329).
24
The difficulty of paclitaxel's administration is its primary limitation in clinical settings. Because
paclitaxel is highly hydrophobic, it must be administered with a combination of dehydrated
alcohol and Cremophor EL (polyoxyethylated castor oil) as an adjuvant. However, this can lead
to serious side effects including neurotoxicity, nephrotoxicity and hypersensitivity reactions (Ma
and Mumper 2013, 1000164). Another clinical limitation of paclitaxel is its poor
pharmacodynamics (cytochrome P450 metabolism) and pharmacokinetics profiles (t1/2 in human:
2.09 h) (Vasantha et al. 2011, 322-328), which may hinder the drug’s accumulation within
tumors and compromise its in vivo efficacy. In contrast, carboplatin is eliminated and
metabolized much slower in vivo because of its heavy metal backbone (van der Vijgh, Wim JF
1991, 242-261), which could lead to long-term drug exposure of cancer cells. However, the bio-
transformations of carboplatin, which form in the bloodstream after hydrolysis and binding to
plasma proteins in the plasm, are directly associated with its acute and chronic hematopoietic
toxicity, hepatotoxicity, and neurotoxicity (Sooriyaarachchi, Narendran, and Gailer 2011, 49-55).
Thus, a more effective paclitaxel/carboplatin combination therapy is needed to encapsulate
paclitaxel without using any harmful organic solvents, improve its bioavailability, and increase
its exposure to tumor cells, while simultaneously minimizing the acute and cumulative long-term
chronic toxic side effects of carboplatin. This could be achieved by modifying the bio-
distributions and tumor uptake of both drugs via a nanoparticle drug delivery system that allows
for the pharmacokinetic and pharmacodynamic behaviors of both drugs to be determined by the
pharmacokinetics and pharmacodynamics of the nanoparticles.
Over the past few decades, significant effort has been devoted to developing nanotechnology for
drug delivery since it allows for the targeted delivery of drugs to tissues of interest and hence, a
significant decrease of toxic, off-target side effects (Wilczewska et al. 2012, 1020-1037; Liu,
25
Rohrs, and Wang 2014, 818-828; Liu et al. 2013, 378380; Zhao et al. 2014, 15319-15325). A
variety of nanocarriers for the delivery of either paclitaxel or carboplatin have been developed
(Danhier et al. 2009, 11-17; Sadhukha and Prabha 2014, 1029-1038; Nanjwade et al. 2010, 176-
180). However, the main challenge is to encapsulate these two drugs, which have distinct
physiochemical properties, into a single nanocarrier. Even though a few studies of the co-
delivery of cisplatin with paclitaxel have been published recently (Xiao et al. 2012, 6507-6519;
Desale et al. 2013, 339-348; Cai et al. 2015, 456-468), research on the co-delivery of carboplatin
with paclitaxel has rarely been reported.
In this study, we investigated whether the cMLVs reported in our previous studies (Joo et al.
2013, 3098-3109; Liu et al. 2014, 1651-1661) could achieve the synergistic combinatorial
delivery of hydrophobic paclitaxel and hydrophilic carboplatin. Different combination ratios of
paclitaxel and carboplatin were encapsulated into cMLVs and the corresponding combination
effects (antagonistic, additive, or synergistic) were determined by XTT assays. We then
determined whether cMLVs could minimize the in vivo toxicity of the drug combination. Lastly,
we determined the synergistic effects of drug combination-loaded cMLVs in ovarian cancer
mouse models.
2.3 Materials and Methods
2.3.1 Materials
OVCAR8 and NCI/ADR-RES cell lines were kindly provided as gifts by Dr. Nouri Neamati
(University of Southern California, School of Pharmacy, and Los Angeles, CA) and maintained
in RPMI-1640 with supplemented with 10% fetal bovine serum (FBS) and 2 mM of L-glutamine
in 5% CO2 environment.
26
All lipids were obtained from NOF Corporation (Japan): 1,2-dioleoyl-sn-glycero-3-
phosphocholine (DOPC), 1,2-dioleoyl-sn-glycero-3-phos-pho-(10-rac-glycerol) (DOPG), and
1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[4-(p-maleimidophenyl) butyr-amide
(maleimide-headgroup lipid, MPB-PE). Carboplatin, paclitaxel were purchased from Sigma-
Aldrich (St. Louis, MO).
BALB/c and Athymic mice (Charles River Laboratories) were used for in vivo toxicity and anti-
tumor studies. Mice were held under specific pathogen-reduced conditions in the Animal Facility
of the University of Southern California (Los Angeles, CA, USA). All experiments were
performed in accordance with the guidelines set by the National Institute of Health and the
University of Southern California on the Care and Use of Animals.
2.3.2 Preparation of cMLVs
Liposomes were prepared based on the conventional dehydration−rehydration method. All lipids
were combined in chloroform, at a molar lipid ratio of DOPC−DOPG−MPB = 4:1:5, and the
chloroform in the lipid mixture was evaporated under argon gas. The lipid mixture was further
dried under vacuum overnight to form dried thin lipid films. In order to prepare cMLVs (CPT),
cMLVs (PTX) and cMLVs (CPT +PTX), paclitaxel was mixed with the lipid mixture in organic
solvent before formation of the dried thin lipid films. The resultant dried film was hydrated in 10
mM Bis-Tris propane at pH 7.0 with carboplatin by vigorous vortexing every 10 min for 1 h and
then applied with four cycles of 15 s sonication (Misonix Microson XL2000, Farmingdale, NY)
on ice at 1 min intervals for each cycle. To induce divalent-triggered vesicle fusion, MgCl2 was
added at a final concentration of 10 mM. The resulting multilamellar vesicles were further cross-
linked by addition of dithiothreitol (DTT, Sigma-Aldrich) at a final concentration of 1.5 mM for
27
1 h at 37 °C. The resulting vesicles were collected by centrifugation at 14 000 g for 4 min and
then washed twice with phosphate-buffered saline (PBS). For pegylation of cMLVs, the particles
were incubated with 1 μmol of 2 kDa PEG-SH (Laysan Bio Inc. Arab, AL) for 1 h at 37 °C. The
particles were then centrifuged and washed twice with PBS. The final products were stored in
PBS at 4 °C.
2.3.3 Characterization of cMLVs
The hydro-dynamic size and size distribution of cMLVs were determined by dynamic light
scattering (Wyatt Technology, Santa Barbara, CA).
2.3.4 In vitro drug encapsulation and release
Concentrations of CPT and PTX were then measured by C-18 reverse-phase high-performance
liquid chromatography (RP-HPLC) (Beckman Coulter, Brea, CA) at 227 nm. To study the
encapsulation capacity of cMLVs corresponding to CPT, cMLVs (CPT) and cMLVs (CPT+PTX)
were collected and washed twice with PBS, followed by lipid extraction of vesicles with 1%
Triton X-100 treatment. To determine the encapsulation capacity of cMLVs with respect to PTX,
the cMLVs(PTX) and cMLVs(CPT+PTX) suspensions were diluted by adding water and
acetonitrile to a total volume of 0.5 mL. Extraction of paclitaxel was accomplished by adding 5
mL of tert-butyl methyl ether and vortex-mixing the sample for 1 min. The mixtures were
centrifuged, and the organic layer was transferred into a glass tube and evaporated to dryness
under argon. Buffer A (95% water, 5% acetonitrile) was used to rehydrate the glass tube. To test
CPT and PTX concentration, 1 mL of the solution was injected into a C18 column, and the CPT
and PTX were detected at 227 nm after different retention times (flow rate 1 mL/min). To obtain
the release behavior of CPT and PTX from cMLVs, the releasing media was removed from
28
cMLVs incubated in 10% FBS-containing media at 37 °C and replaced with fresh media daily.
The concentration of CPT and PTX in the removed media was quantified by HPLC.
2.3.5 In vitro cytotoxicity and data analysis
To evaluate the cytotoxicity, the viability of cell treated by drug-loaded cMLVs was assessed
using the Cell Proliferation Kit II (XTT assay) from Roche Applied Science according to the
manufacturer’s instruction.
The OVCAR8 and NCI/ADR-RES cells in 100 µl of 10% FBS-containing media were seeded in
the well (5 × 10
3
cells per well) of a 96-well plate and incubated at 37 ℃ for 6 h. The cells were
then exposed to a series of concentrations of free drugs and drug-loaded cMLVs, at different
molar ratios of combined drugs for 48 h. Then 50 µL of XTT labeling mixture (Roche Applied
Science) was added. After incubation at 37 ℃ for 4 h, the absorbance of the solution was
measured at 570 nm using microplate reader (Molecular Devices) to determine the OD value.
The data were given as mean ± standard deviation (SD) based on 3 independent measurements.
The cell viability was calculated by subtracting absorbance values obtained from media-only
wells from drug-treated wells and then normalizing to the control cells without drugs (Zhang et
al. 2015, 2251-2256). The cell viability was calculated as follows: Cell viability=OD treated/OD
control×100%, where OD treated was obtained from the cells treated by a particular agent, and
OD control was obtained from the cells without any treatments. The fraction of cells affected (fa)
at each drug concentration was subsequently determined for each well. The data was analysed by
nonlinear regression to get the IC50 value. The median-effect method assesses the drug-drug
interaction by a term called the “combination index” (CI), which is based on the concentration-
response relationship. CI was calculated by the equation: CI=D1/Dm1+D2/Dm2,
where D1 and
29
D2 are the doses of drug 1 and drug 2 that in combination produce some specified effect (e.g., 50%
inhibition of cells) and Dm1 and Dm2 are the doses of the drugs at which the drugs have the
same effect when administered singly. The CI values lower than, equal to, and higher than 1
denote synergism, additivity, and antagonism, respectively.
2.3.6 Flow cytometry and fluorescence-activated cell sorting
FACS analysis was performed MACS Quant analyzer (Miltenyi Biotec Inc., San Diego, CA).
OVCAR8 and NCI/ADR- RES cells were seeded into 6-well plates at 6 × 10
6
cells per well.
When cells reached about 70% confluence, the cells were incubated with PBS control,
cMLVs(10µM CPT), cMLVs(10µM PTX), cMLVs(8µM CPT+2µM PTX), cMLVs(5µM
CPT+5µM PTX) and cMLVs(2µM CPT+8µM PTX) for 24 h. For ALDH staining, OVCAR8
and NCI/ADR-RES cells (3 × 10
5
) from the same cMLVs treatment conditions were harvested
and an aldefluor kit was used according to the manufacturer’s instructions (Stem Cell
Technologies, Vancouver, Canada). For analysis of autofluorescent subpopulations, OVCAR8
and NCI/RES-ADR cells (2 × 10
5
) from the same cMLVs treatment conditions were harvested
and detected Using SORP LSR II (BD Biosciences, San Jose, CA). Autofluorescent cells are
excited with a 488-nm blue laser and best selected as the intersection with filters 525/50 and
575/26. A proper distance between gates for autofluorescent and non-autofluorescent cells is
required to achieve high purity during sorting (Kim et al. 2015).
2.3.7 Evaluation of the acute toxicity
Six-week-old female BALB/c mice were randomized into 5 groups (n=3) based on body weight.
All the Mice were administered with PBS, CPT+PTX (10mg+20mg/kg and 15mg+30mg/kg) and
cMLVs (CPT+PTX) (10mg+20mg/kg and 15mg+30mg/kg) through intravenous injection once.
30
After the injection, the body weight and physical states of the mice were monitored every day.
On day 7 after injection, animals were euthanized via CO2 overdose, kidneys and livers were
harvested and fixed in 4% paraformaldehyde. Then livers and kidneys were frozen and cut into
sections and stained with hematoxylin and eosin (H&E) for pathology analysis.
2.3.8 In vivo antitumor study
Athymic mice were inoculated subcutaneously with 5 ×10
5
OVCAR-8 cells in logarithmic
growth phase from cell culture. Tumor volume was calculated according to the formula tumor
volume (in millimeters cubed) = D × d
2
/2, where D and d are the longest and shortest diameters,
respectively. The tumors were allowed to grow for 2 month to a volume of ∼100 mm
3
before
treatment. The mice were injected intravenously through the tail vein with PBS, cMLVs (1.8
mg/kg CPT), cMLVs (4 mg/kg PTX), cMLVs (1.8 mg/kg CPT + 4 mg/kg PTX) every 3 days for
four rounds (four mice per group). Tumor growth and body weight were monitored until the end
of the experiment. The length and width of the tumor masses were measured with a fine caliper
every 3 days after injection. 12 days after injection, tumors were harvested, fixed, frozen,
sectioned. Frozen sections were treated using the In Situ Cell Death Detection kit (Roche,
Indianapolis, Indiana) as recommended by the manufacturers. Fluorescence images were taken
using a Yokogawa confocal scanner system (Solamere Technology Group, Salt Lake City, UT)
with a Nikon Eclipse Ti-E microscope. Illumination powers at 405, 491, 561, and 640 nm solid-
state laser lines were provided by an AOTF (acousto-optical tunable filter)-controlled laser-
merge system with 50 mW for each laser.
31
2.3.9 Statistical analysis
Data are presented as means ± standard error (SEM). Statistical analysis for comparison of two
groups and multiple groups was performed by Student's t-test and one-way analysis of variance
(ANOVA) respectively.
2.4 Results and discussion
2.4.1 Drug encapsulation and release from cMLVs
As shown in Figure 2-1, the hydrophilic drug, carboplatin and hydrophobic drug, paclitaxel was
encapsulated into the aqueous core and lipid membranes of cMLVs respectively. As a result,
cMLVs could encapsulate binary therapeutic drugs with very different chemo-physical
properties and mechanisms of action into a single nanocarrier so as to synergistically enhance the
efficacy of treatment. Furthermore, CMLVS dispersion could be stabilized by PEG-coating,
which minimizes the interactions between the CMLVS and blood components.
Figure 2-1. Schematic Illustration of the Codelivery of Carboplatin and Paclitaxel via cMLVs.
32
Table 2-1. Drug encapsulation properties and mean diameter of cMLVs
Formulation
Encapsulation capacity (w/w %)
Diameter
(nm)
PDI
CPT PTX
cMLVs(CPT) 13.2±0.3 - 206±9.68 0.10
cMLVs(PTX) - 29.3±0.7 212±8.13 0.08
cMLVs(CPT+PTX) 12.8±0.2 28.7±0.5 225±10.97 0.11
As shown in Table 2-1, the single drug encapsulation capacity for CPT and PTX were
13.2 w/w% and 29.3 w/w% respectively. Interestingly, the binary drug-loaded cMLVs
showed similar encapsulation capacities compared with its single drug-loaded
formulation. This result could be explained by that different compartments of cMLVs
occupied by CPT and PTX. Therefore, we have confirmed that loading multiple drugs
into cMLVs has no effect on the encapsulation capacity of the individual drugs.
Moreover, Table 2-1 shows that there is no significant difference in size between single
drug-loaded cMLVs and binary drug-loaded cMLVs. As shown in Figure 2-2, both
paclitaxel and carboplatin displayed sustained release behaviors from cMLV.
Figure 2-2.Release behavior of Carboplatin and Paclitaxel from cMLVs.
33
2.4.2 In vitro cytotoxicity of drug-loaded cMLVs
The cytotoxicity of the free drugs and their cMLVs formulations was determined in human
ovarian carcinoma OVCAR8 cells using XTT assay. In addition, it has been reported that the
drug combination could have a positive effect on overcoming multi-drug resistance in cancer
cells (Villedieu et al. 2007, 373-384; Liu et al. 2014b). Hence, the effectiveness of each regimen
on inducing the apoptosis of NCI/ADR-RES has also been included by this study. All the
calculated IC50 values are summarized in Table 2-2.
Table 2-2.IC50 and CI values of free drug and cMLVs therapeutics against OVCAR8 and NCI/ADR-RES.
Formulation
CPT/PTX
( molar ratio)
OVCAR8 NCI/ADR-RES
IC 50 (µM) CI 50 IC 50(µM) CI 50
CPT 21.09 - 79.36 -
PTX 5.85 - 9.86 -
CPT+PTX 1.52/1.52 0.33 7.13/7.13 0.81
cMLVs(CPT) 16.54 - 18.88 -
cMLVs(PTX) 2.69 - 8.44 -
cMLVs(4CPT+PTX) 0.92/0.23 0.14 6.05/1.51 0.49
cMLVs(CPT+PTX) 0.29/0.29 0.13 1.65/1.65 0.28
cMLVs(CPT+4PTX) 0.16/0.63 0.24 0.93/3.73 0.50
As shown in Table 2-2 and Figure 2-2A-B, PTX (IC50, 5.85µM, 9.86 µM) is more potent at
inducing apoptosis than CPT (IC50, 21.09µM, 79.36 µM) against OVCAR8 and NCI/ADR-RES.
34
Importantly, both cMLVs(CPT) and cMLVs(PTX) exhibit slightly more toxicity against
OVCAR8 and NCI/ADR-RES cells than their corresponding free drugs.
The anticancer efficacy of the free drug combination, CPT+PTX and cMLVs(CPT+PTX) were
evaluated. Both CPT+PTX (Figure 2-3(A, C)) and cMLVs(CPT+PTX) (Figure 2-3(B, D))
exhibited an enhancement of combination potency. The IC50 of free CPT alone and free PTX
alone against OVCAR8 was 21.09 µM and 5.85µM respectively, while the IC50 of CPT (1.52
µM) and PTX (1.52 µM) in the free CPT+PTX combination are both reduced. Moreover, in
contrast to the IC50 of cMLVs (CPT)
(16.54 µM) and the cMLVs (PTX) (2.69 µM) against
OVCAR8, the IC50 of CPT and PTX in the cMLVs(CPT+PTX) formulation are reduced even
more significantly.
These data suggest synergistic cytotoxicity in free CPT+PTX combination against OVCAR8
cells. The synergistic effect is more evident when the component drugs are encapsulated into one
nanocarrier due to the nanocarrier's ability to ensure that both of the component drugs are
delivered into the cell at a desired ratio.(Liu et al. 2014, 1651-1661) A similar synergistic effect
was also observed for NCI/ADR-RES cells. Importantly, as shown in Figure 2-2, the CPT+PTX
combination delivery via cMLVs exhibits improved potency compared to the free CPT+PTX
combination against OVCAR8 and NCI/ADR-RES cells. The enhanced synergism of
cMLVs(CPT+PTX) is attributed to the cMLVs’ morphology since their rate of endocytosis is
often more efficient than the rate of passive diffusion for small molecular drugs (Rowinsky et al.
1991, 1692-1703).
35
Figure 2-3. In vitro cytotoxicity of cMLVs (CPT/PTX) and CPT/PTX against ovarian cancer cells. In
vitro cytotoxicity profiles of CPT/PTX and cMLVs (CPT/PTX) against OVCAR8 (A, B) and NCI/ADR-RES
(C, D) cells.
It has been widely reported that the dose ratio of the drug combination plays a very important
role in the therapy's combination effect, synergy, additivity, or antagonism (Mayer et al. 2006,
1854-1863). In order to determine the optimal ratio that can induce the strongest synergy, the
cytotoxicitiy of CPT and PTX combinations at three different molar ratios (CPT: PTX, 1:4, 1:1,
4:1) encapsulated by cMLVs were assessed in OVCAR8 and NCI/ADR-RES cells. As shown in
Figure 2-5, the cytotoxicity of cMLVs (CPT/PTX) with CPT/PTX ratios at 1:4 and 1:1 were
significantly stronger than that of the 4:1 molar ratio in both OVCAR8 and NCI/ADR-RES cells.
Table 2-2 displays the IC50 of the binary drugs-loaded cMLVs at the three different molar ratios.
36
Figure 2-4. In vitro cytotoxicity of cMLVs with various CPT/PTX ratios against ovarian cancer cells. In
vitro cytotoxicity profiles of cMLVs (CPT/PTX) with various CPT/PTX ratios against OVCAR8 (A) and
NCI/ADR-RES (B) cells.
To further study the synergy in cMLVs (CPT/PTX), we determined its combination index (CI).
CI values lower than, equal to, and higher than 1 indicate synergism, additivity, and antagonism,
respectively. As shown in Table 2-2, at 50% cell killing effect (CI50), synergistic effects were
observed in both OVCAR8 and NCI/ADR-RES cancer cells treated with cMLVs (CPT/PTX)
with three different molar ratios.
However, the strongest synergistic cytotoxicity was observed at a 1:1 molar ratio of CPT/PTX in
the co-loading cMLVs formulation showing the lowest CI50 values of 0.13 and 0.28 on OVCAR8
and NCI/ADR-RES cells respectively.
It has been reported that the prevalence of ovarian cancer stem cells (CSCs) associates with
recurrence in early-stage ovarian cancer (Whitworth et al. 2012, 226-230). In addition, aldehyde
dehydrogenase activity (ALDH+) is a known CSC biomarker for all human ovarian cancers (Liu
et al. 2013). This provides an approach to investigate the ability of drug-loaded cMLVs to
eliminate CSCs in OVCAR8 and NCI/ADR-RES cell lines.
37
Figure 2-5. In vitro cytotoxicity against CSCs. ALDH+ cells in unsorted OVCAR8 and NCI/ADR-RES cell
lines determined by FACS analyses using ALDH activity assay.
FACS analyses showed that untreated OVCAR8 and NCI/ADR-RES cell lines contained 37.6%
and 57.3% ALDH+ cells, respectively (Figure 2-6). Ovarian cancer cells treated by cMLVs(CPT)
reduced the population of ALDH+ cells significantly in OVCAR8 (13.2%) and NCI/ADR-RES
(24.7%), whereas, treatment of cMLVs(PTX) did not show a strong ability to alter ALDH+
population (32.4% and 44.8%, in OVCAR8 and NCI/ADR-RES cells respectively) (Figure 2-5).
The stronger cytotoxicity of CPT against CSCs could be explained by the different mechanisms
of action of PTX and CPT. PTX is an M-phase specific antitumor drug and acts on highly
proliferative cells. However, it has been reported that the cancer stem cells exhibit a quiescent
slow-cycling phenotype.(Zeuner et al. 2014, 1877-1888; Moore and Lyle 2011, 10.1155/2011/396076.
Epub 2010 Sep 29) Thus, PTX is not able to kill the cancer stem cells efficiently. This is also the
reason why cancer stem cells can induce resistance to conventional chemotherapies that are
dependent on cell cycle inhibition.(Kim et al. 2014, 132702) In contrast, cisplatin is a cell cycle
38
independent antitumor drug that covalently binds to DNA and leads to efficient elimination of
CSCs. Due to the distinct effects induced by CPT and PTX on eliminating the CSCs, the
combination treatments via cMLVs with different ratios of CPT/PTX also showed different
effectiveness at inhibiting the growth of ALDH+ populations in OVCAR8 and NCI/ADR-RES
cells. As shown in Figure 2-6, the ability of cMLVs (CPT/PTX) at molar ratios of CPT/PTX at
4:1 and 1:1 to eliminate ALDH+ cells were significantly stronger than that of cMLVs
(CPT/PTX) prepared at 1:4 molar ratio of CPT/PTX in both OVCAR8 and NCI/ADR-RES cells.
However, in vitro cytotoxicity assays showed that cMLVs (CPT/PTX) at CPT/PTX ratios of 1:4
and 1:1 displayed stronger cytotoxicity in OVCAR8 and NCI/ADR-RES cell lines. Taken
together, these result indicate that 1:1 is a desired molar ratio for cMLVs (CPT/PTX)
combination treatment to target the bulk and ALDH+ CSCs population. Hence, all the CPT and
PTX loaded cMLVs were prepared at 1:1 molar ratio of CPT/PTX in the following studies.
2.4.3 In vivo toxicity
On day 2, as shown in Figure 2-6, the groups treated with the free drug combinations at 20 and
30 mg/kg of PTX dosage experienced a loss of 13.2 and 15.5 % of their body weight respectively.
In contrast, the two cMLVs(CPT+PTX) administration groups showed a much smaller body
weight loss(2-4%). It was noted that all mice receiving free drug combination at all dose levels
showed convulsion over 2 minutes after injection. Whereas, no convulsion was observed with all
dose levels of nanotherapeutics. There was no difference in the general behavior between mice in
the cMLVs(CPT+PTX) groups and mice in the PBS group in the following days, but the mice
treated with free drug combinations were significantly less active.
39
Figure 2-6.In vivo toxicity (body weight change).The body weight changes of BALB/c mice treated with a
single dose of free drug mixture of CPT+PTX and cMLVs(CPT+PTX) at 20 and 30 mg PTX/kg levels in
comparison with PBS control group. Error bars represent standard error of the mean, n=3 for each treatment
group.
To determine whether cMLVs could reduce systemic toxicity, the acute toxicity of the
combination therapeutics was studied. According to previously published data, the maximum
tolerated dose of free PTX for cancer treatments was set at 20mg/kg or lower in mice models (Lu
et al. 2013, 1591-1600). Thus, we tested the efficacy of the free drug combination (CPT+PTX)
and nanotherapeutics cMLVs(CPT+PTX) with a PTX dosage of 20 mg/kg and 30 mg/kg - a 50%
increase. These treatments were injected into BALB/c mice intravenously through the tail vein.
The weights and overall behavior of the mice were monitored for 7 days after the single injection.
Light microscopic examination of H&E stained liver sections from sacrificed animals showed
that liver had no morphological changes after the treatment with cMLVs(CPT+PTX) compared
with control group, while obstruction of sinusoids in livers were observed from all animals
treated with free drug combination.
40
Figure 2-7. In vivo toxicity (Histopathological change).Histopathological changes in liver from the acute
toxicity studies in BALB/c mice. on day 7 after one intravenous treatment with PBS, free drug mixture of
CPT+PTX and cMLVs(CPT+PTX) at 20 and 30 mg PTX/kg levels in comparison with PBS control group.
2.4.4 In vivo anticancer efficacy of drug-loaded cMLVs
We evaluated the efficacy of our cMLVs(CPT+PTX) combination therapy using OVCAR8
xenograft nude mice. When tumors were about 100 mm
3
in size, the mice were randomly divided
into 4 groups and were given four injections of drug-loaded cMLVs intravenously on days 0, 3, 6,
and 9, with the day of the first injection as day 0. The tumor volume and body weight were then
monitored every three days for 12 days. The treatments for the 4 groups were as follows: (a) PBS
control, (b) cMLVs(CPT) (1.8 mg/kg), (c) cMLVs(PTX) (4.0 mg/kg) and (d)
cMLVs(CPT+PTX) (1.8 mg+4.0 mg/kg). No weight change was observed in all treatment
groups for the duration of the experiment, indicating the absence of systematic toxicity from the
cMLVs nanotherapeutics.
41
Figure 2-8.In vivo tumor growth inhibition. (A), and body weight changes (B) of mice bearing OVCAR8
ovarian cancer xenografts after intravenous treatment with PBS, cMLVs(CPT)(1.8mg/kg),
cMLVs(PTX)(4.0mg/kg)and cMLVs(CPT+PTX)(1.8mg/kg+4.0mg/kg) on days 0, 3, 6 and 9. Error bars
represent standard error of the mean, n=4 for each treatment group (**p < 0.01; ***p < 0.001).
Meanwhile, in terms of tumor inhibition, mild tumor inhibitory effects (p < 0.01) were exhibited
by cMLVs(CPT) (1.8 mg CPT/kg) and cMLVs(PTX) (4.0 mg PTX/kg) compared to the control
group. However, no difference in tumor growth inhibition (p=0.09) were seen between
cMLVs(CPT) (1.8 mg CPT/kg) and cMLVs(PTX) (4.0 mg PTX/kg). More importantly,
cMLVs(CPT+PTX) (1.8 mg CPT+4.0 mg PTX/kg) displayed stronger tumor inhibitory effects (p
< 0.01) than both cMLVs(CPT) (1.8 mg CPT/kg) and cMLVs(PTX) (4.0 mg PTX/kg). On day 12,
the relative tumor volume median for cMLVs(CPT+PTX) (1.8 mg CPT+4.0 mg PTX/kg) was
1.1. Comparatively, the relative tumor volume median for mice treated with cMLVs(CPT) (1.8
mg CPT/kg), cMLVs(PTX) (4.0 mg PTX/kg) and PBS were 2.0, 1.9 and 3.5, respectively. The
superior tumor inhibition of cMLVs(CPT+PTX) further confirms the synergistic effects between
CPT and PTX.
42
Figure 2-9.TUNEL staining of apoptotic positive cells in the OVCAR8 tumor. Mice beard OVCAR8
ovarian cancer xenografts were treated with PBS, cMLVs(CPT)(1.8mg/kg), cMLVs(PTX)(4.0mg/kg) and
cMLVs(CPT+PTX)(1.8mg/kg+4.0mg/kg) on days 0, 3, 6 and 9. On day 12, tumors were excised. Apoptotic
cells were detected by a TUNEL assay (green) and costained by nuclear staining DAPI (blue). The scale bar
represents 50 µm.
A TUNEL assay was then conducted to observe the number of apoptotic cells in OVCAR8
tumors treated with CPT and PTX and CPT+PTX in cMLV formulations for 4 rounds, in order
to further investigate whether the synergistic effects could also be induced by the in vivo co-
delivery of cMLVs. As shown in Figure 2-9, OVCAR8 tumors treated with cMLVs(CPT) and
cMLVs(PTX) induced a moderate amount of cell apoptosis compared to controls. Additionally,
cMLVs(CPT+PTX) promoted a more remarkable tumor cell apoptosis than both cMLVs(CPT)
43
and cMLVs(PTX). These results are consistent with results of the inhibitory effect on tumor
growth (Figure 2-8).
2.5 Discussion
CPT and PTX, which have been widely used as a combined platinum-taxane regimen in ovarian
cancer, to achieve synergistic antitumor activity. A number of clinical studies showed the limited
efficacy of this combination due to the distinct pharmacokinetics and noncoordinated
biodistribution profiles of this combination which caused by their different lipophilicities. In the
present study, we apply the CMLVs to enable the codelivery of CPT and PTX. Our in vitro
cytotoxicity result revealed the CMLVs could improve efficacy of the combination comparing to
it was released as free drugs, this may be due to the sustained release behaviors obtained from
cMLVs (Figure 2-2), which can prolong the exposure time of either CPT or PTX to the
OVCAR8 and NCI/ADR-RES cells.
It also has been revealed that the effects of antitumor drug combinations is determined by the
ratio of the combined drugs exposed to cells. Therefore, it is highly desirable to maintain a
synergistic ratio of combined drugs when the drug combination delivered to cancer cells. Here,
in vitro results showed that cMLVs allows us to coload CPT and PTX with predefined ratios and
induce a ratio-dependent synergy in cancer cells. This property of CMLVs could also enable
combination therapy to kill bulk cancer cells and CSCs simultaneously.
Furthermore, in vivo results revealed that the enhanced combinatorial efficacy of cMLVs and
attenuated systemic toxicity compared to cocktail combination is be attributed to its
macromolecular size and the augmented accumulation of drugs at tumor sties.
44
These results, taken together, demonstrated that the applicability of cMLVs in therapy is not only
attributed to the prolonged exposure of drugs to tumor cells, but also to codelivery of drug
combination at synergistic dose ratios to the site of action without inducing systemic toxicity.
2.6 Conclusion
In this study, we have utilized a cross-linked multilamellar liposome for the effective co-delivery
of CPT and PTX as an ovarian cancer combination therapy. These multi-compartment and cross-
linked nanostructures enable the highly encapsulation efficacy for both CPT and PTX and
prolonged stability upon systemic administration. When combined at the optimal drug-loading
ratio, the different mechanisms of action of CPT and PTX allow for synergistic, cytotoxic effects
against ovarian cancer cells. In vivo study showed that the binary drugs combination therapy
administrated via cMLVs displayed a safe and effective inhibition toward OVCAR8 tumor
growth. Overall, this CPT and PTX co-delivery system could offer its potential use in clinic for
ovarian cancer treatment.
45
CHAPTER 3. ANTI-PD-1 ANTIBODY ENHANCES ANTITUMOR IMMUNITY
OF CAR-MODIFIED T CELLS IN B-CELL LYMPHOMA
46
3.1 ABSTRACT
Despite favorable responses of chimeric antigen receptor (CAR)-engineered T cell therapy in
patients with certain hematologic malignancies, the outcome has been far from satisfactory in the
treatment of solid tumors or lymphoma, partially owing to the development of an
immunosuppressive tumor microenvironment. To overcome this limitation, we combined CAR-
T cells with checkpoint inhibitor (CPI) anti-PD-1 antibody and evaluated the antitumor efficacy
in a murine lymphoma model. In an effort to evaluate the effector function and expansion
capacity of CAR T cells combining with PD-1 blockade in vitro, we measured the production of
IFN-γ and cell proliferation marker Ki-67 following antigen-specific stimulation. Furthermore,
the antitumor efficacy of CAR-T cells, anti-PD-1 antibody and CAR-T cells combined with anti-
PD-1 antibody was determined using a murine lymphoma model. Finally, the underlying
mechanism of enhanced tumor eradication of combination therapy was investigated by analyzing
the expansion and tumor infiltration of adoptively transferred T cells. We demonstrated that PD-
1 blockade significantly enhanced CAR T cell expansion and effector function in vitro. In the
animal study, we demonstrated that the anti-PD-1 antibody enhanced the antitumor activity of
CAR-T cells and prolonged overall survival. In conclusion, our study supports the
immunotherapeutic approaches that combine immune checkpoint blockade with CAR-T cell
therapy to achieve better antitumor immunity, especially in the treatment of tumors with
suppressive microenvironment.
47
3.2 INTRODUCTION
The development of ex vivo culture technology and genetically engineered T cells has led to
rapid generation of chimeric antigen receptor (CAR) T cells, thereby broadening the applicability
of cancer immunotherapy(Vera et al. 2010, 305-315; Hollyman et al. 2009, 169-180; Somerville
et al. 2012, 1). Typically, a CAR, consisting of an extracellular antigen-recognition domain, a
hinge, a transmembrane domain, and an intracellular signaling domain, is genetically engineered
on the T cell to enable tumor-associated antigen (TAA) recognition in a major histocompatibility
class (MHC)- independent manner(Ahmed et al. 2015, 1688-1696).
So far, CAR-T cell therapy has consistently demonstrated significant antitumor capacity in
patients with acute lymphoblastic leukemia (ALL) (Maude et al. 2015, 4017). It shows that up to
90% of children and adults with ALL, who had either relapsed or failed to respond to standard
therapies, achieved a complete remission after CAR-T cell therapy (Maude et al. 2014, 1507-
1517). However, the outcome has been less inspiring in treatment of other hematological tumors
such as lymphoma or solid malignancies. For example, in recent clinical trials of non-Hodgkin
lymphoma (NHL), only 33% complete remission was achieved with CAR-T cell therapy (Locke
et al. 2015, 3991). This can be attributed, in part, to the establishment of an immunosuppressive
microenvironment within the tumors. Such milieu involves the upregulation of a number of
intrinsic inhibitory pathways mediated by increased expression of inhibitory receptors (IRs) in T
cells reacting with their cognate ligands expressed on cancer cell surface (Pardoll 2012, 252-
264).
Programmed dearh-1 (PD-1) is one of the most important IRs characterized in T cells. Unlike
other IRs, PD-1 is upregulated shortly after T cell activation, which in turn, inhibits T cell
48
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) (Kondo et al. 2010, 1124).
Additionally, PD-L1 is also shown to be expressed abundantly in a wide variety of solid tumors
(Dong et al. 2002b, 793-800; Brown et al. 2003, 1257; Konishi et al. 2004, 5094) and
lymphomas especially classical Hodgkin lymphoma (CHL) and aggressive virus-associated B-
cell lymphomas (Gatalica et al. 2015, 3899). In contrast, the expression of PD-L1 in normal
tissues is undetectable (Dong et al. 2002a, 793-800). As a consequence of its critical role in
immunosuppression, PD-1 has been the focus of recent research. It aims to neutralize PD-1’s
negative effect on T cells and enhance their antitumor responses. Clinical studies have
demonstrated that PD-1 blockade significantly enhanced tumor regression in colon, renal and
lung cancers, melanoma, CHL and diffuse large-B-cell lymphoma (DLBCL) (Dong et al. 2002b,
793-800; Goodman, Patel, and Kurzrock 2017, 203-220).
A recent study shows tumor-induced hypofunction of CAR T cells as well as upregulation of
PD-1 on the CAR T cells and demonstrates the contribution of PD-1 to the dysfunction of tumor-
infiltrating CAR T cells (Moon et al. 2014, 4262), thereby suggesting a potential strategy
whereby CAR T therapy could be combined with PD-1 blockade in cancer treatment (Chong et
al. 2017, 1039). Herein, in this study, in order to expand the application of CAR T cells in cancer
immunotherapy, we combined PD-1 blockade and CAR T cells and examined the antitumor
activity of the combined therapy in the treatment of lymphoma by using A20 murine B-cell
subcutaneous lymphoma model. We demonstrated that PD-1 blockade significantly improved the
antitumor capacity and enhanced the antitumor immunity of CAR-T cell therapy.
49
3.3 MATERIALS AND METHODS
3.3.1 Plasmid
The MSGV-1D3-28Z.1-3 recombinant retroviral vector was kindly provided as a gift from Dr.
Steven A. Rosenberg. It encodes the MSGV (mouse stem cell virus-based splice-gag vector)
retroviral backbone and the 1D3-28Z.1-3 CAR. The 1D3-28Z.1-3 CAR consists of an anti-CD19
scFv, a portion of the murine CD28 molecule, and the intracellular component of the murine
TCR-ζ molecule in which the first and third ITAMs were inactivated.
3.3.2 Mice
BALB/c mice and the congenic strain CByJ.SJL (B6) (The Jackson Laboratories) were used for
in vivo efficacy studies. Mice were fed ad libitum and kept in air-conditioned rooms at 20°C ±
2°C with a 12-hour light–dark period. Animal care and manipulation were in conformity with
USC institutional guidelines, which were in accordance with the Guidelines for the Care and Use
of Laboratory Animals.
3.3.3 Cell culture
293T cells were maintained in a 5% CO2 environment with Dulbecco’s modified Eagle’s
medium supplemented with 10% FBS, 2 mM L-glutamine, 100 U/ml penicillin and 100μg/ml
streptomycin. A20 cells (ATCC number: TIB-208
TM
) was maintained in 5% CO2 environment
with RPMI-1640 medium supplemented with 10% FBS, 2 mM of L-glutamine, 100 U/ml
50
penicillin, 100μg/ml streptomycin and 0.05mM β-mercaptoethanol (Sigma-Aldrich, St. Louis,
MO). All cell culture media and additives were purchased from Hyclone.
3.3.4 Retroviral T-cell Transduction
A total of 8-10 × 10
7
splenocytes were harvested from CByJ.SJL (B6) mouse and activated in
vitro using the T-cell activation Kit from e-Bioscience, and supplemented with 15ng/ml mIL-2.
Two days later, anti-mouse-CD19-CAR-encoding retroviral vector was harvested, and added to
non-tissue culture plates that had been coated overnight with 15 μg/mL retronectin (Takara Bio
Inc) at 4°C. Retroviral vector was spin-loaded onto plates by centrifuging 2 hours at 2000g at
32°C. The activated T cells were washed and resuspended at 10
6
/mL in R10, and 10
6
activated T
cell were added per well to the retrovirus-loaded plates. Plates were spun at 1000g at 32°C for 10
minutes and incubated overnight at 37°C, 5% CO2. The next day (day 3), the transduced T cells
were transferred to a new 24-well plate at 10
6
/mL in R10. Before treatment, CD19 CAR-
transduced mouse T cell were evaluated for expression of the appropriate CD19 CAR by Anti-
Fab staining and flow cytometric analysis, and cell function was evaluated by overnight co-
culture with cognate antigen-bearing target cells (5 × 10
5
:0.5 × 10
5
) and intracellular staining of
interferon-γ (IFN-γ) was performed subsequently. T cells for treatment were washed in saline
before infusion into BALB/c mice intravenously.
3.3.5 Anti-CD19 CAR staining
To detect anti-CD19 CAR expression on the cell surface, cells were stained with goat anti-mouse
IgG (H+L) conjugated with FITC (Jackson ImmunoResearch). Before FACs staining, 5 × 10
5
were harvested and washed three times with FACs buffer (PBS containing 5% bovine serum
albumin fraction V). Cell were then stained with 20 μg of goat anti-mouse IgG (H+L) conjugated
51
with FITC at 4°C for 30 minutes. Cells were washed and then fixed with transfix cellular antigen
stabilizing reagent (Thermo Scientific, Waltham, MA) at 4°C for 10 minutes. Cells were then
washed twice and stained with PE-anti-CD3, FITC-anti-CD4, PE/Cy5-anti-CD8 (BioLegend,
San Diego, CA) at 4°C for 10 minutes. Cells were washed and resuspended in PBS.
Fluorescence was assessed using a MACSquant cytometer (Miltenyi Biotec, San Diego, CA),
and all the FACs data were analyzed using FlowJo software (Tree Star, Ashland, OR).
3.3.6 Surface immunostaining and flow cytometry
Splenic T cells (1 × 10
6
)
were cultured with target cells for overnight at 37°C and 5% CO2 in 96-
well round bottom plates. PE-Cy5.5-anti-CD3, Pacific Blue
TM
-anti-CD8, PE-anti-PD-1, FITC-
anti-TIM-3 and APC-anti-LAG-3 antibodies were used for the cell surface staining.
3.3.7 Intracellular staining
Splenic T cells (1 × 10
6
)
were cultured with target cells for 16 hours at 37°C and 5% CO2 with
(IFN-γ) or without (Ki-67 and p-Akt) GolgiPlug (BD Biosciences, San Jose, CA) in 96-well
round bottom plates. PE-Cy5.5-anti-CD3, FITC-anti-CD4, Pacific blue-CD8, PE-anti-IFN-γ, PE-
anti-Ki-67 and APC-anti-p-Akt antibodies were used for the intracellular staining.
Cytofix/Cytoperm Fixation and Permeabilization Kit (BD Biosciences) was used to permeabilize
the cell membrane and perform intracellular staining according to the manufacturer’s instruction.
3.3.8 Specific cell lysis assay
A20 cells were labeled by suspending them in PBS+0.1% BSA with 5 μM Carboxyfluorescein
succinimidyl ester (CFSE) fluorescent dye at a concentration of 1 × 10
6
cells/mL. The cells were
incubated for 30 minutes at 37°C. After incubation, the same volume of FBS was added into the
52
cell suspension and then incubated for 2 minutes at room temperature. The cells were then
washed twice and suspended in fresh R10 medium. Cocultures were set up in round bottom 96-
well plates in triplicate at the following effector-to-target ratios: 1:1, 5:1 and 10:1. The cultures
were incubated for 6 hours at 37°C, followed by 7-AAD labeling, according to the
manufacturer’s instructions (BD Biosciences). Flow cytometric analysis was performed to
quantify the percent survival (7-AAD-negative) of target cells. In the wells containing only target
cells without effector cells, the percentage of viable A20 cells was calculated and used to correct
the variation in the starting cell numbers and spontaneous cell death. The cytotoxicity was
determined in triplicate and presented in mean ± SEM.
3.3.9 ELISA
IFN-γ was measured using mouse IFN-γ ELISA kit (BD Biosciences, San Jose, CA) according to
the manufacturer’s instructions respectively. Briefly, 96-well ELISA plates (Thermo Scientific,
Waltham, MA) were coated with 200 ng/well of capture antibodies against the indicated proteins
at 4°C overnight. On the next day, plates were washed with wash buffer (PBS containing 0.05%
Tween 20) and blocked with assay buffer (PBS containing 10% FBS) for 2 hours at room
temperature. Equal volume of serum or protein extract was added to the plate and incubated for 2
hours at room temperature. Plates were then washed and incubated with detection antibodies for
1 hour at room temperature.
3.3.10 In vivo study
BALB/c mice were preconditioned (5 Gy) and then injected with 10 million A20 cell on the right
flank on Day 0. On the day 6, the mice were injected with scFv-anti-CD19-transduced T cells or
non-transduced T cells (6×10
6
/dose) from congenic CByJ.SJL (B6) mice on days 6 in
53
combination with either anti-PD-1 or isotype antibodies (250 µg/injection on days 6, 9, and 13).
Control groups were left untreated. Tumor growth in mice was monitored every 2 to 4 days.
Tumor size was measured by calipers and calculated by the following formula: W
2
× L / 2. Mice
were euthanized when they displayed obvious weight loss, ulceration of tumors, or tumor size
larger than 1000 mm
3
.
3.3.11 Statistical analysis
Statistical analysis was performed in GraphPad Prism (GraphPad Software, San Diego, CA,
USA), version 5.01. One-way ANOVA with Tukey’s multiple comparison was performed to
assess the differences among different groups in the in vitro assays. Tumor growth curve was
analyzed using one-way ANOVA with repeated measures (Tukey’s multiple comparison
method). Mouse survival curve was evaluated by the Kaplan-Meier analysis (log-rank test with
Bonferroni correction). A P value less than 0.05 was considered statistically significant.
Significance of findings was defined as: ns = not significant, P > 0.05; *, P < 0.05; **, P < 0.01;
***, P < 0.001.
3.4 RESULTS
3.4.1 Expression and activation of 1D3-28Z.1-3 anti-CD19 CAR in mouse splenic T cells
The schematic design of the 1D3-28Z.1-3 anti-CD19 CAR used in this study is shown in Figure
3-1A and has also been reported by the others (Kochenderfer et al. 2010, 3875-3886). To
examine the expression of 1D3-28Z.1-3 in primary lymphocytes, mouse splenic T cells were
transduced by retrovirus expressing 1D3-28Z.1-3 CAR. As shown in Figure 3-1B, reproducible
54
levels of 1D3-28Z.1-3 were observed in transduced primary splenic T cells. The median
percentage of CAR-expressing T cells was 76%.
Figure 3-1. Expression and activation of 1D3-28Z.1-3 anti-CD19 CAR in mouse splenic T cells. (A)
Schematic of retroviral construct encoding 1D3-28Z.1-3 anti-CD19 CAR. scFv, single chain variable fragment;
CD28, a portion of mouse CD28 molecule; CD3, the intracellular component of the mouse TCR-ζ molecule;
LTR, long terminal repeat. (B) CAR expression was detected in murine splenic T cells by flow cytometry
following FITC-conjugated goat anti-mouse IgG (H+L) staining. A viable CD3
+
lymphocyte gating strategy was
used. CD8
+
T cells were shown in each panel. CAR-expressing CD8 T cells were gated and shown in each
scatterplot. NT indicates non-transduced T cells, which were used as a control. (C) Intracellular expression of p-
Akt in T cells. p-Akt-expressing CD8 T cells were gated and shown in each scatterplot. (D) The summarized
statistics of (C) were shown in bar graphs (n=3, mean ± SEM; **P < 0.01).
Co-stimulation through either CD28 or 4-1BB has been shown to regulate multiple aspects of T
cell function by activating PI3K/Akt signaling pathway (Stärck et al. 2005, 1257-1266; Parry et al.
55
1997, 2495-2501). To evaluate the activation of downstream signaling pathway of CAR19 T cells
upon antigen-specific stimulation, non-transduced or CAR19 T cells were co-cultured with
CD19-expressing murine A20 B-cell lymphoma cells without exogenous IL-2 for 16 h before
they were subjected to intracellular staining for the expression of phosphorylated Akt. We found
that p-Akt (ser473) in CAR19 CD8
+
T cells was significantly increased upon antigen-specific
stimulation, compared to non-transduced CD8
+
T cells (15.9±3.0% versus 7.6±2.1%; Figure 3-
1C and 3-1D).
3.4.2 PD-1 expression is upregulated on anti-CD19-CAR-transduced T cells following
antigen-specific stimulation
It has been shown that PD-1 expression on T cells is increased following T-cell activation via
TCR recognition of MHC/peptide (Day et al. 2006, 350-354; Petrovas et al. 2006, 2281-2292;
Trautmann et al. 2006, 1198-1202). In addition, the expression of PD-1 was also shown to be
upregulated on anti-HER2 CAR T cells with antigen-specific stimulation (John et al. 2013a,
5636-5646). To evaluate if antigen-specific stimulation of T cells would have similar effect and
enhance the PD-1 expression on anti-CD19 CAR T cells, both the non-transduced and CAR19 T
cells were cultured with or without A20 cells for overnight. We found that without antigen
stimulation, the cell surface expression of PD-1 on CAR19 T cells is similar to that on the non-
transduced T cells. However, upon CAR stimulation, the expression of PD-1 on CAR19 CD8
+
T
cells was increased and significantly higher when compared to the non-transduced T cells
(15.6±1.9% versus 5.2±0.25%; Figure 3-2A).
In addition to PD-1, other cell surface inhibitory molecules, including lymphocyte activation
gene 3 protein (LAG-3), T cell immunoglobulin domain and mucin domain-containing protein 3
56
(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, 252-264). In order to evaluate whether the expression of other
T cell exhaustion markers is regulated by CAR stimulation, we measured the expression of
LAG-3 and TIM-3 in CAR19 T cells. We found that compared to non-transduced T cells,
without antigen stimulation, the basal level of LAG-3 on CAR19 CD8
+
T cells was significantly
higher (8.54±0.5% versus 4.59±1.1%; Figure 3-2B). With antigen-specific stimulation, the
expression of LAG-3 on both T cells was upregulated and the expression between non-
transduced and CAR19 T cells showed little difference (Figure 3-2B). The expression of TIM-3
on both non-transduced and CAR19 T cells was undetectable (data not shown).
Figure 3-2. Upregulation of PD-1 expression on CAR T cells following antigen-specific stimulation. Non-
transduced T cell or CAR19 T cells were stimulated with or without A20 B-cell lymphoma cells at an E/T ratio
of 1:1 for overnight. The cell surface expression of (A) PD-1 and (B) LAG-3 was detected by flow cytometry
following cell surface staining. The percentage of PD-1 (A) or LAG-3 (B) expressing CD8 T cells over total CD8
+
T cells was shown in bar graphs (n=3; ns, not significant, P > 0.05; **P < 0.01).
3.4.3 PD-1 blockade enhances the antigen-specific immune responses of 1D3-28Z.1-3 CAR
T cells
57
We have shown that PD-1, the expression of which plays an important role in inhibiting T cell
activation and inducing T cell exhaustion, was significantly upregulated on CAR19 T cells
following antigen-specific stimulation. Next, to assess the effect of PD-1 blockade on the
functional capacity of CAR19 T cells, the T cells that transduced with 1D3-28Z.1-3 were co-
cultured with A20 cells in combination with either anti-PD-1 or control isotype antibodies and
the expression of T cell function marker IFN-γ and cell proliferation marker Ki-67 was
evaluated. Notably, it revealed that PD-1 blockade significantly increased the expression of IFN-
γ and Ki-67 on CAR19 T cells, compared to the group that treated with control isotype antibody
(Figure 3-3A and 3-3B).
Figure 3-3. Anti-PD-1 enhanced the antigen-specific immune responses of CAR T cells. To examine the
effects of PD-1 blockade on T-cell function, non-transduced T cells or CAR19 T cells were co-cultured with
A20 cells at an E/T ratio of 10:1 (IFN-γ) or 1:1 (Ki-67) for overnight in the presence of anti-PD-1 or isotype
control antibodies. The T cells were then harvested and stained intracellularly for IFN-γ (A) and Ki-67 (B) as
58
determined by flow cytometry. (C) The ability of CAR19 T cells to lyse A20 cells with or without the presence
of anti-PD-1 antibody was determined by flow cytometry after 6-hour co-culture at 1:1, 5:1, and 10:1 effector-
to-target ratios. Non-transduced (NT) T cells were used as a control. Error bars represent standard error of the
mean for each treatment group (n = 3; *P < 0.05; **P < 0.01).
Furthermore, the cytolytic activity of CAR19 T cells with or without PD-1 blockade was
examined in a 6-hour cytotoxicity assay. The cytotoxic activity of T cells transduced with 1D3-
28Z.1-3 against A20 cells was evaluated at E/T ratios of 1, 5 and 10. We found that CAR19 T
cells mediated significant cell lysis of target cells at E/T ratios of 5 and 10, but not at E/T ratio of
1, compared to the non-transduced T cells. Interestingly enough, with PD-1 blockade, CAR19 T
cells induced significant cytotoxicity of target cells at E/T ratio of 1, compared to that without
PD-1 blockade (Figure 3-3D).
3.4.4 PD-1 blockade enhances 1D3-28Z.1-3 CAR T cell-mediated tumor regression in
established tumor model
Next, to evaluate the antitumor efficacy of combined PD-1 blockade with anti-CD19 CAR T
cells, we adoptively transferred 6 x 10
6
CAR19 T cells into BALB/C mice bearing established
A20 subcutaneous B-cell lymphoma (~30 mm
3
). 6-hour later on the same day, we intravenously
injected anti-PD-1 or control isotype antibodies. The antibody injection was continued twice a
week for two weeks. The experimental design for animal study is shown in Figure 3-4A (upper
panel). We found that anti-PD-1 antibody or CAR19 T cells treatment alone significantly
inhibited tumor growth, compared to the control group. Notably, combined PD-1 blockade and
CAR19 T cells further enhanced the antitumor efficacy, compared to either treatment given alone
(Figure 3-4A, lower panel). Moreover, the combination therapy significantly enhanced the long-
59
term survival, compared to monotherapy (Figure 3-4B).
Figure 3-4. PD-1 blockade enhanced CAR19 T cells mediated tumor regression of established tumor. (A)
Adoptive transfer of CAR19 T cells in combination with PD-1 blockade enhances growth inhibition of
established tumor. BALB/c mice were preconditioned (5 Gy) and then injected with 10 million A20 cell on the
right flank on Day 0. On the day 6, the mice were injected with CAR19 T cells or non-transduced T cells
(6×10
6
/dose) from congenic from congenic CByJ.SJL (B6) mice on days 6 in combination with either anti-PD-1 or
isotype antibodies (250 µg/injection on days 6, 9, and 13). Control groups were left untreated. Tumor growth in
mice was monitored every 2 to 4 days. Tumor size was measured by calipers and calculated by the following
formula: W
2
× L / 2. Mice were euthanized when they displayed obvious weight loss, ulceration of tumors, or tumor
size larger than 1000 mm
3
. Tumor growth curve was shown for mice without treatment, treated with non-transduced
(NT) plus anti-PD-1 injection, CAR19 plus isotype control antibody, or CAR19 plus anti-PD-1 antibody injection.
Data were presented as mean tumor volume ± standard error of the mean (SEM) at indicated time points (n = 6;
*P<0.05). (B) Survival of A20 tumor-bearing mice after indicated treatment. Overall survival curves were plotted
60
using the Kaplan-Meier method and compared using the log-rank (Mantel-Cox) test (n ≥ 7). The percentage of
transferred CD45.1
+
T cells in the tumor, blood, and spleen of A20 tumor-bearing mice that were adoptively
transferred with non-transduced (NT) plus anti-PD-1 injection, CAR19 plus isotype control antibody, or CAR19
plus anti-PD-1 antibody injection was investigated by flow cytometry on day 3 (C) or day 9 (D) post-therapy (n = 3,
mean ± SEM; *P<0.05; **P < 0.01; ***P < 0.001).
Further analysis of the engraftment and expansion of CAR T cells were assessed. Three days
following T cell infusion, mice were euthanized, and different organs and tissues, including the
tumor, blood, and spleen, were harvested for transferred-T cell staining. We found that certain
percentage of CD45.1
+
T cells (2-5%) circulated in the blood, whereas most of the CD45.1
+
T
cells (15-20%) homed to the spleen in all groups. Particularly, compared to CAR19 T cells
monotherapy, the combination therapy resulted in significantly higher infiltration of transferred
T cells into the spleen. The infiltration of CD45.1
+
T cell in the tumor was low and showed little
difference across all the examined tissues (Figure 3-4C). One-week post-T cell infusion, on day
9, we observed a significant expansion of transferred CD45.1
+
T cells in the circulation of mouse
treated with CAR19 T cells or combined CAR19 T cells and anti-PD-1 antibody. Remarkably,
compared to either treatment given alone, in the combined treatment group, CD45.1
+
T cells
specifically accumulated and expanded in the tumor, whereas the population of CD45.1
+
T cells
had little variation across all the treatment groups in the blood and spleen (Figure 3-4D).
3.4.5 The antitumor efficacy of combined therapy is associated with tumor
microenvironment modulation
It has been well established that both cytotoxic CD8
+
T cells and helper CD4
+
T cells were
involved in antitumor immunity (Fridman et al. 2012, 298-306). To further understand the
underlying mechanism of enhanced antitumor activity of combination therapy, we therefore
61
assessed the effects of combined therapy on tumor infiltrating lymphocytes. We found that in the
combined treatment group, PD-1 blockade significantly enhanced the accumulation of CD8
+
T
and CD4
+
T cells in the tumor microenvironment, in comparison with the control isotype
antibodies treatment (Figure 3-5A and 3-5B).
Figure 3-5. The enhanced antitumor efficacy of combined therapy is correlated with tumor
microenvironment modulation. Tumors were taken on day 13 post-therapy from mice treated with non-
transduced (NT) T cells plus anti-PD-1 antibody, CAR19 T cells plus isotype control antibody, or CAR19 T
cells plus anti-PD-1 antibody. Non-treated mice were used as a control. The percentage of CD8
+
T (A) and
CD4
+
T (B) cells at the tumor sites was examined by flow cytometry. Error bars represent standard error of the
mean for each treatment group (n=3; ns, not significant, P > 0.05; **P < 0.01).
3.4.6 Combination therapy specifically regulates local immune responses within tumor
To further substantiate the observed changes of tumor microenvironment, we examined the
expression of T-cell activation marker in tumor-bearing mice by ELISA. It showed that CAR19
T cells monotherapy significantly increased the expression of IFN-γ in the tumor, compared to
the control group. Moreover, the PD-1 blockade further enhanced CAR19 T cell-mediated IFN-γ
up-regulation (Figure 3-6A). However, the measurement of IFN-γ expression in the serum and
62
spleen showed that the combined therapy had little effects on IFN-γ expression, compared to
CAR19 T cells monotherapy (Figure 3-6B and 3-6C).
Figure 3-6. Combined therapy specifically regulates immune responses at local tumor site. (A) Tumors,
(B) Serum and (C) Spleens were harvested on day 9 post-therapy from mice treated with non-transduced (NT)
T cells plus anti-PD-1 antibody, CAR19 T cells plus isotype control antibody, or CAR19 T cells plus anti-PD-
1 antibody. Non-treated mice were used as a control. The expression of IFN-γ was quantified using ELISA.
Error bars represent standard error of the mean for each treatment group (n=3; ns, not significant, P > 0.05; *P <
0.05).
3.5 DISCUSSION
Adoptive T cell therapy using gene-modified T cells has emerged as a promising method for
cancer immunotherapy. It has achieved successful responses in patients with certain
hematopoietic malignancies. However, in the treatment of other blood-borne tumors, such as
63
follicular lymphoma (FL) and DLBCL, or solid tumors, the outcome has been less promising,
partly owing to the immunosuppressive properties and establishment of an immunosuppressive
microenvironment within the tumors (Vazquez-Cintron, Monu, and Frey 2010, 7133). The PD-
1/PD-L1 regulatory pathway has demonstrated a particularly antagonistic effect on the antitumor
response of tumor-infiltrated lymphocytes (TILs). For example, the FL TILs were shown to have
high PD-1 expression and PD-L1
+
histiocytes were also found within the T cell-rich zone of the
neoplastic follicles in FL (Myklebust et al. 2013, 1367). In addition to FL, the other lymphomas
or solid tumors with poor prognosis also showed upregulation of PD-L1 expression, with
increasing PD-1 expression on TILs (Myklebust et al. 2013, 1367; Myklebust et al. 2013, 1367;
Xia, Jeffrey Medeiros, and Young 2016, 58-71). The combined effect of these two leads to tumor
escape. However, this can be disrupted by the use of checkpoint inhibitors (CPIs), targeting the
PD-1/PD-L1 pathway (Dong et al. 2002a, 793-800; Ansell et al. 2015, 311-319; Brahmer et al.
2012b, 2455-2465). As a result, in this study, we combined anti-PD-1 antibody and CAR19 T
cells to evaluate the effects of PD-1 blockade in infused CAR19 T cells and CAR19 T cell-
mediated antitumor immunity.
Herein we have demonstrated that the expression of PD-1 was significantly increased on CAR19
T cells following antigen-specific stimulation and that PD-1 blockade significantly increased the
functional capacity of CAR19 T cells both in vitro and in vivo. In the adoptive T cell transfer
experiment, we demonstrated that combined PD-1 blockade and CAR19 T cells significantly
enhanced the CAR T cell-mediated inhibition of established tumor growth. Interestingly, we
found that the enhanced antitumor efficacy of combined therapy was correlated with increased
local accumulation and proliferation of adoptive transferred T cell and elevated modulation of
tumor microenvironment and immune responses within tumor.
64
It has been shown that the expression of PD-1 is increased shortly after T-cell activation through
TCR recognition of MHC/peptide (Day et al. 2006, 350-354; Petrovas et al. 2006, 2281-2292;
Trautmann et al. 2006, 1198-1202). In order to investigate the possibility that PD-1 can be
similarly upregulated on CAR19 T cells following antigen-specific stimulation, we co-cultured
the T cells with target cells for overnight and measured PD-1 expression. In consistent with
previous observation, which showed that the expression of PD-1 in anti-Her2 CAR T cells is
elevated upon antigen-specific activation (John et al. 2013b, 5636-5646), we demonstrated that
CAR stimulation significantly enhanced PD-1 expression on anti-CD19-CAR-transduced CD8
+
T cells. This observation suggests that the up-regulation of PD-1 may contribute to limiting the
full potential of CAR T cells in inducing anti-tumor immunity. It has been shown that co-
expression of multiple inhibitory receptors is a cardinal feature of T cell exhaustion (Wherry and
Kurachi 2015, 486-499). Thus, in addition to PD-1, other cell surface inhibitory molecules were
also examined. We found that the expression level of TIM-3 on both rested T cells and activated
T cells was minimal and undetectable. Interestingly, we observed that without stimulation, the
basal level of LAG-3 on CAR19 T cells was significantly higher than that on non-transduced T
cells. Upon antigen stimulation, LAG-3 expression was upregulated on both T cells, whereas
little difference was seen between non-transduced and CAR19 T cells T cells, suggesting that the
upregulation of LAG-3 may also contribute to limiting the activity of CAR T cells as PD-1 does,
but through different mechanisms (Buchbinder and Desai 2016, 98-106).
The PD-1/PD-L1 pathway involves the regulation of cytokine production by T cells, inhibiting
production of IFN-γ, TNF-α and IL-2 (Taylor, Lee, and Schiemann 2011, 117-132). In this
study, to unleash the inhibitory effect of tumor-induced upregulation of PD-1 on CAR T cells,
we combined anti-PD-1 antibody with CAR19 T cells, and demonstrated that PD-1 blockade
65
significantly enhanced T cell functional capacity by increasing the production of IFN-γ. In
addition to cytokine production, PD-1 can also inhibit T cell proliferation (Keir et al. 2008b, 677-
704). With CAR-specific stimulation in the presence of PD-L1
+
cancer cells, we found that anti-
PD-1 antibody significantly increased the expression of Ki-67 in CAR19 T cells, implying that
PD-1 blockade confers the CAR19 T cells to be more proliferated. Taken together, these data
imply that PD-1/PD-L1 signaling blockade results in more functional CAR19 T cells with higher
proliferation capacity.
Our in vivo study showed that the tumor growth could be inhibited by CAR19 T cells treatment
or anti-PD-1 antibody treatment given alone. However, in comparison, combined PD-1 blockade
and CAR19 T cells further enhanced the inhibitory effect on tumor growth and pro-longed the
survival. To understand the underlying mechanism of enhanced antitumor efficacy of the
combined therapy, we analyzed the expansion and infiltration of adoptively transferred T cells in
different tissues. We found that PD-1 blockade specifically increased the accumulation of
adoptively transferred T cells to local tumor tissues. However, in the other tissues, such as blood
and spleen, the accumulation of T cell is similar in groups with and without PD-1 blockade.
From our current study, it is unclear whether the administrated anti-PD-1 antibody enhances
CAR T cells infiltration, enhances T cell retention, or enhances T cell proliferation in the tumor
microenvironment, but also the effect could be a result of a combination of these effects. A
previous study by Wong et al. (2007) reported that PD-1 blockade augments proliferation (Wong
et al. 2007, 1223-1234). This data corresponds with our in vitro experiment that PD-1 blockade
enhanced cell proliferation of CAR19 T cells. Moreover, it has been reported that PD-1/PD-L1
blockade reduces T cell motility (Zinselmeyer et al. 2013, 757), which may lead to higher T cell
retention in the tumor microenvironment. In addition to adoptively transferred T cells, we also
66
examined the endogenous tumor infiltrating T cells and found that PD-1 blockade significantly
increased the population of tumor infiltrating CD8
+
T and CD4
+
T cells. The population of
cytotoxic CD8
+
T and helper CD4
+
T cells among TILs is critical in eliciting antitumor immunity
(Ceeraz, Nowak, and Noelle 2013, 10.1016/j.it.2013.07.003). Taken together, the increased
CAR19 T cells trafficking to tumor and the modulation of tumor microenvironment may both
contribute to the enhanced antitumor immunity of the combined therapy, which was also
supported by the specifically increased expression of IFN-γ in the tumor.
In conclusion, our study supports the immunotherapeutic approaches that combine immune
checkpoint blockade with CAR-T cell therapy. Tumor-induced upregulation of PD-1 on the
surface of CAR T cells and the low tumor infiltration of cytotoxic T cells lead to tumor evasion
to CAR-T cell therapy. In this study, we have demonstrated that PD-1 blockade allows enhanced
CAR T cells functional capacity, increased accumulation of CAR T cells, and infiltration of
endogenous cytotoxic CD8
+
and helper CD4
+
T cell into local tumor tissue, thereby leading to
enhanced antitumor immunity of CAR-T cell therapy.
67
CHAPTER 4. TARGETED DEPLETION OF TUMOR STROMAL CELLS CAN
ENHANCE PD-1 IMMUNOTHERAPY AGAINST MELANOMA IN MICE
68
4.1 ABSTRACT
The immunosuppressive tumor microenvironments (TMEs) in solid tumors undermine the
therapeutic effect of PD-1 blockades, despite the promising results exhibited by PD-1 blockades
in clinical trials. Fibroblast activation protein (FAP) is highly expressed on stromal cells in
TMEs and can be used as a therapeutic target to overcome the immunosuppressive TMEs. Here,
we developed a less immunogenic, mutant immunotoxin (αFAP-PE38), based on the original
αFAP-PE38 described in our previous study (Fang et al. 2016, 1013-1023), that can that
specifically target FAP
+
stromal cells. We found that mutant αFAP-PE38 bound to FAP with a
high binding affinity, specifically killed FAP
+
cells in vitro, and suppressed tumor growth with
the same extent as the original αFAP-PE38 in vivo. In addition, combinatorial therapy of mutant
αFAP-PE38 and PD-1 blockade exhibited further tumor regression and changed the
immunosuppressive TMEs to immune stimulatory TMEs. These results indicate that the less
immunogenic mutant αFAP-PE38 is capable of combating immunosuppressive TMEs,
facilitating the antitumor activities of cancer therapeutics and thus make it a promising approach
in clinical developments.
69
4.2 INTRODUCTION
Checkpoint inhibitors have revolutionized the field of cancer immunotherapy by allowing
treatments to circumvent the effects of the immunosuppressive tumor microenvironment.
Programmed death-1 (PD-1) is a major checkpoint receptor that has been studied extensively. It
has been shown that PD-1 expressed on T cells downregulates signaling induced by the
interaction between antigens and T cell receptors, reduces secretion of pro-inflammatory
cytokines when bound to either of its two ligands (PD-L1 and PD-L2) on antigen presenting cells
(APCs) and tumor cells, and therefore hinders T cell effector function(Keir et al. 2008a, 677-
704). As a result, blockade of the interaction between PD-1 and its ligands will restore T cell
effector functions (Sharma and Allison 2015, 56-61). Antibodies that block PD-1 binding have
been evaluated in clinical trials and have resulted in tumor regression in 30%-50% of patients
with various diseases, ranging from melanoma to lung cancer (Lipson et al. 2013, 462-468;
Brahmer et al. 2012a, 2455-2465; Wolchok et al. 2010, 155-164).
Nonetheless, only a minority of patients experience a significant response with PD-1 therapy.
Often, the blockade of a single inhibitory pathway is not sufficient enough to overcome the
immunosuppressive tumor microenvironments (TMEs) that impair T cell functions and promote
angiogenesis and metastasis(Sharma and Allison 2015, 56-61; Mellman, Coukos, and Dranoff
2011, 480-489; Drake 2012, viii41-viii46; Topalian et al. 2012, 2443-2454; Larkin et al. 2015,
23-34). Cancer associated fibroblasts (CAFs) are the major non-hematopoietic cell types that
play crucial roles in the immunosuppressive functions of TMEs (Kalluri and Zeisberg 2006, 392-
401). CAFs promote the polarization of M2 macrophages, which are known to promote
70
angiogenesis and induce proliferation of blood endothelial cells (BECs)(Coussens, Zitvogel, and
Palucka 2013, 286-291; Leek et al. 1996, 4625-4629). It has been reported that BECs upregulate
PD-L1, which suppresses T cell functions through the inhibitory PD-1 pathway (Zang et al.
2010, 1104-1112; Rodig et al. 2003, 3117-3126). Furthermore, the reduced expression of
adhesion molecules on tumor-associated BECs hinders lymphocyte extravasation (Griffioen et
al. 1996, 1111-1117; Griffioen et al. 1996, 667-673). Therefore, CAFs restrain the recruitment of
T cells (Turley, Cremasco, and Astarita 2015, 669-682). Additionally, CAFs enhance recruitment
of myeloid cells, including macrophages and myeloid derived suppressor cells (MDSCs),
through the secretion of chemokines (Kim et al. 2012, 60-66; Paunescu et al. 2011, 635-646).
FAP is the major phenotypic marker of CAFs and studies have shown that depletion of FAP
+
cells result in reduced tumor growth and metastasis (Loeffler et al. 2006a, 1955-1962; Liao et al.
2009, e7965). Therefore, therapeutics targeting FAP
+
stromal cells might become a promising
way of overcoming immunosuppressive TMEs and enhancing antitumor activities.
We have previously developed a novel approach to eliminate FAP
+
stromal cells using a
recombinant immunotoxin named αFAP-PE38 and experiments showed that the immunotoxin
bound to FAP with high affinities, specifically killed FAP
+
cells in vitro, and restrained tumor
growth in vivo (Fang et al. 2016, 1013-1023). However, PE38, which is a toxin derived from
bacteria, has been shown to be highly immunogenic especially in patients with solid tumors and
a fully functional immune system (Pai et al. 1991, 2095-2103; Pai et al. 1996, 350-353; Hassan
and Ho 2008, 46-53). The first clinical trial with PE immunotoxins led to 100% of patients
developing antibodies against the toxin. Subsequent clinical trials of PE-based immunotoxins all
resulted in an immunogenicity rate of at least 50% (Hassan and Ho 2008, 46-53; Mazor, Onda,
and Pastan 2016, 152-164). Thus, there is an urgent need to reduce the immunogenicity of these
71
bacterial toxins. Liu et al have designed a PE-based recombinant immunotoxin with low
immunogenicity through the identification and replacement of human B-cell epitopes (Liu et al.
2012, 11782-11787).
In this study, we adopted the point mutations developed by Liu et al (Liu et al. 2012, 11782-
11787) and constructed the mutant recombinant immunotoxin αFAP-PE38. We examined the
specificity and cytotoxicity of the mutant αFAP-PE38 in vitro and measured its effects on tumor
growth in a B16-F10 melanoma mouse tumor model. Afterwards, we examined the effects of
mutant αFAP-PE38 combined with a PD-1 blockade treatment on the enhancement of tumor
growth suppression (Figure 4-1) and tested the ability to alter immunosuppressive TMEs.
Figure 4-1. Illustration of the Combination Therapy of αFAP-PE38 and PD-1 Blockade.
72
4.3 MATERIALS and METHODS
4.3.1 Mice, cell line construction and cell culture
Female C57BL/6 mice were purchased from Charles River Laboratories (Wilmington, MA) and
housed in the animal facility in accordance with institute regulations. All animal experiments and
protocols were performed according to the guidelines set by the NIH and the University of
Southern California on the Care and Use of Animals. B16-F10 and 293T cells were purchased
from ATCC (Manassas, VA) and cultured in high-glucose Dulbecco’s modified Eagle medium
(Hyclone, Logan, UT) with L-glutamine (Hyclone Laboratories, Omaha, NE) supplemented with
10% fetal bovine serum (Sigma-Aldrich, St. Louis, MO). The 293T–hFAP and 293T–mFAP cell
lines were generated by stable transduction of 293T cells with lentivirus pseudotyped with
vesicular stomatitis virus glycoprotein, as described previously (Yang et al. 2008, 326-334).
4.3.2 Plasmid construction and protein purification
The sequences encoding both the original and the mutated truncated Pseudomonas exotoxin A
(PE38) described by Liu et al (Liu et al. 2012, 11782-11787)fused with a reported sequence of
species-crossreactive FAP-specific scFv (MO36) (Brocks et al. 2001, 461-469) were cloned into
the pET-28a(+) vector (Life Technologies, Grand Island, NY) separately. The plasmids were
transformed to Escherichia coli BL21 (DE3; Invitrogen) and the bacteria were grown in luria
broth media containing 100 µg/ml of kanamycin at 37°C. When OD 600 reached 0.6, isopropyl-β-
D-1-thiogalactopyranoside (Sigma-Aldrich) was added to 1 mM for 4 hours. Cells were then
harvested and the recombinant fusion protein was isolated from inclusion bodies by washing
with 2M urea buffer and dissolving in 8M urea. After renaturation by dialysis in gradient urea
73
buffer, the recombinant fusion protein was purified by Ni
2+
IDA column for His-tag purification
(Qiagen, Valencia, CA).
4.3.3 Dye labeling of αFAP-PE38
Purified αFAP-PE38 protein was incubated with 50 nmol of Alexa488-TFP ester (Invitrogen) for
2 hr in 0.1 M sodium bicarbonate buffer (pH = 9.3). The unbound dye molecules were removed
via buffer exchange into PBS (pH = 7.4) using a Zeba desalting spin column (Thermo Fisher
Scientific).
4.3.4 In vitro cytotoxicity of αFAP-PE38
Standard XTT assays were performed to measure the dose-dependent cytotoxicity of αFAP-PE38
in cultured cells using a commercial kit (Roche Scientific). Cells were seeded on 96-well plates
one day before the treatment, treated with αFAP-PE38 on day 2 and XTT assay was conducted
on day 4. PBS was used as a control for 0% cell death. The OD values were normalized between
the 100% cell death (0% line) and PBS controls (100% alive) and fit to a standard 4-parameter
sigmoidal curve with a variable slope using the GraphPad Prism (version 5.03; GraphPad
Software) program to obtain the concentration of immunotoxin at which there was 50% cell
death (IC50).
4.3.5 Tumor challenge and treatment
Female C57BL/6 mice (n = 6 per group) were subcutaneously inoculated with 2 ×10
5
B16-F10
cells on the right flank. Tumor growth was evaluated every other day by measuring tumor
diameter with calipers. Tumor volume was defined as (smallest diameter) × (longest diameter) ×
(height). Original αFAP-PE38 at the dose of 0.5 mg/kg, mutant αFAP-PE38 at the dose of
74
0.5 mg/kg or 1.5 mg/kg and PD-1 antibody at the dose of 10 mg/kg were injected to mice via i.v.
injection at day 12 post injection respectively.
4.3.6 Pharmacokinetics
Two groups of three female C57BL/6 mice were injected with 10 µg of original αFAP-PE38 and
mutant αFAP-PE38 in 0.2 mL of PBS with 0.2% HSA respectively. Blood samples were taken
from 3 separate mice within each group at time intervals of 2, 5, 10, 20, 30, and 60 minutes from
the time of injection, and each mouse was bled twice. Groups of 3 mice were bled at time
intervals of 2 and 60 minutes, 5 and 30 minutes, or 10 and 20 minutes. Serum was harvested
from the blood samples and analyzed by His Tag ELISA Detection Kit (GenScript) following the
manufacturer's protocol.
4.3.7 Flow cytometry analysis
Tumor tissue from treated mice was harvested, minced to single suspension cells and filtered
through 0.7 μm nylon strainers (BD Falcon, Franklin Lakes, NJ). The filtered cells were washed
twice with cold PBS and then incubated for 10 minutes at 4 °C with rat anti-mouse CD16/CD32
mAbs (BD Biosciences) to block nonspecific binding. Cells were then stained with monoclonal
antibodies conjugated with fluorescent dyes. All staining antibodies and isotype controls were
purchased from eBioscience or BioLegend, including anti-CD45 (30-F11), anti-CD3 (145-
2C11), anti-CD4 (RM4-5), anti-CD8 (53–6.7), anti-F4/80 (BM8), anti-ly-6G (1A8), anti-ly-
6C(1A8), anti-PD-1 (RMP1-30), anti-CD25 (PC61), anti-FoxP3 (FJK-16S), anti-CD11b
(M1/70). Tregs were identified by CD3
+
CD45
+
CD4
+
CD25
+
Foxp3
+
markers; CD4 and CD8 T
cells were identified by CD3
+
CD45
+
CD4
+
and CD3
+
CD45
+
CD8
+
markers, respectively. MDSCs
were identified by CD45
+
CD11b
+
F4/80
+
ly-6C
+
ly-6G
+
markers. Data were acquired on a
75
MACSquant cytometer (Miltenyi Biotec, San Diego, CA), and the analysis was performed using
FlowJo software (Tree Star, Ashland, OR).
4.3.8 Immunofluorescence imaging and Immunohistochemical analysis
For immunofluorescent staining, the frozen tumor sample slides were fixed with 4%
formaldehyde, permeabilized with 0.1% Triton X-100, stained with TUNEL antibody, and
followed by counterstaining with DAPI. All fluorescence images were acquired on a Yokogawa
spinning-disk confocal scanner system (Solamere Technology Group) using a Nikon eclipse Ti-E
microscope (Nikon) equipped with an x60/1.49 Apo TIRF oil objective and a Cascade II: 512
EMCCD camera (Photometrics, Tucson). Then livers and kidneys of mice were frozen and cut
into sections and stained with hematoxylin and eosin (H&E) for pathology analysis.
4.3.9 RNA isolation and transcripts analysis by RT-qPCR
Total tissue RNA was extracted from the flank tumor tissue using an RNeasy Mini Kit (Qiagen,
Valencia, CA), according to the manufacturer’s protocol. The cDNAs were synthesized from
equal amounts of total RNAs using the High-Capacity RNA-to-cDNA Kit (Applied Biosystems,
Grand Island, NY). Real-time qPCR with the appropriate primers was used to measure the
expression of Perforin, IL-12p35, IL-12p40, ICOS, and TNF-α genes. An ABI 7300 Real-Time
PCR System (Applied Biosystems) was used for real-time qPCR to measure the incorporation of
SYBR Green (Applied Biosystems). The ΔΔCt method was used to calculate changes in gene
expression level, and the raw values were normalized to the levels of GAPDH as a reference
gene.
76
4.3.10 Statistical analysis
Statistical analysis was performed by GraphPad (Prism) software to determine p values by
Student’s t-test where two groups were compared. When more than two groups were compared,
an ANOVA with the Tukey posttest was used to determine significant differences between
individual groups. Kaplan-Meier analysis was used to evaluate the survival of mice. A p value
below 0.05 was considered statistically significant, and data were presented as means ± SEM.
4.4 RESULTS
4.4.1 Construction, purification and in vitro cytotoxicity of original and mutant αFAP-PE38
There are two changes made to the mutant αFAP-PE38 compared with the original αFAP-PE38
described previously (Fang et al. 2016, 1013-1023). First, the linker was changed from the
human CD8 hinge to a furin-cleavable linker (Figure 4-2A). Second, domain II of the truncated
PE 38 was removed and seven point mutations described by Liu et al (Liu et al. 2012, 11782-
11787) were made on the truncated PE38 (Figure 4-2A). We purified the protein with a Ni-IDA
column. SDS PAGE analysis of the purified samples showed that the original αFAP-PE38 and
mutant αFAP-PE38 had the expected molecular weight of ~75KDa and ~53 KDa respectively
(Figure 4-2B).
The binding specificity of the mutant αFAP-PE38 to human and murine FAP-expressing 293T
cells was determined by using flow cytometry to measure the fluorescence of dye-labeled
immunotoxin bound to FAP. The KD of the interaction between the original/mutant αFAP-PE38
and the FAP-expressing 293 T cells was determined by Lineweaver-Burk analysis (Xiao et al.
2013, 967-977; Benedict, MacKrell, and Anderson 1997, 223-231). The KDs of mutant αFAP-
77
PE38 against human FAP (hFAP) and murine FAP(mFAP) were 7.77±0.4x10
-9
M and
2.41±0.6x10
-9
M respectively (Figure 4-1D, Figure 4-1E). Mutant αFAP-PE38 and original
αFAP-PE38 showed similar binding affinities toward FAP, indicating that the mutant αFAP-
PE38 binds both hFAP and mFAP efficiently and the modification made to PE38 do not affect
binding affinities.
The cytotoxic effect of mutant αFAP-PE38 and FAP-expressing cells in vitro was evaluated by
XTT assay. Mutant αFAP-PE38 decreased the viability of FAP-expressing target cells, but not
that of 293T cells (Figure 4-2C). However, the IC50s were calculated to be 255ng/ml for mutant
αFAP-PE38 against 293T-mFAP and 3.18 µg/ml for mutant αFAP-PE38 against 293T-hFAP
(Figure 4-2C), which are approximately 50-fold higher than those of original αFAP-PE38 (Fang
et al. 2016, 1013-1023).
78
Figure 4-2. Construction, purification and characterization of mutant αFAP-PE38. A). Schematic drawings of
original αFAP-PE38 and the mutant αFAP-PE38. B). SDS-PAGE of purified immunotoxins. Lane 1, purified
original αFAP-PE38 (75kDa) after His-tag affinity chromatography; Lane 2, purified mutant αFAP-PE38 (53kDa)
after His-tag affinity chromatography. C). Measurement of cell cytotoxicity of mutant αFAP-PE38. The cell
cytotoxicity of mutant αFAP-PE38 against 293T, 293T-mFAP and 293T-hFAP cells was performed by a standard
XTT assay with a 48-hr treatment procedure. Data are given as an IC 50 value, the concentration of immunotoxin that
causes a 50% inhibition of cell death after a 48-hr incubation with immunotoxin. All the assays were conducted in
triplicate for each cell line. Data are representative of mean±SEM. D-E). Measurement of binding specificity of
mutant αFAP-PE38 toward FAP
+
cells in vitro. The K D value of the interaction between mutant αFAP-PE38 and
cell-surface mFAP/hFAP, as determined by Lineweaver-Burk analysis. All the assays were conducted in triplicate
for each cell line. Data are representative of mean±SEM.
79
4.4.2 Both mutant and original αFAP-PE38 restrain tumor growth in vivo
Figure 4-3. Antitumor efficacy of original and mutant αFAP-PE38 in B16-F10 tumor-bearing mice. Effect of
original and mutant αFAP-PE38 on the growth of established B16-F10 melanoma cancer model was evaluated.
Female C57BL6 mice were inoculated s.c. with 2×10
5
B16-F10 cells in the right flank and then treated with original
αFAP-PE38 (0.5 mg/kg) or mutant αFAP-PE38 (0.5 mg/kg)/ (1.5 mg/kg) 12 days after tumor implantation through
i.v. injection for total of four times at the indicated days. Tumor volume(A) and body weight(B) were monitored
every other day posttreatment. Error bars, average tumor volume±SEM, n=6 for each treatment group (*p < 0.05;
**p < 0.01; ***p< 0.001). Arrows indicate time of the immunization. C). H&E staining was performed on the
kidney and liver of untreated mice and treated mice with original αFAP-PE38 or mutant αFAP-PE38 from day 22.
The scale bar represents 20 µm.
To determine the antitumor effect of the mutant αFAP-PE38 compared with the original αFAP-
PE38 in vivo, we first subcutaneously inoculated C57BL/6 mice with 2x10
5
B16-F10 cells at the
right flank. Original αFAP-PE38 (0.5 mg/kg) or mutant αFAP-PE38 (0.5 mg/kg and 1.5 mg/kg)
80
were then intravenously injected every other day from 12 days after tumor inoculation for a total
of four injections. Tumor volume and body weight were measured throughout the injections.
PBS was injected as a control group. Mice group receiving immunotoxin injections showed
apparent tumor growth inhibition compared with control group and there is no obvious
difference between original αFAP-PE38 and mutant αFAP-PE38 in terms of tumor inhibition
capacity (Figure 4-3A). The mice receiving the higher dose of mutant αFAP-PE38 did not show
further restraint of tumor growth (Figure 4-3A). Therefore, the dose of mutant αFAP-PE38 was
set to be 0.5 mg/kg for subsequent experiments. No significant weight loss was observed in any
group throughout the experiment (Figure 4-3B). Results of H&E stainings of liver and kidney
tissue also indicated that no off-target toxicity was present (Figure 4-3C).
81
4.4.3 Pharmacokinetics showed a faster metabolic rate of mutant αFAP-PE38
Figure 4-4. Pharmacokinetics of original and mutant αFAP-PE38. C57BL/6 mice were injected intravenously
with 10µg of either original or mutant αFAP-PE38 and bled at several intervals between 2 and 60 minutes from the
time of injection. The concentration of the immunotoxin in the plasma at the various intervals was determined by
His-Tag ELISA and fit to a single exponential decay function. The corresponding half-life (t 1/2) is indicated.
C57BL/6 mice were injected intravenously with a single dose of 10 µg of either mutant αFAP-
PE38 or original αFAP-PE38. Blood samples were drawn at different time intervals between 2
mins and 60 mins after injection and the concentration of immunotoxin was measured by His-
Tag ELISA. Data were fit into a single exponential decay function (Figure 4-4). The mutant
αFAP-PE38 exhibited a shorter half-life time (t1/2) compared with that of original αFAP-PE38,
which are 9.1 min and 15.3min respectively.
82
4.4.4 Combinatorial therapy of PD-1 blockade and mutant αFAP-PE38 shows improved
antitumor activity
Figure 4-5. Combinatorial antitumor efficacy of mutant αFAP-PE38 and αPD-1. Female C57BL6 mice were
inoculated s.c. with 2×10
5
B16-F10 cells in the right flank and then treated with mutant αFAP-PE38 (0.5 mg/kg),
αPD-1(10mg/kg) or combined injections 12 days after tumor implantation through i.v. injection for a total of four
times at the indicated days. Tumor volume (A) and body weight (B) were monitored every other day posttreatment.
Error bars, average tumor volume ±SEM, n = 6 for each treatment group (*p < 0.05; **p < 0.01;
***
p< 0.001).
Arrows indicate the time of immunization. C). Representative images of apoptosis in tumor sections. Cell apoptosis
was detected by TUNEL staining (nuclei stained with DAPI, blue; apoptotic cells stained with FITC, green). The
scale bar represents 50 µm.
Although PD-1 blockade therapy has been shown to have promising clinical results, blockade of
a single inhibitory pathway is not sufficient to overcome immunosuppressive TMEs that
83
attenuate T cell effector function(Sharma and Allison 2015, 56-61; Mellman, Coukos, and
Dranoff 2011, 480-489; Larkin et al. 2015, 23-34). To confirm our proposal that mutant αFAP-
PE38 can eliminate FAP
+
stromal cells in TMEs and boost the efficacy of PD-1 blockade to help
recover T cell effector functions, C57BL/6 mice were injected with 2x10
5
B16-F10 cells at the
right flank. Mutant αFAP-PE38 (0.5mg/kg) or PD-1 antibody (10mg/kg) or both were then
intravenously injected every other day from 12 days after tumor inoculation for a total of four
injections. PBS was injected as a control group. Combinatorial injection of PD-1 antibody and
mutant αFAP-PE38 showed a significant 4-fold tumor reduction (p< 0.001) compared with the
control group (Figure 4-5A). Combinatorial therapy also displayed enhanced tumor suppression
compared with single injection groups (p< 0.01) (Figure 4-5A). No significant change of body
weight was observed (Figure 4-5B). Immunofluorescent imaging of tumor tissues from day 22
also indicated that both PD-1 blockade and mutant αFAP-PE38 led to cell death and
combinatorial therapy showed enhanced effect of inducing the apoptosis of tumor cells (Figure
4-5C).
4.4.5 Combinatorial therapy of PD-1 blockade and mutant αFAP-PE38 alters
immunosuppressive TMEs to promote immune stimulatory TMEs.
To further verify that mutant αFAP-PE38 can alter immunosuppressive TMEs to help PD-1
antibody to restore T cell effector function, we analyzed the tumor-infiltrating lymphocytes
(TILs) of treated mice at day 22. The mutant αFAP-PE38 group showed a noticeable increase in
the percentage of CD4
+
T
cells and the PD-1 antibody group displayed a pronounced rise in the
percentage of CD8
+
T cells (Figure 4-6A and Figure 4-6E), which is consistent with the results
from Curran et al (Curran et al. 2010, 4275-4280). Meanwhile, there was an evident additional
84
increase in the percentage of CD8
+
and CD4
+
T cells in the combinatorial treatment group (p<
0.001) (Figure 4-6A and Figure 4-6E).
Figure 4-6. Combinatorial therapy of mutant αFAP-PE38 and αPD-1 increased tumor-infiltrating T cells, the
ratio of effector T cells versus Treg cells and versus myeloid-derived suppressor cells and PD-1 expression in
effector T cells within the tumor. Tumor tissues of treated mice from day 22 were harvested and purified. Cell
population was stained by various makers, and analyzed by flow cytometry for the composition of various subsets of
immune cells. (A, B) Percentages of CD8
+
and CD4
+
T cells within CD45
+
TILs. (C, D) The ratios of CD8
+
and
CD4
+
T cells to CD4
+
CD25
+
FoxP3
+
Treg cells. (E, F). The ratios of CD8
+
and CD4
+
T cells to
CD45
+
CD11b
+
F4/80
+
ly-6C
+
ly-6G
+
MDSCs. (G, H) Percentages of CD8
+
and CD4
+
T cells expressing PD-1 within
CD45
+
TILs. The data shown were individually analyzed from mice that received the indicated therapy, and t-tests
were performed to determine the statistical significance between samples (n = 6, mean ± SEM; *P < 0.05, **P <
0.01,
***
P< 0.001).
The ratio of T cells to T regulatory cells (Tregs) has been related to tumor development and
repression in both of mice and humans (Mandl et al. 2012, 19-29; Chen et al. 2012, e47219). The
mutant αFAP-PE38 group demonstrated an enhancement of both CD8/Treg and CD4/Treg ratios,
85
and the PD-1 antibody group also experienced a noticeable increase in CD8/Treg ratio (Figure 4-
6B, Figure 4-6F). The combinatorial treatment group exhibited a further improvement in
CD8/Treg and CD4/Treg ratios compared with single groups (p< 0.001) (Figure 4-6B, Figure 4-
6F). Besides Tregs, MDSC ratios were also investigated as a major composition of
immunosuppressive TMEs. The mutant αFAP-PE38 led to an enhancement of CD8/MDSC and
CD4/MDSC ratios and the combinatorial group displayed an additional increase of both ratios
compared with the control group (p< 0.001) (Figure 4-6C, Figure 4-6E). Finally, the expression
of PD-1 functioning as an inhibitory pathway receptor on TILs was measured. The PD-1
antibody displayed an evident upregulation of PD-1 in CD8
+
TILs and PD-1 in CD4
+
TILs and
the combined group showed similar results (Figure 4-6D, Figure 4-6H).
Figure 4-7. Combinatorial therapy of mutant αFAP-PE38 and αPD-1 altered immunosuppressive tumor
microenvironment. mRNA expression levels of ICOS, IL-12p35, IL-12p40, Perforin, and TNF-α were measured
from treated mice at day 22. Three tumors from each group were resected, homogenized and pooled. Total RNA was
extracted, and the mRNA expression levels were determined by real-time qPCR. Graph depicts relative levels of
mRNA after normalizing to GAPDH mRNA levels (mean ± SEM; *P < 0.05, **P < 0.01,
***
P< 0.001).
86
Next, we measured the gene expression of T cell activation markers and associated cytokines to
further investigate changes of TMEs by using real time-qPCR (RT-qPCR). Inducible T-cell
costimulator (ICOS) expression level was remarkably increased in the combined treatment group
compared with both single treatment groups (p<0.001) (Figure 4-7A). IL-12p70 is a heterodimer
that consists of IL-12p35 and IL-12p40 subunit and promotes Th1 polarization and cell-mediated
immunity (Fewell et al. 2009, 718-728; Del Vecchio et al. 2007, 4677-4685). The combined
group elevated the expression levels of both IL-12p35 and IL-12p40 (Figure 4-7B, Figure 4-7C).
Perforin, a pore forming protein in the granules of cytotoxic T lymphocytes, and TNF-α, a
cytokine that can cause acute and hypoxic death of cancer cells (Kraman et al. 2010, 827-830),
were all upregulated in both single treatment groups. Combinatorial groups displayed further
elevation of expression levels (Figure 4-7D, Figure 4-7E).
4.5 DISCUSSION
Our previous experiments with αFAP-PE38 exhibited its high affinity for FAP in vitro, specific
killing of FAP
+
cells, and robust antitumor activity in a 4T1 mouse breast cancer model, which
makes it a promising translational approach for treating human cancers (Fang et al. 2016, 1013-
1023). However, the formation of neutralizing antibodies due to the immunogenicity of
immunotoxins is a major challenge that must be addressed in order to achieve successful clinical
applications. Liu et al used phage display and point mutations to identify and remove human B-
cell epitopes in the PE38 immunotoxin and greatly reduced its immunogenicity (Liu et al. 2012,
11782-11787). In this study, we developed an improved FAP-targeting immunotoxin, mutant
αFAP-PE38, by adopting point mutations found by Liu et al (Liu et al. 2012, 11782-11787) to
silence human B cell epitopes and reduce immunogenicity. We compared the mutant αFAP-
PE38 with the original αFAP-PE38 and found that the mutant αFAP-PE38 bound FAP-
87
expressing cells with high affinities (Figure 4-2D, Figure 4-2E), specifically targeted FAP-
expressing cells in vitro (Figure 4-2C), rendered the same in vivo antitumor effect as the original
αFAP-PE38 (Figure 4-3), altered immunosuppressive TMEs to promote immune stimulatory
TMEs (Figure 4-6, Figure 4-7), and finally showed improved antitumor efficacy with PD-1
blockade (Figure 4-5).
Effective endocytosis of immunotoxins through antibody binding is crucial for the efficacy of the
therapy. We measured the binding affinity of mutant αFAP-PE38 and compared it with that of
the original αFAP-PE38 to investigate whether point mutations of PE38 will have any effect on
the binding affinity. Our results indicated similar binding affinities between original αFAP-PE38
and mutant αFAP-PE38 (Figure 4-2D, Figure 4-2E), confirming that point mutations of PE38 did
not significantly impact the binding affinity of PE38. In this study, the cytotoxicity of mutant
αFAP-PE38 was greatly compromised in vitro - about 50 times lower than the original αFAP-
PE38. However, in our animal study, there was no significant difference in the extent of tumor
regression between the original αFAP-PE38 and the mutant αFAP-PE38 (Figure 4-2C, Figure 4-
3A). There are two possible reasons for this discrepancy. First, the furin cleavable site of mutant
αFAP-PE38 is placed in the linker region, rather than hidden inside domain II of PE38, which
was the case for the original αFAP-PE38. Second, furin is a type I transmembrane protein that is
secreted by 293T cells (Seidah et al. 1998, 9-24). Therefore, the highly exposed furin cleavable site,
which is very susceptible to cleavage by furin secreted in the medium, may cause the
immunotoxin to lose its targeting specificity before endocytosis takes place and drastically
decrease its cytotoxicity. We also did not observe any off-target toxicity of mutant αFAP-PE38
with body weight measurement and H&E staining (Figure 4-2B, Figure 4-3C, Figure 4-5B),
88
which is consistent with the results from previous studies involving FAP-targeting therapeutics
(Liao et al. 2009, e7965; Loeffler et al. 2006b, 1955-1962; Wen et al. 2010, 2325-2332).
While it is undeniable that checkpoint inhibitor therapy such as PD-1 blockade have
demonstrated an efficacious inhibition of tumor growth from clinical trials, immunosuppressive
TMEs can undermine the therapeutic effect through various inhibitory pathways and promote
tumor growth (Tlsty and Coussens 2006, 119-150; Bhowmick, Neilson, and Moses 2004, 332-
337). For instance, immunosuppressive TMEs secrete paracrine factors to inhibit effector T cell
recruitment and inhibit pro-inflammatory cytokine secretion to further impair effector T cell
function (Nummer et al. 2007, 1188-1199). CAFs secrete CCL2, CCL3, CCL4 and CCL5 to
increase the recruitment of myeloid cells such as MDSCs and repel T cells by secreting CXCL12
(Kim et al. 2012, 60-66; Paunescu et al. 2011, 635-646). CAFs also reduce the expression of E-
selectin on effector T cells and lower the expression of intercellular adhesion molecule 1 (ICAM-
1) and vascular cell adhesion molecule 1 (VCAM-1) on BECs to hinder effector T cell
extravasation. CAFs also upregulate mucosal vascular addressin cell adhesion molecule 1
(MADCAM1) and CD166, which bind α4β7 integrin and CD6 respectively on Tregs to promote
their extravasation (Nummer et al. 2007, 1188-1199). In this study, mutant αFAP-PE38, when
combined with PD-1 blockade, successfully eliminated FAP
+
stromal cells, reduced the
recruitment of MDSCs and Tregs, increased the recruitment of effector T cells and pro-
inflammatory cytokines, and reduced tumor growth (Figure 4-5, Figure 4-6, Figure 4-7). PD-1
expression in CD8
+
and CD4
+
T effector cells was upregulated in the combined group, indicating
that effector T cells were accumulated and proliferated in the intratumoral region (Curran et al.
2010, 4275-4280). Thus, we have shown that eliminating FAP
+
stromal cells with an FAP-
targeting immunotoxin, combined with PD-1 blockade, can restrain tumor growth by destroying
89
a barrier to effector T cell recruitment and promoting the activation and expansion of effector
TILs.
To sum up, we successfully developed a mutant αFAP-PE38 by adopting point mutations and
deleting domain II of PE38 to reduce the immunogenicity, as described by Liu et al (Liu et al.
2012, 11782-11787). Mutant αFAP-PE38 bound to FAP expressing cells with high affinities,
specifically killed FAP expressing cells in vitro, overcame immunosuppressive tumor
microenvironments and showed improved antitumor effects when combined with PD-1 blockade.
Our pharmacokinetic data indicated that mutant αFAP-PE38 has a half-life of 9.1 min while
original αFAP-PE38 has a half-life of 15.3 min (Figure 4-4). The decreased half-life of mutant
αFAP-PE38 is due to its smaller molecular weight of ~53 KDa compared to that of original
αFAP-PE38 which is 75 KDa (Figure 4-2A) and thus makes mutant αFAP-PE38 susceptible to
glomerular filtration (Kontermann 2011, 868-876). PEGylation, which can improve circulation
time and antitumor activity of a therapeutic, is a possible avenue for improvement of the current
immunotoxin (Filpula et al. 2007, 773-784). Chimeric antigen receptor (CAR) T cells have
shown potent in vivo antitumor effects in clinical trials of blood malignancies (Park, Geyer, and
Brentjens 2016, 3312-3320; Grupp et al. 2013, 1509-1518; Kochenderfer et al. 2012, 2709-2720).
However, in solid tumors, immunosuppressive TMEs sabotaged CAR T cell function through
various pathways (Newick, Moon, and Albelda 2016, 16006; Gilham et al. 2012, 377-384). A
future study of the combinatorial therapy of CAR T cells and immunotoxins would explore the
possible effect of overcoming immunosuppressive TMEs and expand the antitumor effect of
CAR T cell therapy.
90
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Abstract (if available)
Abstract
Combination therapy, a treatment modality that combines two or more therapeutic agents, is a cornerstone of cancer therapy. The amalgamation of anti-cancer therapies enhances efficacy compared to the mono-therapy approach because it targets key pathways in a characteristically synergistic or an additive manner. This approach potentially reduces drug resistance, while simultaneously providing therapeutic anti-cancer benefits, such as reducing tumor growth and metastatic potential, arresting mitotically active cells, boosting immune response, reducing cancer stem cell populations, and inducing apoptosis. In this dissertation, one combination chemotherapy and two combination immunotherapies are presented. ❧ In the combination chemotherapy study, we used cross-linked multilamellar liposomal vesicles (cMLVs) to encapsulate and deliver the combination of carboplatin (CPT) and paclitaxel (PTX) to ovarian cancer cells at a controlled ratio. In vitro cytotoxic assays determined the strongest anti-tumor synergism to be when the drug combination was delivered at a 1:1 CPT/PTX molar ratio. Moreover, we demonstrated that our co-encapsulation strategy reduced cytotoxity and resulted in a stronger anti-tumor effect when compared to free drug combinations and individual drug-loaded cMLVs in an OVCAR8 ovarian cancer xenograft mouse model. ❧ In the first combination immunotherapy study, we demonstrated that PD-1 blockade significantly enhanced CAR T cell expansion and effector function in vitro. In the animal study, we demonstrated that the anti-PD-1 antibody enhanced the antitumor activity of CAR-T cells and prolonged overall survival. Our study supports the immunotherapeutic approaches that combine immune checkpoint blockade with CAR-T cell therapy to achieve better antitumor immunity, especially in the treatment of tumors with suppressive microenvironment. ❧ In the second combination immunotherapy study, we developed a less immunogenic, mutant immunotoxin (αFAP-PE38), based on the original αFAP-PE38 described in our previous study, that can that specifically target FAP⁺ stromal cells. We found that mutant αFAP-PE38 bound to FAP with a high binding affinity, specifically killed FAP⁺ cells in vitro, and suppressed tumor growth with the same extent as the original αFAP-PE38 in vivo. In addition, combinatorial therapy of mutant αFAP-PE38 and PD-1 blockade exhibited further tumor regression and changed the immunosuppressive TMEs to immune stimulatory TMEs. These results indicate that the less immunogenic mutant αFAP-PE38 is capable of combating immunosuppressive TMEs, facilitating the antitumor activities of cancer therapeutics and thus make it a promising approach in clinical developments.
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Creator
Zhang, Xiaoyang
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Core Title
Combination therapy for solid tumor
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Viterbi School of Engineering
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Doctor of Philosophy
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Chemical Engineering
Publication Date
11/28/2017
Defense Date
10/17/2017
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CAR T cell,chemotherapy,immunotherapy,immunotoxin,nanoparticle,OAI-PMH Harvest,PD-1 blockade
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