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Engineering immunotoxin and viral vectors for cancer therapy

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Engineering immunotoxin and viral vectors for cancer therapy

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Engineering immunotoxin and viral vectors for cancer therapy

by

Jinxu Fang

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 2015

Copyright 2015 Jinxu Fang
ii

Acknowledgements
I would like to express my sincere gratitude to my advisor and committee chair, Dr.
Pin Wang, who gave me the opportunity to join this lab to enjoy science, and for his
continuous support of my Ph.D. study and research. I would like to thank Dr. Noah
Malmstadt and Dr. Matthew Robert Pratt for serving as my dissertation committee
members and providing valuable suggestions in refining my dissertation. I would also like
to thank Dr. Katherine Shing and Dr. Malancha Gupta, who offered me valuable advice
regarding my research project and served on my qualifying exam committee.
I am grateful for working with all the talented members in the Laboratory. I would
like to thank Dr. Liang Xiao and Dr. Bingbing Dai for teaching me the techniques in
immunology. I would like to thank Dr. Kye-il Joo, Dr. Paul Bryson, Dr. Biliang Hu, Dr.
Yarong Liu, Dr. Chupei Zhang, Dr. Alex Lei, Dr. Steven Lee, Dr. Steven Frolich, and Dr.
April Tai for their help and support. I would like to thank my fellow graduate students
Xiaolu Han, Si Li, Xiaoyang Zhang, John Mac and Natnaree Siriwon who have shared this
valuable journey with me, for technical assistance, insightful discussion and support in the
lab. I would also like to thank Matthew Lim for his technical assistance.
Last but not least, I would like to thank my husband Guideng Li for his love and
encouragement. He is always there to encourage me to look forward to the bright future.
Without him, I would never have made it to where I am today.

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Table of Contents
Acknowledgements ...................................................................................................... ii

List of Figures.............................................................................................................. vi

Abstract ......................................................................................................................viii

Chapter 1. Introduction ...................................................................................................... 1
1.1 Tumor and its microenvironment .......................................................................... 1
1.2 Tumor stroma and FAP targeting in cancer therapy.............................................. 3
1.3 Immunotoxin in cancer therapy ............................................................................. 6
1.4 Lentiviral vector based cancer vaccine.................................................................. 8
1.5 Vesicular stomatitis virus..................................................................................... 10

Chapter 2. A novel fibroblast activation protein (FAP) targeting immunotoxin for the
treatment of malignant breast cancer................................................................................ 14
2.1 Introduction.......................................................................................................... 14
2.2 Materials and Methods......................................................................................... 18
2.2.1 Mice, cell line construction and cell culture.............................................. 18
2.2.2 Plasmid construction and protein purification .......................................... 19
2.2.3 Dye labeling of αFAP-PE38 and immunofluorescence imaging .............. 20
2.2.4 In vitro cytotoxicity of αFAP-PE38........................................................... 20
2.2.5 Tumor challenge and treatment ................................................................. 21
2.2.6 Immunohistochemical analysis ................................................................. 21
2.2.7 Flow cytometry analysis of TAM.............................................................. 22
2.2.8 Radiolabeling, PET imaging and biodistribution of αFAP-PE38.............. 23
2.2.9 RNA isolation and transcripts analysis by qRT-PCR................................ 24
2.2.10 Statistical analysis ......................................................................... 24
2.3 Results.................................................................................................................. 25
2.3.1 Construction, purification and in vitro cytotoxicity of FAP-PE38.......... 25
2.3.2 Biodistribution of FAP-PE38 in 4T1 tumor-bearing mice...................... 28
2.3.3 FAP-PE38 treatment slows 4T1 breast tumor growth in vivo................. 30
2.3.4 FAP-PE38 treatment alters tumor microenvironment...............................33
2.3.5 Combined FAP-PE38 and paclitaxel therapy increases antitumor
activity.................................................................................................................35
2.4 Discussion............................................................................................................ 38
2.5 Acknowledgements.............................................................................................. 42

iv

Chapter 3. Combination of a multiple antigen vaccine regimen with tumor fibroblast
targeting therapy in a melanoma mouse model.................................................................44
3.1 Introduction.......................................................................................................... 45
3.2 Materials and Methods......................................................................................... 48
3.2.1 Mice and cell line....................................................................................... 48
3.2.2 Plasmid construction and protein purification............................................ 48
3.2.3 Lentiviral vector production....................................................................... 49
3.2.4 Immunization and Tumor challenge study................................................. 49
3.2.5 Flow cytometry analysis............................................................................ 50
3.2.6 Immunohistochemical and immunofluorescence analysis ........................ 50
3.2.7 RNA isolation and transcripts analysis by qRT-PCR................................. 51
3.2.8 Statistical analysis...................................................................................... 52
3.3 Results.................................................................................................................. 52
3.3.1 Immunization with LV-3Ag confers anti-melanoma tumor activity in a
prophylactic model............................................................................................. 52
3.3.2 LV-3Ag immunization potently inhibits tumor growth in a therapeutic
model.................................................................................................................. 54
3.3.3 FAP-PE38 treatment significantly enhances the therapeutic activity of
LV-3Ag immunization........................................................................................ 56
3.3.4 Combined LV-3Ag immunization and FAP-PE38 treatment greatly
reduce cell proliferation and induces apoptosis.................................................. 59
3.3.5 The combination therapy modulates infiltration of immune cells into
tumor tissues....................................................................................................... 56
3.3.6 The combination therapy alters cytokine profile and fosters a local
immune stimulatory tumor microenvironment................................................... 62
3.4 Discussion............................................................................................................ 64

Chapter 4. Effective immunization against Mouse Melanoma by engineered dendritic
cell targeting Vesicular Stomatitis Virus and lentivector....................................................67
4.1 Introduction...........................................................................................................68
4.2 Materials and Methods..........................................................................................70
4.2.1 Mice and cell line........................................................................................70
4.2.2 Generation of BHK-SVGmu cell line..........................................................70
4.2.3 Virus............................................................................................................71
4.2.4 Confocal imaging of SVGmu pseudotyped VSV........................................71
4.2.5 BMDC culture and transduction..................................................................72
4.2.6 Intracellular Cytokine Staining ...................................................................72
4.2.8 Statistical analysis.......................................................................................73

4.3 Results...............................................................................................................73
v

4.3.1 Pseudotyping of rVSV with Sindbis virus glycoprotein..............................73
4.3.2 VSVΔG-SVGmu-GFP targets to DC-SIGN-expressing cell line and
BMDCs................................................................................................................75
4.3.3 VSVΔG-SVGmu-GFP infection promotes the maturation of DCs in vitro.76
4.3.4 Heterologous prime/boost with DCLV-gp100 and VSVΔG-SVGmu-
gp100 induces CD8+ T cell response in mice.......................................................79
4.3.5 Heterologous prime/boost with DCLV-gp100 and VSVΔG-SVGmu-
gp100 inhibit B16 melanoma growth in vivo........................................................80

REFERENCES..................................................................................................................82

vi

List of Figures
Figure 1.1 FAP: a model for tumor-stromal epithelial interactions....................................... 5

Figure 1.2 Trafficking pathway and work mechanism of recombinant immunotoxin..........7

Figure 1.3 VSV virion structure and genome organization.................................................11

Figure 1.4 VSV life cycle....................................................................................................13

Figure 2.1 Preparation and characterization of recombinant anti-FAP immunotoxin.......25

Figure 2.2 Imaging and Biodistribution of
64
Cu-labeled FAP-PE38................................28

Figure 2.3 Antitumor efficacy of FAP-PE38 in 4T1 tumor-bearing mice........................31

Figure 2.4 Targeting of FAP-expressing cells results in decreasing TAM population
in tumor and altering the release of cytokines, chemokines, growth factors and matrix
metalloproteinases..............................................................................................................33

Figure 2.5 Combined treatment with FAP-PE38 and paclitaxel displays enhanced
antitumor activity...............................................................................................................36

Figure 3.1 Protection of mice against B16 melanoma tumor cell challenge after
immunization with the LV-3Ag.........................................................................................51

Figure 3.2 Therapeutic efficacy of LV-3Ag immunization against B16 melanoma
tumor..................................................................................................................................53

Figure 3.3 Combined LV-3Ag immunization with FAP-PE38 treatment increase the
anti-tumor activity .............................................................................................................55

Figure 3.4 Combined LV-3Ag immunization and FAP-PE38 treatment inhibit tumor
cell proliferation and induces apoptosis in vivo..................................................................57

Figure 3.5 LV-3Ag immunization in combination with depletion of FAP+ stromal
cells enhances tumor infiltrating CTL activity and increase the ratio of Teffs/Treg and
Teffs/MDSCs with in tumor...............................................................................................59

Figure 3.6 LV-3Ag immunization and depletion of FAP+ stromal cells alters the
tumor immune microenvironment......................................................................................61

Figure 4.1 rVSV genome, protein expression and pseudotyping........................................72

Figure 4.2 SVGmu bearing rVSV can selectively target DC-SIGN and infect
dendritic cells in vitro.........................................................................................................74
vii

Figure 4.3 SVGmu bearing rVSV infection induces BMDCs maturation ..........................76

Figure 4.4 Heterologous DCLV prime/rVSV boost immunization elicits CD8+ T
cells response.....................................................................................................................78

Figure 4.5 Heterologous DCLV prime/rVSV boost immunization inhibit tumor growth...79

viii

Abstract
In the past two decades, the selective mechanism-based therapeutics had been
developed for cancer treatment with an improved understanding of cancer pathogenesis.
Targeted approaches can both inhibit molecular pathways, which are critical for
tumorigenesis and tumor progression, and also stimulate host immune response against
tumor growth. Recently, many investigations have evaluated the effect of the combined
therapy with targeting agents and cytotoxic molecules and found effectively improved
clinical outcomes. Herein this thesis, I explored the potential of use targeting immunotoxin
based on fibroblast activation protein (FAP), a type II transmembrane cell surface serine
protease that highly expressed in tumor-associated fibroblasts (TAFs) in most human
epithelial cancers, some soft tissue and bone sarcomas, in cancer therapeutics. I evaluated
a novel immune-based approach to specifically targeting FAP-expressing TAFs in a mouse
4T1 metastatic breast cancer model. Upon treatment with FAP-targeting immunotoxin, the
levels of various growth factors, cytokines and matrix metalloproteinases were changed in
the tumor and the recruitment of tumor infiltrating immune cells in the tumor
microenvironment was decreased. In addition, combined treatment with FAP-PE38 and
cytotoxic agent paclitaxel effectively inhibited tumor growth in vivo. These findings
highlight the potential use of TAFs targeting therapy in cancer treatment. Next, I assessed
a strategy of utilizing multi-antigens vaccine by administration of three melanoma-
associated antigens (gp100, TRP1, TRP2) in lentiviral vectors (termed LV-3Ag). The LV-
ix

3Ag immunization dramatically increased functional T cell infiltration into tumors and
generated protective and therapeutic antitumor immunity. I also studied the combination
therapy with aFAP-PE38 and LV-3Ag and they exhibited significantly enhanced antitumor
effects on tumor growth in established B16 melanoma model. The mechanism likely
involves in modulation of immune suppressive tumor microenvironment and,
consequently, activation of cytotoxic CD8+ T cells, which are capable of specifically
recognizing and destroying tumor cells. In my third study, I attempted to use dendritic cells
(DCs) targeting vesicular stomatitis virus (VSV), which displays engineered sindbis virus
glycoprotein mutant (SVGmu) on its surface to selectively binding to the DC-SIGN
protein, to express a mouse melanoma antigen (gp100) in cancer therapy. This recombinant
rVSV specifically infected DCs and stimulateed their activation and maturation in vitro. A
heterologous DCLV-gp100 prime/rVSV-Gag boost immunization elicited gp100-specific
CD8 T cell response, therefore greatly inhibiting the tumor growth and prolong the medium
survival. These findings suggest that DC-specific rVSV vectors encoding tumor antigens
can be an effective therapeutic agents and also potentiate the advantage of heterologous
immunization regimen with lentivirus to achieve more effective immune response against
cancer.

1

Chapter 1. Introduction
1.1 Tumor and its microenvironment
Cancer is a systemic disease encompassing a variety of components both tumor cells
themselves and host stromal cells. In 1889, Stephen Paget first proposed that an appropriate
host microenvironment (the "soil") is indispensable for the growth and metastases of tumor
cells (the ‘‘seed’’) (Paget 1989). Over the years, most researchers have focused on the
tumor cell itself, the genetic change, the signaling pathway and the biochemical interaction
within the cells and the cell-cell communication. Although genetic and signaling
alterations in tumor cells are critical for tumor development, they are not sufficient to
endow cancer cells with malignant properties. Over the last two decades, researchers
started to consider tumor cells and their microenvironment as a functional whole (Kunz-
Schughart and Knuechel 2002, De Wever and Mareel 2003). Accumulating evidence has
shown that the tumor microenvironment is actively participated in tumorigenesis, tumor
progression, invasion and metastasis.
The tumor microenvironment is composed of cancer cells, stromal cells, cytokines,
chemokines, extracellular matrix (ECM), and other components. Tumor progression
requires a cooperative interplay between stromal cells and cancer cells and reciprocal
feedback between ECM molecules, stromal cells and cancer cell. Stromal cell consists of
many cell types, including fibroblasts, immune cells, endothelial cells, which comprise the
blood and lymphatic circulatory systems, various bone marrow derived cells (BMDCs),
2

including neutrophils, mast cells, myeloid cell-derived suppressor cells (MDSCs) and
mesenchymal stem cells (MSCs) (Albini and Sporn 2007, Balkwill, Capasso et al. 2012).
The interactions among cells in the tumor microenvironment create a distinct circumstance
that can confer growth advantages, resist interference, and maintain epithelial
transformation. In normal tissue, the stroma functions as the main barrier against
tumorigenesis, the regulatory cytokine network keep the epithelia cell in a balanced
situation (Kalluri 2003, Nelson and Bissell 2006). In tumor microenvironment, stroma is
one of the most important components. Stromal cells can produce chemokines, growth
factors and matrix-degrading enzymes to promote angiogenesis, basement membrane
damage and tumor invasion. Furthermore, in response to the genetically changing of the
epithelial cells, the stromal cells will change their own cytokine secreting profile to
accommodate the tumor cells proliferation.
The interplay of stroma and cancer cells occurs within a dense ECM network. ECM,
a noncellular compartment of the tumor microenvironment, does not merely function as a
static structure that maintains tissue morphology but also as an important regulator of
cancer evolution. ECM is intensively remodeled (deposition and degradation) during the
whole process of cancer progression and has been shown to have direct effect on cancer
cell malignancy. ECM molecules and their metabolites can regulate cell proliferation,
migration, angiogenesis and cancer metastasis. Perturbation of ECM synthesis,
degradation, density or rigidity can considerably influence the capacity of the tumor
3

microenvironment to promote cancer cell proliferation, migration, and invasion (Barker,
Cox et al. 2012, Noel, Gutierrez-Fernandez et al. 2012).
1.2 Cancer-associated fibroblasts and FAP targeting in cancer therapy
Fibroblasts are widely distributed non-vascular, non-epithelial and non-
inflammatory cells and usually embedded within the fibrous matrix of the connective tissue.
Their functions involved in regulation of epithelial differentiation, production of ECM to
provide the scaffold for other cells, and regulation of local inflammation as well as wound
healing (Tomasek, Gabbiani et al. 2002). Fibroblasts are the resources of the ECM, such
as collegen, fibronectins, and laminin; they are also the resources of ECM-degrading
proteases such as matrix metalloproteinases (MMPs) (Simian, Hirai et al. 2001, Chang,
Chi et al. 2002). Under normal conditions, fibroblasts are in a resting state. Fibroblasts
become activated in wound healing and these activated fibroblasts as well as
myofibroblasts have a different phenotype and secretion profile. Once the wound healing
process is completed, most of the myofibroblasts are removed by apoptosis from the
granulation tissue (Desmouliere, Redard et al. 1995). Cancers have been referred to as
wounds that do not heal because of the similarity with granulation tissue, and activated
fibroblasts in the tumor microenvironment, termed cancer-associated fibroblasts (CAFs),
share some of similarities with myofibroblasts, including expression of α-smooth muscle
actin (α-SMA) and other specific markers. CAFs are the main component of cancer stroma
and studies have demonstrated that fibroblasts contribute to tumor formation and growth
4

rates ，the CAF markers α-SMA and fibroblast activation protein (FAP) are overexpressed
in several type of tumors (Cheng and Weiner 2003, Barber 2004, Nakao, Ishii et al. 2009).
It has been well established that CAFs directly stimulate tumor cell proliferation ，
invasion and metastasis by various growth factors, chemokines and cytokines. The classical
growth factors, such as fibroblast growth factor (FGF), insulin-like growth factor (IGF),
vascular endothelial growth factor (VEGF), epithelial growth factor (EGF) and the
transforming growth factor (TGF-β), contribute to tumor initiation and progression by
stimulating the cell proliferation or survival signaling pathway (Bhowmick, Neilson et al.
2004). In addition to growth factors, pro-inflammatory cytokines, such as interleukins,
interferons and members of the tumor necrosis factor family, which are produced both by
stromal and cancer cells, exert tumor-modulating effects. Chemokines, such as SDF-1 and
MIP-1, which are expressed by CAFs, can recruit immune cells and MDSCs to tumor
microenvironment, thereby promoting angiogenesis and metastasis (Gerber, Hippe et al.
2009, Matsuo, Ochi et al. 2009). Recent studies revealed that the ECM-degrading proteases,
such as the MMPs, are associated with tumor progression and invasion by facilitating
cancer cells passage across tissue boundaries and escape from the primary tumor site
(Lederle, Hartenstein et al. 2010).The gene expression profile of CAFs is much different
from normal fibroblast, implying that CAFs could be a potential target for cancer therapy.
FAP is an atypical serine protease, which exhibits both dipeptidyl peptidase and
endopeptidase activities, and therefore can degrade both gelatin and type I collagen to
5

modulate the tumor environment. As a selectively expressed protein maker on CAFs, FAP
has attracted researchers' attention during last decade (Barber 2004, Dolznig, Schweifer et
al. 2005, Kelly 2005). In order to determine the best method to target FAP-expressing cells,
a number of studies have been designed. A series of small-molecule FAP inhibitors and
prodrugs have been designed to either inhibit FAP enzymatic activity or ablating FAP-
positive cells. Targeting FAP through the use of antibodies, vaccines, and T cells have
also been successfully designed and proved to be valuable in cancer treatment (Cheng and
Weiner 2003, Henry, Lee et al. 2007, Kakarla, Chow et al. 2013, Roberts, Deonarine et al.
2013). These FAP-targeting approaches further confirmed the constant expression of
FAP in tumourigenesis and metastasis. However, the exact molecular mechanisms of FAP
regulate tumor growth and invasion remains unclear. Further research in this area will
enhance our knowledge of FAP and illuminate the physiological role of this protease for
stroma-targeted anticancer therapy.

Figure 1.1 FAP: a model for tumor-stromal epithelial interactions.
(Cheng et al., Clin. Cancer Res. 9: 1639-1647, 2003)

6

1.3 Immunotoxin in cancer therapy
Immunotoxins are engineered therapeutic proteins. They combine an antibody
fragment with a cytotoxic protein that selectively recognize target cells and kill them
depends on intracellular toxin action. The most potent immunotoxins are made from
bacterial and plant toxins. Usually, the antibody binding efficiency determines the target
specificity, and in most of the cases, immunotoxin was designed to kill cancer cells as a
therapeutic approach.
The most commonly used immunotoxins are two bacterial toxins, pseudomonas
aeruginosa exotoxin A (PE) and diphtheria toxin (DT), which have similar cell killing
mechanism. PE has three functional domains: domain Ia. domain II and domain III.
Domain Ia, located at the N terminus, is the cell-binding domain; domain II has
translocation activity; domain III is able to adenosine diphosphate (ADP)-ribosylate
elongation factor 2 and stop protein synthesis at the elongation step, which eventually leads
to cell death (figure 1.3) (Pastan, Hassan et al. 2006). The recombinant immunotoxins are
internalized via receptor-mediated endocytosis and retrograde traffics within the cell to the
endoplasmic reticulum (ER), where the cell-killing domain is translocated across the ER
membrane and delivered into the cytosol. The cell-killing domain then ADP-ribosylates
elongation factor 2 (EF2) in the cytosol, leading to apoptotic cell death. Most currently
used PE-based immunotoxins is the third generation, which only contains the elements
required to recognize and kill cells by replacing the cell-binding domain with the Fv portion
of an antibody and retaining the minimum translocation and cell-killing domains.
7

Much progress has been made on development of immunotoxins as cancer treatment
agents over the last 30 years and a number of successful results on animal tumor model
have led to further large animal pharmacology studies and to the planning and performance
of clinical trials, such as LMB-2 (target metastasis melanoma, leukemia, and lymphoma),
R F B 4 - d g A a n d H D 3 7 - d g A ( t a r g e t B - l i n e a g e a c u t
lymphoblastic leukemia), HA22(target hairy cell leukemia), anti-B4-bR(target B-cell
lymphoma), SS1P(target ovarian and pancreatic cancer), FRP5-ETA(target melanoma,
breast cancer, and colon cancer) and so on (Pai, Bookman et al. 1991, Posey, Khazaeli et
al. 2002, Azemar, Djahansouzi et al. 2003, Furman, Grossbard et al. 2011, Schindler,
Gajavelli et al. 2011). Among these immunotoxins, the targeted immunotoxin DT-IL2
(termed denileukin diftitox—trade name Ontak) has been approved by FDA for patients

Figure 1.2 The trafficking pathway and work mechanism of recombinant
immunotoxin. (Benhar et al., Antibodies. 1: 39-69, 2012)

8

with persistent or recurrent cutaneous T-cell lymphoma (Foss, Bacha et al. 2001, Frankel,
Fleming et al. 2003, Dang, Hagemeister et al. 2004). In addition, there are several reports
of using combination therapy in vivo to enhance immunotoxin-mediated antitumor action.
For example, taxol can enhance enhance immunotoxin access to tumor cells, thereby
leading to improved responses over either agent alone (He, Zhang et al. 2006, Zhang, Xiang
et al. 2007). Similarly, ABT-737, the inhibitor of survival factors Bcl2/Bcl-xl, also
enhanced immunotoxin-mediated antitumor action (Risberg, Fodstad et al. 2011, Mattoo
and FitzGerald 2013). However, several common disadvantages of using immunotoxin still
need to be overcame, such as the stability, resistance, side effect, does limitation, and
immunogenicity (Bridle, Boudreau et al. 2009) .

1.4 Lentiviral vector based cancer vaccine
T cell-mediated anti-tumor immunity is extremely important for the prevention and
control of cancers. Eradication of tumor is mainly reliant on the activation of tumor specific
T cells, especially CD8+ T cells, which can identify and kill tumor cells. Therefore, the
ideal cancer vaccine should be able to elicit potent and long-lasting T cell immune
responses to eliminate existing tumors, recognize and destroy developing tumor cells.
Research on enhancing the antitumor T cell immune response against cancer has been
carried out for decades. So far, numerous viral vectors, including retroviral vectors,
adenoviral vectors, adenoassociated virus (AAV) vectors, and lentiviral vectors (LV), have
9

been established for gene therapy and immunization purposes. Among these viral vectors,
lentiviral vectors have attracted much attention because of their ability to infect a wide
variety of cell types, which include both dividing and nondividing cells, and to integrate
into the genome with long-term foreign gene expression. In addition, lentivectors induce
less anti-vector immunity than other viral vectors(He and Falo 2006) (He, Zhang et al.
2006), which is important for their use as tolerogenic gene therapy agents or for their
repeated use in vaccination. Lentivectors have been progressively improved over time to
increase both their efficiency and biosafety. The third generation lentivectors are rev- and
tat-independent and the U3 region of 3’ LTR was completely removed from these vectors.
Thus, they are self-inactivating and the biosafety was further enhanced (Zufferey, Dull et
al. 1998). Besides, lentivectors show significant T cell adjuvant activities compared to
other vectors, probably by providing roll-like receptor ligands to professional antigen
presenting cells (Breckpot, Escors et al. 2010, Rossetti, Gregori et al. 2011).
LVs have been shown to be excellent vehicles for tumor antigen delivery to elicit
effective cellular immunity and humoral responses. LV-triggered immune responses are
mostly mediated by the viral genome and engage TLR3 and TLR7. LVs can directly
activate murine myeloid DCs (mDCs) after in vivo administration, resulting in secretion of
IFN , TNF , and upregulation of costimulatory molecules (Breckpot, Escors et al. 2010).
Upon transduction by antigen-encoding LVs, DCs are able to efficiently present the
antigens and provide antigen-specific responses either ex vivo or through re-injection to
the host. It has been demonstrated that administration of tumor associate antigen (TAA)-
10

encoded LVs directly in vivo is a promising approach for generating antitumor responses,
especially when these LVs are engineered to target APC and in particular DCs (Yang, Yang
et al. 2008, Xiao, Joo et al. 2012). The infected DC mature, present antigen and then
migrate to the lymph organ where they can efficiently prime T cells that will sequentially
migrate to the tumor site. Besides DCs, other cells of the immune system can be targeted
by lentivectors ex vivo for adoptive cell transfer or in vivo through direct vaccination.

1.5 Vesicular stomatitis virus
Vesicular stomatitis virus (VSV), which belongs to the Rhabdoviridae family of
Vesiculoviruses (Genus), is a negative-stranded RNA virus with an around 11-kb negative
sense genome. VSV is the pathogen of vesicular stomatitis, which cause vesicular lesions
on the oral cavity, nose, feet and teats on cattle, horses and swine (Letchworth, Rodriguez
et al. 1999). Human infected by VSV display flu-like symptoms that resolved within 3-4
days (Fields and Hawkins 1967). VSV has two serotypes, including Indiana (VSVI) and
New Jersy (VSV NJ) and they show similar properties (Rodriguez 2002, Lichty, Power et
al. 2004).
VSV is a bullet shaped virus, which appears to be approximately 170 nm long and
80 nm wide with envelope. The non-segmented VSV genome encodes five structural
proteins: glycoprotein (G), large polymerase protein (L), phosphoprotein (P), matrix
protein (M), and nucleoprotein (N), arbitrating to virus infection, virus gene transcription,
11

genome synthesis, virion assembly and replication. The N protein packages the viral
genome; L and P proteins interact with each other to form the ribonucleoprotein (RNP)
complex; M protein between envelope and genome interacts with the nucleocapsid core to
support the virion shape; G protein is attached to the virus surface by inserting into the lipid
bilayer with a hydrophobic transmembrane domain (Ge, Tsao et al. 2010). The genomic
RNA of VSV lacks the 5’ cap and 3’ poly A tail, but has a non-coding leader sequence of
47 nucleotides on the 3’ and a non-coding trailer sequences of 59 nucleotides on the 5’ end
that are necessary for transcription. Each of these five genes has a start and termination
sequences, as separated by an intergenic space that results in a ∼30% gradually decrease
of mRNA expression at each subsequent gene junction during the sequential transcription
process (Iverson and Rose 1981 , Lawson, Stillman et al. 1995).

Figure 1.3 VSV virion structure and genome organization.
(Jianrong Li and Yu Zhang (2012). ISBN: 978-953-51-0881-8)

12

The VSV replication cycle can be divided into three phases. In the first phase, viruses
attach to the host cell surface as mediated by G protein, then are endocytosed by a clathrin-
dependent pathway, subsequently fused with endosomal membrane at a PH lower than 6.5
and finally released the nucleocapsid core to the cytoplasm. Studies have shown that VSV-
G is essential for binding of VSV to its receptor, fusion with the target cell membrane and
internalization into the cell plasma (Harrison 2008). Earlier studies found that the cell
surface proteins were tolerated to proteolytic digestion for virus binding and the wide
tropism of VSV, suggesting the ubiquitously expressed lipid components, such as
phosphatidylserine, phosphatidylinositol, and ganglioside GM3, on plasma membrane are
the cellular receptor for VSV, because. However, the recent studies showed that LDL
receptor (LDLR) is the major receptor for VSV-G (Coil and Miller 2004, Finkelshtein,
Werman et al. 2013).
The second phase begins with viral RNP dissociates from M protein. The RNA
dependent RNA polymerase (RdRp) complex, which is composed of the L protein and P
proteins, initiates primary transcription of the viral mRNAs. The viral genes are transcribed
sequentially from the extreme 3’ end of the genome with a 47 nucleotides leader RNA,
followed by the synthesis of five individual 5’ capped and 3’ polyadenylated viral mRNAs
in the order of N-P-M-G-L. Each gene has a highly conserved sequences at the start (3’-
UUGUCnnUAG-5’) as well as end (3’-AUACUUUUUUU-5’), which constitutes critical
signals for initiation and termination of mRNA synthesis. During replication, the RdRP
synthesizes a full-length anti-genome, which serves as a template to produce full-length
13

genome for progeny virus assembling.

Figure 1.4 VSV life cycle.
(Lichty et al., TRENDS in Molecular Medicine. 10: 210-216, 2004)

After transcription, the mRNAs are translated to viral protein by host cell ribosomes
for next phase of life cycle. VSV assembly takes place at the plasma membrane, where the
viral genome is encapsidated by N protein and assembled with other viral components into
virions, which then transported to the budding site and eventually released from the cellular
plasma membrane.

14

Chapter 2
A novel fibroblast activation protein (FAP) targeting immunotoxin for
the treatment of malignant breast cancer
Fibroblast activation protein (FAP) is mainly expressed in the tumor-associated
fibroblasts (TAFs) of most human epithelial cancers. Studies have shown that FAP plays a
critical role in tumorigenesis and cancer progression, which makes it a promising selective
target for novel anticancer therapy. However, mere abrogation of FAP enzymatic activity
by small molecules is not very effective in inhibiting tumor growth. In this study, we have
evaluated a novel immune-based approach to specifically deplete FAP-expressing TAFs in
a mouse 4T1 metastatic breast cancer model. Depletion of FAP-positive stromal cells by
FAP-targeting immunotoxin FAP-PE38 altered levels of various growth factors,
cytokines, chemokines and matrix metalloproteinases, decreased the recruitment of tumor
infiltrating immune cells in the tumor microenvironment and suppressed tumor growth. In
addition, combined treatment with FAP-PE38 and paclitaxel potently inhibited tumor
growth in vivo. Our findings highlight the potential use of immunotoxin FAP-PE38 to
deplete FAP-expressing TAFs and thus provide a rationale for the use of this immunotoxin
in cancer therapy.
2.1 Introduction
Tumorigenesis is a complex multistep process involving not only genetic and
epigenetic alterations in tumor cells, but also other cells in the dysregulated
microenvironment surrounding the tumor, e.g., immune and inflammatory cells,
endothelial cells and tumor-associated fibroblasts (TAFs), collectively termed herein as
15

stromal cells. Stromal cells communicate with tumor cells, as well as inflammatory and
immune cells, directly via cell interaction and indirectly via paracrine/exocrine signaling,
protease activity and modulation of extracellular matrix (ECM) properties that alter cell-
cell tension (Tlsty and Coussens 2006). Such complex crosstalk results in a tumor
microenvironment that supports tumorigenesis, angiogenesis and metastasis (Bhowmick,
Neilson et al. 2004, Tlsty and Coussens 2006).
TAFs are primarily responsible for the synthesis, deposition, and remodeling of the
ECM, as well as the production of growth factors, cytokines and chemokines, promoting
tumor growth and metastasis (Balkwill 2002, Balkwill 2004, Kalluri and Zeisberg 2006).
The degradation of extracellular matrix depends on matrix metalloproteinases (MMPs)
(Kessenbrock, Plaks et al. 2010). MMP-2 and MMP-9 are expressed in tumor-derived
fibroblasts (Singer, Kronsteiner et al. 2002), and their expression is associated with the
invasiveness of many human cancers (Roomi, Monterrey et al. 2009). MMP-2 and MMP-
9 play an important role in tumor angiogenesis by regulating the release of vascular
endothelial growth factor (VEGF), which is the most potent inducer of tumor angiogenesis
(Kessenbrock, Plaks et al. 2010). Primary TAFs extracted from breast carcinoma patients
displayed increased expression of SDF-1 and TGF-β mRNA and enhanced TGF-β
bioactivity (Kojima, Acar et al. 2010). Autocrine TGF-β and SDF-1 signaling drives the
differentiation of tumor-promoting TAFs during tumor progression (Kojima, Acar et al.
2010). TAFs also express tumor necrosis factor-alpha (TNF- ), which can induce acute,
hypoxic death of both cancer and stromal cells (Kraman, Bambrough et al. 2010), and the
16

chemokine monocyte chemotactic protein-1 (MCP-1), which critically mediates the
recruitment of macrophage cells (Mishra, Banerjee et al. 2011). Given the essential roles
of TAFs in tumor progression and metastasis, they have recently emerged as new
promising therapeutic targets (Brennen, Isaacs et al. 2012, Liu, Li et al. 2012, Kakarla,
Chow et al. 2013).
Fibroblast activation protein (FAP), a type II transmembrane cell surface serine
protease of the dipeptidyl peptidase IV family, is expressed selectively in TAFs in most
human epithelial cancers, including malignant breast, colorectal, skin and pancreatic
cancers, as well as in some soft tissue and bone sarcomas (Liu, Li et al. 2012). It was also
detected in the stroma of human prostate cancer specimens (Tuxhorn, Ayala et al. 2002)
and in the fibroblasts or pericytes in areas of tumor angiogenesis (Santos, Jung et al. 2009).
FAP has emerged as a key regulator in cancer physiology with multiple biological
functions, such as cell motility, cell adhesion, cell invasion and angiogenesis (Brennen,
Isaacs et al. 2012, Jacob, Chang et al. 2012, Kelly, Huang et al. 2012). Expression of either
catalytically active or inactive FAP enhanced the production of MMP-9 and invasive
behavior of cells (Huang, Simms et al. 2011, Lee, Mullins et al. 2011). In addition, FAP-
expressing stromal cells have been shown to suppress antitumor immunity (Kraman,
Bambrough et al. 2010), adding another layer of complexity in FAP-mediated tumor
growth.
The dipeptidyl-peptidase activity of FAP contributes to tumor progression, as
suggested by the finding that abrogation of FAP enzymatic activity attenuates the growth
17

of HT-29 human colon carcinoma cells (Cheng, Valianou et al. 2005, Wolf, Quan et al.
2008). Most FAP-based therapeutic approaches have focused on the development of small-
molecule inhibitors of enzymatic activity, including Val-boroPro (PT-100/talabostat), Glu-
boroPro (PT-630) and LAF-237 (Narra, Mullins et al. 2007, Santos, Jung et al. 2009, Tsai,
Yeh et al. 2010, Walsh, Duncan et al. 2013). However, inhibition of FAP enzymatic
activity by small molecules, as well as their combined treatment with other chemotherapy
agents, has had little success to date in clinical trials (Kelly, Huang et al. 2012). This could
be attributed to the cyclization of these small molecules, rendering them ineffective at the
tumor site, or the non-enzymatic functions of FAP in tumor progression, based on recent
evidence showing that a catalytically inactive mutant of FAP promotes tumor growth and
invasion of breast cancer cells through non-enzymatic functions (Huang, Simms et al.
2011).
More recently, rationally designed immunotoxins that target cell-surface proteins
have emerged as a promising approach for cancer treatment (Pastan, Hassan et al. 2006,
Pastan, Hassan et al. 2007). An immunotoxin is a chimeric protein that is composed of a
modified antibody or antibody fragment linked to a toxin. Immunotoxins can bind to
specific antigens on the surface of their target cells, enter those cells through endocytosis
and eventually kill the cells by directly interfering with the molecules involved in cell
process, modifying cell membrane or by inducing apoptotic proteins. The most commonly
used immunotoxins are based on diphtheria toxin (DT) and Pseudomonas exotoxin (PE),
both of which prevent protein synthesis by inactivating EF-2 through ADP ribosylation.
18

Immunotoxins have been proven to effectively inhibit tumor growth in vitro and in vivo.
For example, anti-Fc receptor-like 1 (FCRL1) immunotoxin E9(Fv)-PE38 displayed
remarkably selective cytotoxicity on FCRL1-positive malignancies, including chronic
lymphocytic leukemia (CLL), hairy cell leukemia (HCL), mantle cell lymphoma (MCL)
and other B cell non-Hodgkin’s lymphomas (B-NHLs). Immunotoxin IL4-PE potently
inhibited growth of human glioblastoma tumors in vivo (Kioi, Seetharam et al. 2008).
Furthermore, many different immunotoxins have already been clinically evaluated
(Madhumathi and Verma 2012). Among them, DT-IL2, which targets the IL-2 receptor,
has been approved by the Federal Drug Administration (FDA) for treatment of cutaneous
T-cell lymphoma (CTCL) (Pastan, Hassan et al. 2006).
In this study, we constructed the novel immunotoxin FAP-PE38 designed to target
FAP-expressing fibroblasts within the tumor stroma and tested its efficacy in suppressing
tumor growth in a mouse 4T1 metastatic breast cancer model. In addition, we also
investigated the molecular mechanism underlying the inhibition of tumor growth mediated
by FAP-PE38. Finally, we explored the possible combination therapy of FAP-PE38 and
paclitaxel in the same mouse model.

2.2 Materials and Methods
2.2.1 Mice, cell line construction and cell culture
Female BALB/c mice were purchased from Harlan Laboratories and housed in the
19

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. Murine 4T1 mammary carcinoma
cell line was purchased from ATCC and cultured in high-glucose DMEM supplemented
with 10% fetal bovine serum. 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, Yang et al. 2008). Both cell lines were
cultured in DMEM supplemented with 10% fetal bovine serum.
2.2.2 Plasmid construction and protein purification
To generate a construct to express immunotoxin, a reported sequence of species-
crossreactive FAP-specific scFv (MO36) (Brocks, Garin-Chesa et al. 2001), fused with the
sequence encoding the truncated Pseudomonas exotoxin A (PE38), was cloned into the
pET-28a(+) vector. The plasmid was transformed to Escherichia coli BL21 (DE3;
Invitrogen) for protein expression. The bacteria were grown in luria broth media containing
100 g/ml of kanamycin at 37° C until OD600 reached 0.6, followed by the addition of
isopropyl-β-D-1-thiogalactopyranoside (IPTG, 1 mM) to the culture medium to induce
protein expression. Four hours later, cells were harvested, and the recombinant fusion
protein was isolated from inclusion bodies by washing with 2M urea buffer 4 times and
dissolving in 8M urea. After renaturation by dialysis in gradient urea buffer, the
recombinant fusion protein was subject to Ni
2+
-IDA column for His-tag-based purification.
20

The resulting protein was aliquoted and stored in a freezer at -80
o
C. By subcloning the
cDNA of PE38 into the pET28a(+) vector, pET28a-PE38 was generated to express the His-
tag-containing PE38 as a control protein. The recombinant PE38 protein was purified by
the same method as described above.
2.2.3 Dye labeling of αFAP-PE38 and immunofluorescence imaging
To label FAP-PE38 with organic dyes, purified FAP-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). After the incubation, the unbound dye molecules were removed via
buffer exchange into PBS (pH = 7.4) using a Zeba desalting spin column (Thermo Fisher
Scientific). For immunofluorescent staining, the tumor samples were fixed with 4%
formaldehyde, permeabilized with 0.1% Triton X-100, stained with TUNEL antibody and
dye labeled FAP-PE38, 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).
2.2.4 In vitro cytotoxicity of αFAP-PE38
Using a commercial kit from Roche Scientific, standard XTT assays with a 48 hr
treatment procedure were performed to measure the dose-dependent cytotoxicity of FAP-
PE38 in cultured cells. Cells were plated on 96-well dishes one day before the treatment,
21

followed by FAP-PE38 treatment on day 2 and XTT assay on day 4. The absorption at
490 nm was measured with a plate reader. 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). Triplicate sets
were measured and compiled for final data presentation.
2.2.5 Tumor challenge and treatment
BALB/c mice were injected subcutaneously with 2 × 10
5
4T1 cells on the right
flank and randomized into groups of 6 mice. Treatment was started 7 days post-tumor
inoculation. Paclitaxel (PTX) formulated in Cremophor/ethanol (1:1, v/v) and FAP-PE38
diluted with 0.9% NaCl were administered to mice at the dose of 10 mg/kg and 0.5 mg/kg
via i.v. injection, respectively. Tumor size was measured every two days, using fine
calipers, and calculated according the following equation: volume = (L × S
2
)/2, where L is
the long dimension and S is the short dimension. Survival end point was set when the tumor
volume reached 1000 mm
3
. The survival rates are presented as Kaplan-Meier curves. The
survival curves of individual groups were compared by a log-rank test.
2.2.6 Immunohistochemical analysis
Tumor tissues were excised and fixed with 4% formaldehyde for frozen section.
Acetone-fixed 5-μm sections were first treated with 0.3% hydrogen peroxide in PBS for
22

10 min to quench endogenous peroxidase. Nonspecific binding was blocked using PBS
containing 10% serum. Sections were then incubated with biotinylated anti-mouse CD31
(1:50, eBiosciences) and biotinylated anti-mouse F4/80 (1:50, eBiosciences) in blocking
buffer for 2 hr at room temperature, followed by incubation with streptavidin-conjugated
HRP for 30 min. After incubation, the slides were washed 3 times with PBS and then
developed with the DAB substrate (Abcam). After substrate development, the sections
were then washed in water, counterstained with hematoxylin, dehydrated, and mounted
with mounting medium (Richard-Allan Scientific). An in situ cell death detection kit
(Roche) was used to detect apoptotic cells in the tumor area, following the manufacturer’s
instructions.
2.2.7 Flow cytometry analysis of TAM
Tumor tissues from treated mice were harvested, minced, and then incubated with
digestion buffer (RPMI supplemented with 3 mg/ml Dispase II, 1 mg/ml Collagenase I,
Clostridium Histolyticum) for 30 min at 37° C. Digestion mixtures were filtered through
0.7 μm nylon strainers (BD Falcon), washed twice with cold PBS, and then incubated for
10 min at 4° C with rat anti-mouse CD16/CD32 mAb (BD Biosciences) to block
nonspecific binding. Cells were then stained with anti-CD206 antibody conjugated with
Alexa488 (BioLegend) and anti-F4/80 antibody conjugated with APC (BioLegend),
followed by washing with PBS and fixation with 1% paraformaldehyde. Data acquisition
and analysis were performed on a MACSquant cytometer using FlowJo software (Treestar
23

Inc.).
2.2.8 Radiolabeling, PET imaging and biodistribution of αFAP-PE38
Radiolabeling of FAP-PE38 was performed based on a previously reported method
for AmBaSar-mediated
64
Cu labeling of proteins and peptides (Li, Jin et al. 2011). Briefly,
5 μg of AmBaSar (11.1 μmol) dissolved in 100 μl water and 1.9 μg of EDC (10 μmol)
dissolved in 100 μl water were mixed, and the pH was adjusted to 4.00 by 1 N NaOH. N-
hydroxysulfosuccinimide sodium salts (SNHS) (1.9 μg, 8.8 μmol) was then added to the
stirring mixture on ice-bath, and the pH was adjusted to 5.5. The reaction was incubated at
4° C for 30 min. Protein was then added to the AmBaSar-OSSu reaction mixture (pH 8.5)
at a ratio between 1 to 5 and 1 to 20. The reaction was incubated at 4° C overnight, after
which the AmBaSar-conjugated FAP-PE38 was purified by PD-10 column. The
AmBaSar- FAP-PE38 was then labeled with
64
Cu by addition of 1 mCi of
64
Cu in 0.1 N
phosphate buffer (pH 7.5), followed by 45 min incubation at 40° C. The
64
Cu-AmBaSar-
FAP-PE38 was purified through a PD-10 column. Positron emission tomography (PET)
imaging of the mice was performed using a rodent scanner (Concorde Microsystems).
About 100 μCi
64
Cu-AmBaSar- FAP-PE38 was diluted in a total volume of 150 μl of PBS
and injected intravenously into mice bearing established 4T1 at the right flanks (n = 3).
Static scans were obtained at 1, 3, and 24 hr post-injection. The images were reconstructed
by a Two-Dimensional Ordered Subset Expectation Maximization (2D-OSEM) algorithm.
For biodistribution, mice were sacrificed 24 hr post-injection. Tissues and organs were
24

harvested and weighed, and the accumulated radioactivity was measured using a gamma
counter. The amount of radioactivity per gram organ was given as a percentage of the total
injected dose, which was arbitrarily set to 100%.
2.2.9 RNA isolation and transcripts analysis by qRT-PCR
Total tissue RNA was extracted from flank tumor tissues using an RNeasy Mini Kit
(QIAGEN) according to the manufacturer's protocol. The cDNAs were synthesized from
equal amounts of total RNAs using the SMART™ RACE cDNA Amplification Kit (BD
Bioscience). Real-time qPCR with the appropriate primers was used to measure the
expression of VEGF, MMP-2, MMP-9, MCP1, SDF-1, CCL5, TNF-  and TGF-  genes.
The  Ct method was used to calculate change in the level of gene expression, and the
raw values were normalized to the levels of the reference gene GAPDH.
2.2.10 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 post test was used to determine significant
differences between individual groups. Kaplan-Meier analysis was used to evaluate the
survival of mice. A P value less than 0.05 was considered statistically significant, and data
were presented as means ± SEM.

25

2.3 Results
2.3.1 Construction, purification and in vitro cytotoxicity of FAP-PE38
The variable regions of the heavy and light chain of anti-FAP antibody ( FAP) were
fused to the hinge sequence of human CD8 followed by the sequence encoding the
truncated Pseudomonas exotoxin A (PE38) (Figure. S1a). This recombinant cDNA was
subcloned into a bacterial expression plasmid for recombinant protein production. The
immunotoxin FAP-PE38 was isolated from the inclusion body and then purified by Ni2+
chelate affinity chromatography. Purified immunotoxins, migrated as a monomer in non-
reducing gel (Figure. S1b) and SDS-PAGE analysis of the purified recombinant protein
confirmed that recombinant FAP-PE38 and PE38 proteins had the expected molecular
weight of ~75 and ~48 kilodalton (kDa), respectively (Figure 2.1a).
To evaluate the binding specificity of FAP-PE38 to the target FAP, human and
murine FAP-expressing cell lines were generated by stable transduction of 293T cells with
lentiviral vectors encoding the human FAP (hFAP) and murine FAP (mFAP) genes,
respectively. The expression of hFAP and mFAP in 293T cells was confirmed by flow
cytometry analysis with anti-FAP antibody staining. The results showed that more than 95%
of analyzed cells were FAP
+
cells (Figure 2.1b). We next analyzed the binding of
immunotoxin FAP-PE38 to the surface of hFAP- or mFAP-expressing 293T cells (293T-
hFAP or 293T-mFAP) by flow cytometry (Figure 2.1c). More than 94% of cells were
bound with FAP-PE38, compared with the absence of binding to negative control 293T
26

cells, suggesting that the immunotoxin FAP-PE38 binds efficiently with surface FAP in
these cells.

27

Figure 2.1 Preparation and characterization of recombinant anti-FAP immunotoxin.
(a) SDS-PAGE of purified immunotoxins. Lane 1, inclusion bodies containing FAP-
PE38 from Escherichia coli in 8M urea; Lane 2, purified FAP-PE38 after His-tag affinity
chromatography; Lane 3, PE38 inclusion bodies; Lane 4, purified PE38 after His-tag
affinity chromatography. (b) FACS analysis of hFAP or mFAP expressing on 293T cells.
The VSVG pseudotyped lentiviral vectors FUW-hFAP and FUW-mFAP were used to
transduce 293T cells and the stable transduced cells 293T-hFAP and 293T-mFAP were
stained with anti-FAP antibody to analyze FAP expression by FACS. 293T was used as
negative control (shaded area). (c) FACS analysis of immunotoxin FAP-PE38 binding to
293T control (shaded area) or 293T-mFAP/293T-hFAP (solid line) cells. (d) The KD
value of the interaction between FAP-PE38 and cell-surface mFAP/hFAP, as determined
by Lineweaver-Burk analysis. (e) The cell cytotoxicity of FAP-PE38 against 293T,
293T-mFAP and 293T-hFAP cells was performed by a standard XTT assay with a 48 hr
treatment procedure. (f) Summary of cytotoxic activity of FAP-PE38 on 293T-mFAP and
293T-hFAP cells. Data are given as an IC50 value, the concentration of immunotoxin that
causes a 50% inhibition of cell death after a 48-hour incubation with immunotoxin. All the
assays were conducted in triplicate for each cell line. Data are representative of mean 
SEM.

The binding affinity of FAP-PE38 with FAP was measured by a flow cytometry-
based assay, and the KD of the interaction between FAP-PE38 and FAP was determined
by Lineweaver-Burk kinetic analysis (Benedict, MacKrell et al. 1997, Xiao, Hung et al.
2013). The KDs of FAP-PE38 against mFAP and hFAP were 4.68± 0.69× 10
-10
M and
1.47± 0.7× 10
-9
M, respectively, indicating that FAP-PE38 bound to both hFAP and mFAP
at high affinity (Figure 2.1d). Finally, we characterized the cytotoxic effects of FAP-PE38
to target cells in vitro by performing the XTT assay, an effective method to measure cell
viability (Figure 2.1e). The immunotoxin FAP-PE38 efficiently inhibited the viability of
both 293T-hFAP and 293T-mFAP cells, with calculated IC50s of 54 ng/ml and 4 ng/ml,
respectively (Figure 2.1f). It, however, did not affect the viability of 293T cells at
concentrations up to 10 μg/ml. In addition, blocking immunotoxin FAP-PE38 binding to
FAP-positive cells using excess anti-Fab antibody abolished the cell killing activity of
28

FAP-PE38 (Figure S1c).
2.3.2 Biodistribution of FAP-PE38 in 4T1 tumor-bearing mice
To test whether FAP-PE38 specifically targets FAP-positive tumor stromal cells in
vivo, we examined the biodistribution properties of FAP-PE38 and PE38 in BALB/c mice
bearing 4T1 tumors using positron emission tomography (PET) imaging. The bifunctional
chelator AmBaSar was used in the
64
Cu labeling, as the in vivo stability of
64
Cu-AmBaSar
is better than that of other
64
Cu-chelators, such as
64
Cu-DOTA (1,4,7,10-
tetraazacyclododecane-1,4,7,10-tetraacetic acid) (Cai, Fissekis et al. 2009, Mishra,
Banerjee et al. 2011). Static microPET scans were performed in 4T1 tumor-bearing
BALB/c mice, which were injected with
64
Cu-AmBaSar-PE38 or
64
Cu-AmBaSar- FAP-
PE38 prior to imaging, at multiple time points (1 hr, 3 hr and 24 hr). Accumulation of
64
Cu-
AmBaSar- FAP-PE38 in the tumor was observed at 1 hr after injection, and a clear tumor
uptake was detected at 3 hr and 24 hr post-injection, while no obvious accumulation signal
of
64
Cu-AmBaSar-PE38 was seen in the tumor region over the same period of time (Figure
2.2a). To quantify the degree of uptake on a per organ basis, we next quantitatively
analyzed the biodistribution of
64
Cu-AmBaSar-PE38 and
64
Cu-AmBaSar- FAP-PE38 in
those mice at the 24 hr time point. Biodistribution analysis showed that
64
Cu-AmBaSar-
FAP-PE38 accumulated at 3.3-fold higher levels than
64
Cu-AmBaSar-PE38 in tumors
(p<0.01), suggesting that
64
Cu-AmBaSar- FAP-PE38 was retained in 4T1 tumors by the
desired targeted interaction between immunotoxins and FAP expression in the tumor mass
29

(Figure 2.2b). Comparable amounts of
64
Cu-AmBaSar-PE38 and
64
Cu-AmBaSar- FAP-
PE38 were detected in the liver, kidney, and spleen and, to a lesser extent, in the blood,
heart, lung, small intestine and muscle. Thus, immunotoxin FAP-PE38, but not PE38,
specifically targets FAP-positive tumor stroma.

Figure 2.2 Imaging and Biodistribution of 64Cu-labeled FAP-PE38. (a) In vivo PET
images of mice bearing 4T1 tumors at 1h, 3h and 24h after i.v. injection of
64
Cu-AmBaSar-
FAP-PE38 or
64
Cu-AmBaSar-PE38. An obvious accumulation of
64
Cu- FAP-PE38 in
tumor was observed 1 hr after administration. Mice injected with
64
Cu-AmBaSar-PE38
showed no prominent accumulation in tumor (n = 3/group). (b) Biodistribution of FAP-
PE38 and PE38 in different tissues at 24 hr after injection with
64
Cu-AmBaSar- FAP-
PE38 or
64
Cu-AmBaSar-PE38 shown as percentage of injection dose per g of tissues (%
ID/g). Tumor uptake was significantly higher in the mice injected with
64
Cu- FAP-PE38,
as compared to the mice injected with
64
Cu-PE38 (**: p<0.01). Data are representative of
two independent experiments.

30

2.3.3 FAP-PE38 treatment slows 4T1 breast tumor growth in vivo
To examine the antitumor efficacy of FAP-PE38, we tested its tumor inhibition
activity in a 4T1 breast carcinoma model. 4T1 cells, which are insensitive to αFAP-PE38
in vitro (Figure S2), were first inoculated to the right flank of BALB/c mice to allow tumor
outgrowth. Treatment was started on day 8 when the tumors reached an approximate
volume of 50 mm
3
. Mice were given four sequential intravenous injections with 0.5 mg/kg
FAP-PE38 or an equimolar dose of PE38 every other day for a total of 4 cycles, and tumor
growth and body weights were monitored over time. Mice in the group receiving 0.5 mg/kg
FAP-PE38 showed significant retardation of tumor growth (p<0.05), whereas the
treatment with an equimolar dose of PE38 exhibited no inhibition (Figure 2.3a). No
significant weight loss and bone marrow toxicity was seen for the duration of the
experiment (Figure S3a and S3b) and TUNEL staining of tissues, such as lung, liver, spleen
and kidney (Figure S4), which also display accumulation of immunotoxin, showed
no significant difference in apoptotic index between the tissues harvested from mice
treated with PE38 or FAP-PE38, indicating that the side effects of immunotoxin FAP-
PE38, if any, were minimal. Having shown the antitumor activity of FAP-PE38 in a 4T1
breast carcinoma model, we next analyzed cell viability in tumor tissue. Two consecutive
injections of FAP-PE38 or PE-38 were carried out when the tumor volume reached 100
mm
3
, and the tumor tissues were analyzed on the following day. Analysis of the tumor cells
from isolated tumor tissues by 7-AAD staining showed that FAP-PE38, but not PE38,
31

markedly reduced the number of total live cells in the tumor (Figure 2.3b). To further
investigate whether the antitumor effect of FAP-PE38 could be attributed to the induction
of apoptosis, a histopathological examination was performed by in situ TUNEL staining.
The TUNEL assay showed dramatically increased cell apoptosis in the FAP-PE38-treated
tumors as compared with that in the control PE38- treated tumor (Figure 2.3c).
Quantitatively, approximately 23% and 1.1% TUNEL-positive nuclei were observed in
FAP-PE38-treated tumors and control PE38-treated tumor, respectively (Figure 2.3d).
Finally, we examined if FAP-PE38 increases TAF cell apoptosis in the tumor tissues by
co-staining cells with TUNEL and FAP. Indeed, we found increased TUNEL-positive
and FAP-expressing tumor stroma cells in the FAP-PE38-treated tumor (Figure 2.3e).
Further, αFAP-PE38 treatment tumor sample displayed less expression of FAP protein
(Figure S5), indicating reduced population of FAP expressing cells within the tumor after
treatment. Thus, immunotoxin FAP-PE38 induces apoptosis of tumor stromal cells and
effectively inhibits tumor growth in mice.

32

Figure 2.3 Antitumor efficacy of FAP-PE38 in 4T1 tumor-bearing mice. (a) Effect of
FAP-PE38 on the growth of established 4T1 breast cancer model. Female BALB/c mice
were inoculated s.c. with 2 × 10
5
4T1 cells in the right flank and then treated with vehicle,
FAP-PE38 (0.5 mg/kg) or PE38 (0.5 mg/kg) 7 days after tumor implantation through i.v.
injection for total four time at the indicated days. Tumor volume was monitored every 3
days post-initiation of the treatment. Error bars, average tumor volume  SEM, n = 5 for
each treatment group. (b) Percentage of viable cells of isolated tumor from FAP-PE38-
and PE38-treated mice. Tumor tissue harvested from treated mice was minced and digested
33

into single cell suspension. The cells were then filtered through 0.7 μm nylon strainers,
washed twice with cold PBS, and stained with 7-AAD. (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). (d) Quantification of
TUNEL-positive cells represented in (c). Four regions of interest (ROI) were randomly
chosen per image at 10 magnification and areas of TUNEL-positive nuclei and areas of
nuclear staining were counted per region. Data were depicted as mean TUNEL-positive%
area fraction ± SEM (n=3). (e) FAP-positive stromal cells (red) undergo apoptosis as
indicated by TUNEL-positive nuclei (green) following treatment with FAP-PE38 (upper
panel), but not PE38 (bottom panel). Dye labeled FAP-PE38 was used to detect FAP
expression on the tumor section. (*: p<0.05; **: p<0.01). Data are representative of three
independent experiments.

2.3.4 FAP-PE38 treatment alters tumor microenvironment
Tumor stromal cells and infiltrating inflammatory cells, such as tumor-associated
fibroblasts (TAFs), produce excessive and different types of growth factors, cytokines,
chemokines and matrix metalloproteinases, thereby supporting tumor growth and
facilitating metastasis (Bhowmick, Neilson et al. 2004). This prompted us to hypothesize
that depletion of FAP-positive tumor stroma may alter the release of these molecules and,
consequently, change the tumor microenvironment. To test this hypothesis, we first
investigated their mRNA and protein expression in 4T1 tumor tissues collected from
FAP-PE38-treated or PE38-treated control mice (Figure 2.4a and Figure S6). Consistent
with a previous study (Kraman, Bambrough et al. 2010), the expression of TNF-  was
significantly increased in the FAP-PE38-treated tumor. The expression of
proinflammatory cytokine TGF- , which has been implicated in many aspects of
tumorigenesis by directly acting on the tumor cell, as well as influencing the tumor
microenvironment (Mocellin, Marincola et al. 2005, Connolly, Freimuth et al. 2012), was
34

significantly reduced upon FAP-PE38 treatment. MMPs and VEGF play important roles
in vasculogenesis and angiogenesis (Ferrara and Kerbel 2005).

Figure 2.4 Targeting of FAP-expressing cells results in decreasing TAM population
in tumor and altering the release of cytokines, chemokines, growth factors and matrix
metalloproteinases. (a) Relative mRNA levels of cytokines and growth factors in 4T1
tumors from tumor-bearing BALB/c mice receiving 4 i.v. injections of FAP-PE38 or
PE38. Data were normalized to the expression of GAPDH and were depicted as mean fold
change of gene expression in FAP-PE38-treated tumors compared to those in PE38-
treated tumors. Each bar is the mean of triplicate measurement for three mice. Data are
representative of mean  SEM (*: p<0.05; **: p<0.01). (b) Representative flow cytometry
plots of the population of CD206
+
F4/80
+
tumor-associated macrophages (TAMs) in 4T1
tumor tissues harvested from FAP-PE38- and PE38-treated mice. (c) The percentage of
TAMs in 4T1 tumor tissues harvested from FAP-PE38- and PE38-treated mice. Data are
representative of mean  SEM (*: p<0.05). Data are representative of three independent
experiments.

35

Immunotoxin FAP-PE38 reduced the expression of VEGF and MMP-9, but not
MMP-2. It also downregulated the expression of chemokines CCL5, MCP-1 and SDF-1.
Consistent with the downregulation of CCL5 and MCP-1, both of which can mediate the
recruitment of macrophage cells (Keophiphath, Rouault et al. 2010, Mishra, Banerjee et al.
2011), we observed a dramatic decrease of the F4/80
+
/CD206
+
tumor-associated
macrophage (TAM) population in FAP-PE38-treated tumors, as compared with the
control PE38-treated tumors (16.8% versus 5.7%, p<0.05) (Figure 2.4b and 2.4c). Thus,
immunotoxin FAP-PE38 alters the levels of growth factors, cytokines, chemokines and
matrix metalloproteinases and inhibits the recruitment of tumor associated-macrophage
cells in 4T1 tumors.
2.3.5 Combined FAP-PE38 and paclitaxel therapy increases antitumor activity
It is well accepted that macromolecules, such as antibody immunotoxins,
immunocytokines, and other immunoconjugates, enter solid tumors slowly because of the
diffusion barrier. Cytotoxic chemotherapy agents, such as paclitaxel and
cyclophosphamide, can “sensitize” the tumor to the tumoricidal effects of the
immunocytokine in the tumor microenvironment likely by increasing the uptake of
immunotcyokine into tumors (Holden, Lan et al. 2001, Zhang, Xiang et al. 2006).
We next investigated the effectiveness of the combination of FAP-PE38 and
paclitaxel in inhibiting tumor growth in vivo. The mice bearing 4T1 tumors were treated
with vehicle, two doses of paclitaxel (10 mg/kg) alone every seven days, six doses of
36

FAP-PE38 (0.5 mg/kg) alone every other day, or both FAP-PE38 and paclitaxel. When
both agents were used, paclitaxel was given 1 day before the first dose of FAP-PE38.
Mice receiving the combination of FAP-PE38 and paclitaxel treatments showed a
significantly reduced mean tumor volume (418± 77 mm
3
) compared with mice receiving
FAP-PE38 alone (770± 122 mm
3
, p<0.01), paclitaxel alone (684± 56.2 mm
3
, p<0.05), or
untreated group (1194± 62.7 mm
3
, p<0.001) on day 29 (Figure 2.5a, left panel). Further,
the survival study showed that the groups that received the combination treatment had a
median survival of 35 days and lived significantly longer than mice treated with either
agent alone (log-rank p<0.05) or those in the control group (log-rank p<0.001) (Figure 2.5a,
right panel).
Angiogenesis is critical for invasive tumor growth and metastasis (Folkman 2002).
We next evaluated the effects of FAP-PE38 on angiogenesis by performing
immunohistochemical staining of CD31, which is a superior vascular marker for
angiogenesis, on the 4T1 tumors. FAP-PE38 and paclitaxel alone inhibited angiogenesis
in 4T1 tumors, as shown by the significantly decreased vascular density in those tumors
(Figure 2.5b and 2.5c). This is consistent with the reduction of MMP-9 and VEGF
expression in FAP-PE38-treated tumors. Importantly, combined treatment with FAP-
PE38 and paclitaxel virtually abrogated the angiogenesis in 4T1 tumors. In addition, we
observed a dramatic decrease in the F4/80
+
/CD206
+
macrophage population in 4T1 tumors
treated by FAP-PE38 alone (p<0.001), but not paclitaxel alone. Combined treatment with
paclitaxel did not show further decrease of macrophage population, as compared with
37

FAP-PE38 treatment only (Figure 2.5b and 2.5d). Thus, combinatorial therapy with
FAP-PE38 and paclitaxel inhibits angiogenesis and increases antitumor activity.

Figure 2.5 Combined treatment with FAP-PE38 and paclitaxel displays enhanced
antitumor activity. (a) Inhibition of tumor growth by treatment with immunotoxin in
combination with chemotherapy. Mice were inoculated s.c. with 2 × 10
5
4T1 cells in the
right flank and treated with paclitaxel (PTX) 10mg/kg, FAP-PE38 0.5mg/kg, or the
combination of both agents through i.v. injection. Treatment regimens are depicted for all
groups. The tumor volume was assessed every 3 days and data are displayed as mean
38

tumor volume ± SEM (n = 6 mice/group). FAP-PE38/PTX versus FAP-PE38, p<0.01;
versus PTX, p<0.05; versus untreated, p<0.001. Mice received FAP-PE38/PTX treatment
demonstrated a significantly prolonged survival than mice received FAP-PE38 or PTX
alone (*: p<0.05, Kaplan–Meier survival analysis). (b) Immunohistochemical analysis of
4T1 tumors harvested from untreated mice or mice treated with FAP-PE38 alone, PTX
alone, or both PTX and FAP-PE38. Tumor sections were stained for CD31 (blood vessels)
or F4/80 (macrophages). (c,d) Quantification of CD31-positive blood vessels and F4/80-
positive macrophages in 4T1 tumors from the treated and untreated mice, as in (b). Data
are representative of two independent experiments. (*: p<0.05).

2.4 Discussion
In this study, we developed a novel FAP-targeting immunotoxin, FAP-PE38, to
target FAP-expressing stromal cells. We have constructed and purified recombined
immunotoxin FAP-PE38 and demonstrated that it bound with FAP at high affinity. Using
direct cell killing assays, we have shown that FAP-PE38 specifically killed cell lines
constitutively expressing FAP. In addition, we have also used positron emission
tomography (PET) imaging to show the targeting of FAP-PE38 to FAP-positive tumor
stroma in mice. Furthermore, we have shown that depletion of FAP-positive stromal cells
reduced tumor growth by inhibiting angiogenesis and inducing apoptosis, with altered
levels of various growth factors, cytokines, chemokines and matrix metalloproteinases in
the tumor microenvironment and a concomitant decreased recruitment of tumor-associated
macrophage cells. Finally, we have shown that paclitaxel enhanced the antitumor activity
of FAP-PE38 in mice.
Tumor stromal cells, which play an important role in all stages of carcinogenesis,
from initiation to metastasis, represent a reservoir of potential chemotherapeutic targets
(Hofmeister, Schrama et al. 2008). Strategies that endeavor to exploit cellular targets within
39

the tumor stromal cells offer several potential advantages over traditional approaches. First,
the target is more genetically stable and thus less likely to acquire resistance to a cytotoxic
agent. Second, many solid tumor malignancies share common alterations in their tumor
microenvironment; therefore, approaches that target these alterations may be widely
applied to many of these tumors.
FAP is selectively expressed on the surface of fibroblasts in the stroma of many
epithelial cancers and has emerged as a promising target for cancer treatment (Brennen,
Isaacs et al. 2012, Liu, Li et al. 2012). However, merely blocking FAP enzymatic activity
is not beneficial for inhibition of tumor growth (Huang, Simms et al. 2011, Kelly, Huang
et al. 2012), suggesting that FAP might contribute to cancer progression through additional
mechanisms other than FAP enzymatic activity per se. In our study, we have adopted a
more viable strategy by using the novel immunotoxin FAP-PE38 to specifically target
FAP-expressing stroma cells. Independent of the potentially uncertain role of FAP in
tumorigenesis, this strategy would be effective since the therapeutic effect of FAP-PE38
relies on stroma-restricted distribution of FAP, rather than strictly on inhibition of its
function. In addition to the restricted distribution of FAP in stroma, efficient internalization
of an antibody upon binding is a prerequisite for FAP in targeted therapy. Our data showed
that FAP-PE38 bound to the cell-surface expressed FAP antigen with high affinity, as
shown by the low KD of FAP-PE38 against FAP (as low as 4.68± 0.69× 10
-10
M).
Importantly, our study showed that depletion of tumor stromal cells by FAP-PE38 results
in significant inhibition of tumor growth in a mouse model and prolongs survival in a tumor
40

challenge assay. The ability of FAP-PE38 to target tumor microenvironment is confirmed
by our biodistribution study showing that FAP-PE38 was significantly accumulated at
tumor sites, as compared with that of PE38 (Figure 2.2), and our immunofluorescence
staining showing that the majority of apoptotic cells in FAP-PE38-treated tumor were
FAP-expressing tumor stroma (Figure 2.3e).
The tumor and its microenvironment exist in a dynamic and interconnected network
of reciprocal interactions that can influence cell proliferation, survival, migration, invasion
and angiogenesis (Albini and Sporn 2007, Swartz, Iida et al. 2012). Such effects are
mediated via both paracrine and autocrine stimulation by a variety of growth factors,
cytokines, chemokines and matrix metalloproteinases. Our data showed that FAP-PE38
treatment reduced the recruitment of infiltrating tumor-associated macrophages, decreased
angiogenesis to deprive tumor cells of required nutrients and oxygen, and inhibited tumor
cell growth, suggesting that elimination of the FAP-expressing population could lead to an
antitumor effect through multiple mechanisms. Such effects are likely mediated through
altering the tumor microenvironment, as suggested by the altered expression of TNF- ,
TGF-β, CCL5, MCP-1, SDF-1, VEGF and MMPs, upon FAP-PE38 treatment (Figure
3.4a). Decreased angiogenesis upon FAP-PE38 treatment is likely due to both reduced
TAFs and infiltrating TAMs, which can both release of angiogenic molecules, such as
VEGFs. In fact, one key aspect of tumor stroma formation in solid neoplasms is the
production of new blood vessels (angiogenesis) to provide an enhanced tumor blood
(Hanahan and Folkman 1996). Our finding that αFAP-PE38 immunotoxin significantly
41

reduce TAFs and TAMs, and subsequent the level of VEGF, which regulates blood vessel
growth and maturation (Conway, Collen et al. 2001, Jain 2003), suggest that αFAP-PE38
immunotoxin treatment likely prevent development of vessels.
Consistent with several previous preclinical studies targeting FAP, in which no
significant toxicities were reported (Loeffler, Kruger et al. 2006, Liao, Luo et al. 2009,
Kraman, Bambrough et al. 2010, Wen, Wang et al. 2010), FAP-PE38 treatment does not
result in significant off-target toxicity, such as weight loss and bone toxicity. In another
study, Tran and his colleagues reported bone toxicity and cachexia upon immune targeting
of FAP-expressing cells using FAP5-CAR-transduced T cells (Tran, Chinnasamy et al.
2013). Such discrepancy between this study and ours’ likely reflects differences in the
mechanism of FAP-targeting therapies. For example, the antitumor T cell response
mediated by adoptively transferred T cells is more robust and this T cells likely circulating
in the body for longer time than targeting immunotoxin, which typically have maximal
several hours half-life (Pastan, Hassan et al. 2007). It is also possible that high-dose is
associated with significant toxicity in their study as they also found that the majority of
those mice treated with 5 × 10
6
FAP5-CAR-transduced T cells did not succumb to
treatment-related toxicities.
Despite recent advances in chemotherapeutic agents for cancer, their clinical
applications are often limited by systemic toxicity. The combined use of an immunotoxin
and a chemotherapeutic agent could provide a new strategy to not only minimize potential
systemic toxicity but also maximize efficacy. Our data showed that mice treated with a
42

combination of FAP-PE38 and paclitaxel displayed a significantly increased inhibition
of tumor growth and prolonged survival as compared to that of either agent alone, and no
obvious systemic toxicity was observed. It is likely that the enhanced antitumor activity of
combined treatment can be attributed, at least in part, to increased “uptake” of
immunotoxin FAP-PE38 by stroma cells. It is also possible that the direct
immunosuppressive properties of paclitaxel may decrease formation of neutralizing
antibodies to immunotoxins and, therefore, enhance anti-tumor activity. The combined
treatment resulted in a significant inhibition of angiogenesis, but not the recruitment of
macrophage cells, suggesting that modulation of angiogenesis may be one of the important
mechanisms underpinning the enhanced antitumor activity of combined therapy.
Taken together, our findings prove the rationale for using FAP-targeting
immunotoxin to deplete FAP-expressing TAFs and demonstrate the feasibility of using this
immunotoxin to inhibit tumor growth in vivo, thus paving the way for applying this potent
and selective FAP-targeting immunotoxin for cancer therapy. During the treatment with
immunotoxins, the early production of neutralizing antibodies to immunotoxins is one of
the major limiting factors for their efficacy. Recent study has identified eight helper T-
cell epitopes, which are required for T-Cell activation and subsequent generation of
neutralizing antibodies, in PE38 (Mazor, Eberle et al. 2014). It will be interesting to make
new FAP-PE38, in which three epitopes were deleted and five others diminished by point
mutations in key residues, and to exam if this modification will obtain better antitumor
activity.
43

2.5 Acknowledgements
This work was supported by National Institutes of Health grants (R01AI068978,
R01CA170820, R01EB017206 and P01CA132681), a translational acceleration grant from
the Joint Center for Translational Medicine, the National Cancer Institute (P30CA014089),
and a grant from the Ming Hsieh Institute for Research on Engineering Medicine for
Cancer.

44

Chapter 3
Combination of a multiple antigen vaccine regimen with tumor
fibroblast targeting therapy in a melanoma mouse model
Optimum efficacy of therapeutic cancer vaccines not only requires generation of
effective antitumor immune responses but also overcoming tolerance mechanisms as
mediated by progressing tumor itself. Previous studies showed that glycoprotein 100
(gp100), tyrosinase related protein 1 (TRP1) and tyrosinase related protein 2 (TRP2) are
promising targets for melanoma immunotherapy. In this study, we have first adopted a
strategy of utilizing multi-antigens vaccine by administration of these three melanoma-
associated antigens in lentiviral vectors (termed LV-3Ag). We found that LV-3Ag
immunization dramatically increases functional T cell infiltration into tumors and
generates protective and therapeutic antitumor immunity. We have previously shown that
FAP-PE38, a novel immunotoxin FAP-PE38 designed to target FAP-expressing
fibroblasts within the tumor stroma, possess promising antitumor activity. When combined
with FAP-PE38, LV-3Ag exhibits greatly enhanced antitumor effects on tumor growth
in established B16 melanoma model. The mechanism likely involves in modulation of
immune suppressive tumor microenvironment and, consequently, activation of cytotoxic
CD8
+
T cells, which are capable of specifically recognizing and destroying tumor cells.
Thus, these results provide a strong rationale to combine immunotoxin with cancer vaccine
for the treatment of patients with advanced cancer.
45

3.1 Introduction
Therapeutic cancer vaccines and adoptive T cell immunotherapy have been
considered very attractive therapeutic approaches for the treatment of human cancer.
Identification of melanoma specific antigens, such as glycoprotein 100 (gp100), tyrosinase
related protein 1 (TRP1) and tyrosinase related protein 2 (TRP2), has given a significant
boost to develop novel melanoma targeting vaccines (Girardet, Ladisch et al. 1983,
Rosenberg 1996). Several immunization strategies have been tested to break tolerance to
melanoma differentiation antigens, including the use of viral or bacterial vectors expressing
cancer antigens, and HLA-binding peptides derived from tumor-specific antigens (Jaeger,
Bernhard et al. 1996, Overwijk, Tsung et al. 1998, Xiang, Lode et al. 2000). Immunization
with lentiviral vectors encoding TRP1 induces potent CD8
+
T cell immune responses and
antitumor immunity that prevents and inhibits B16 tumor growth (Liu, Peng et al. 2009).
Delivering melanoma antigen gp100 DNA vaccine into mice induces T cell dependent
immune responses and provides protection against subsequent tumor challenge
(Rakhmilevich, Imboden et al. 2001, Yang, Hu et al. 2011). Notably, immunization with
peptides derived from gp100 can induce a measurable antitumor immune response in
patients (Salgaller, Marincola et al. 1996). Simultaneous immunization of mice with two
melanosome antigens, hgp100 and mTRP-2, greatly protected mice from subsequent B16
melanoma tumor challenge, suggesting that immunization with the multi-antigen vaccine
will be highly effective (Mendiratta, Thai et al. 2001).
46

Despite the identification of a number of tumor-associated antigens that are
recognized by the immune system, single antigen cancer vaccines have yielded
disappointing clinical results in the last two decades. Recently, it has become apparent that
cancer vaccines alone may not work well for cancer treatment due to the immune
suppression established in the tumor lesions (Rosenberg, Yang et al. 2004). Thus, many
current therapeutic investigations on cancer focus on vaccine-based immunotherapeutic
strategies combined with conventional and novel methods of treatment (Zitvogel, Tesniere
et al. 2008, Curran, Montalvo et al. 2010). Several combination strategies are being tested
to enhance the immune response and improve the clinic outcomes in patients. For example,
cancer vaccines have been combined with other immunotherapy, with standard cancer
drugs, targeted small-molecule drugs, and local and systemic radiation of tumors, or even
laser therapy (Emens and Jaffee 2005, Gulley, Madan et al. 2007). In preclinical murine
studies, the chemotherapy agents including cyclophosphamide, doxorubicin, paclitaxel,
and docetaxel (Chu, Wang et al. 2006) enhanced antitumor immune response to a whole
tumor-cell vaccine (Machiels, Reilly et al. 2001). Combination of lentivector
immunization and low-dose chemotherapy or PD-1/PD-L1 blocking primes self-reactive T
cells and induces anti-tumor immunity (Sierro, Donda et al. 2011). A small trial combining
granulocyte–macrophage colony-stimulating factor (GM-CSF)-secreting tumor cell
vaccines with CTLA4 blockade found increased inflammatory infiltrates and tumor
regression (Hodi, Butler et al. 2008). However, combination of cancer vaccine and
immunotoxin, which are highly specific and very potent molecules that kill tumor cells
47

directly at very low concentrations, is much less studied.
Tumors consist of heterogeneous populations of cells, including both transformed
and untransformed cells, which vary among different tumors and at different tumor
progression stages (Marx 2008). However, they contain some regular cell types, such as
infiltrating inflammatory and immune cells, endothelial cells and myeloid-derived
suppressor cells (MDSCs), pericytes, and tumor-associated fibroblasts (TAFs). TAFs, a
heterogeneous fibroblast population that is phenotypically distinguished from normal
fibroblast cells, are primarily responsible for the synthesis, deposition, and remodeling of
the ECM as well as production of growth factors, cytokines and chemokines that promotes
tumor growth and metastasis (Bhowmick, Neilson et al. 2004, Loeffler, Kruger et al. 2006).
These cells selectively express high levels of fibroblast activation protein (FAP), a type II
transmembrane cell surface serine protease of the dipeptidyl peptidase IV family. As FAP-
expressing fibroblast cells have been shown to suppress antitumor immunity (Kraman,
Bambrough et al. 2010), this protein has gained great attention for targeting therapy in
recent years. In our previous study, we have shown that depletion of FAP-expressing
stromal cells reduced the recruitment of infiltrating tumor-associated macrophages,
attenuated angiogenesis to deprive tumor cells of required nutrients and oxygen, and
inhibited tumor cell growth. In addition, we and others (Kraman, Bambrough et al. 2010)
have shown that depletion of FAP-positive stromal cells can also modulate the expression
profile of growth factors, cytokines and chemokines, therefore altering the tumor
microenvironment.
48

In this study, we tested the efficacy of LVs immunization with a combination of
three antigens gp100, TRP1, TRP2 (LV-3Ag) in tumor prevention and inhibition in a
prophylactic and therapeutic B16 melanoma model. We also investigated the combination
therapy of FAP-PE38 and LV-3Ag vaccine in the same mouse model. Finally, we
explored the mechanism underlying the enhanced antitumor activity of combination
therapy.

3.2 Materials and Methods
3.2.1 Mice and cell line
C57BL/6 mice were purchased from Charles River Laboratories (Wilmington, MA).
All mice were housed in an animal facility at the University of Southern California in
accordance with institute regulations. B16-F10 and 293T cells were purchased from ATCC
(Manassas, VA) and cultured in high glucose DMEM (Hyclone, Logan, UT) with L-
glutamine (Hyclone Laboratories, Inc., Omaha, NE) supplemented with 10% FBS (Sigma-
Aldrich, St. Louis, MO).
3.2.2 Plasmid construction and protein purification
The lentiviral backbone plasmids FUW-mgp100, FUW-mTRP1 and FUW-mTRP2
was constructed by insertion of the cDNA of mouse melanoma antigen mgp100, mTRP1
and mTRP2 into the lentiviral backbone plasmid FUW downstream of the human ubiquitin
C promoter. The FAP-PE38 protein was purified as previous reported. Briefly, the
sequence encoding the truncated Pseudomonas exotoxin A (PE38) was cloned to the
49

downstream of a reported sequence of species-crossreactive FAP-specific scFv (MO36)
(Brocks, Garin-Chesa et al. 2001) in pET-28a(+) (Life Technologies, Grand Island, NY).
vector. The plasmids were transformed to the host Escherichia coli BL21 (DE3), which
was then grown in LB containing 100 g/ml of kanamycin at 37° C. At an OD600 of 0.6,
isopropyl-p-Dthiogalactopyranoside (IPTG) (Sigma-Aldrich, St. Louis, MO) was added to
a final concentration of 1mM, the culture was further incubated for 4 hr. Cells were
harvested and the recombinant protein was purified by applying to Ni
2+
IDA column
(Qiagen, Valencia, CA).
3.2.3 Lentiviral vector production
To produce lentiviral vector (LV-3Ag), the standard calcium phosphate precipitation
procedure was used in the transient transfection of virus producing cells for the making of
lentiviral vectors. HEK293T cells seeded in 15-cm culture dish (BD Biosciences, San Jose,
CA) were transiently transfected with lentiviral backbone plasmid (5 mg), packaging
plasmids (pMDLg=pRRE, 2.5 mg; pRSV-REV, 2.5 mg), and the envelope plasmid
pVSVG (2.5 mg) or pSVGmu(2.5 mg). Two days after transfection, the viral supernatant
was collected and filtered through a 0.45-μm pore size filter (Nalgene, Rochester, NY).
The supernatant was then ultracentrifugated at 25,000 rpm for 90 min, using an Optima L-
90 K preparative ultracentrifuge and an SW28 rotor. The pellets were resuspended in an
appropriate volume of cold HBSS (Hyclone, Logan, UT) for in vivo study.
3.2.4 Immunization and Tumor challenge study
For prophylactic experiments, female C57BL/6 mice (n = 5 per group) were
immunized with LV-3Ag at indicated dosage. On day 14 post immunization, the mice were
challenged with indicated number of B16-F10 cells s.c. on the right flank. Tumor growth
50

was evaluated every other day by measuring tumor diameter using a caliper. Tumor volume
was defined as (smallest diameter) ×(longest diameter) ×(height). For therapeutic
experiment, mice were challenged with indicated number of B16-F10 cells s.c. on the right
flank and immunized with LV-3Ag at day 3 post tumor challenge. The FAP-PE38
treatment was started 7 day after the immunization. Tumor volume was measured as
described above. In the survival experiment, mice were considered be dead when the
volume of tumors reached 2000 mm
2
.
3.2.5 Flow cytometry analysis
Tumor tissue from treated mice were harvested, minced to single cell, filtered
through 0.7 μm nylon strainers (BD Falcon, Franklin Lakes, NJ) and then purified by
Percoll (Sigma-Aldrich, St. Louis, MO) density gradient separation. The purified cells were
washed twice with cold PBS, and then incubated for 10 min at 4° C with rat anti-mouse
CD16/CD32 mAbs (BD Biosciences, San Jose, CA) to block the nonspecific binding. The
cells were then stained with monoclonal antibodies conjugated with fluorescent dyes. All
Staining antibodies and isotypes were purchased from eBioscience or BioLegend,
including αCD45 (30-F11), αCD3 (145-2C11), αCD4 (RM4-5), αCD8 (53-6.7), αF4/80
(BM8), αCD206 (C068C2), αPD-1 (RMP1-30), αNK1.1 (PK136), αCD25 (PC61), αFoxP3
(FJK-16S), αCD11b (M1/70), αCD11c (N418), and αGr-1 (RB6-8C5). Data was acquired
on a Macsquant cytometer (Miltenyi Biotec Inc., San Diego, CA) and the analysis were
performed using Flowjo software (Tree star Inc. Ashland, OR).
3.2.6 Immunohistochemical and immunofluorescence analysis
Tumor tissues were excised and fixed with 4% formaldehyde for frozen section.
Acetone-fixed 5-μm sections were first treated with 0.3% hydrogen peroxide in PBS for
51

10 min to quench endogenous peroxidase. Nonspecific binding was blocked using PBS
containing 10% serum. Sections were then incubated with rabbit anti-mouse Ki67 (Abcam,
Cambridge, MA) in blocking buffer for 2 hr at room temperature, followed by incubation
with HRP conjugated anti-rabbit IgG secondary antibody (Santa Cruz Biotechnology, Inc,
Santa Cruz, CA) for 30 min. After incubation, the slides were washed 3 times with PBS
and then developed with the DAB substrate (Abcam, Cambridge, MA).). After substrate
development, the sections were then washed in water, counterstained with hematoxylin,
dehydrated, and mounted with mounting medium (Richard-Allan Scientific).
H&E staining and immunohistochemistry images were acquired by EVOS® XL Cell
Imaging System (Life Technologies, Carlsbad, CA). An in situ cell death detection kit
(Roche, Indianapolis, IN) was used to detect apoptotic cells in the tumor area, following
the manufacturer’s instructions. The slides were analyzed with laser scanning by Nikon
Eclipse Ti-E microscopy with Yokogawa spinning-disk confocal scanner system
(Solamere Technology Group, Salt Lake City,UT). Quantitation of the TUNEL and Ki67
positive cells was performed using ImageJ software.
3.2.7 RNA isolation and transcripts analysis by qRT-PCR
Total tissue RNA was extracted from 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 IL-2, IFN- γ, Perforin, Granzyme B, IL-12p35, IL-
12p40, TRAIL, TNF-  and TGF-  genes. ABI 7300 real time PCR system (Applied
Biosystems, Grand Island, NY) was used for real time qPCR to measure the incorporation
52

of SYBR
®
Green (Applied Biosystems, Grand Island, NY). The  Ct method was used to
calculate the level change of the gene expression and the raw values were normalized to
the levels of GAPDH as a reference gene.
3.2.8 Statistical analysis
Statistical analysis was performed by either GraphPad
®
(Prism) or Excel
®

(Microsoft) software to determine P values by Student’s t-test. Kaplan-Meier analysis was
used to evaluate the survival of mice. A P value less than 0.05 was considered statistically
significant and data was presented as means ± SEM.

3.3 Results
3.3.1 Immunization with LV-3Ag confers anti-melanoma tumor activity in a
prophylactic model
We first investigated the effectiveness of lentiviral vaccine LV-3A in tumor
protection in a prophylactic B16 melanoma model. C57BL6 mice were primed with a
footpad injection of LV-3Ag/VSVG (5 × 106 transduction units (TU) each antigen),
followed by a boost injection consisting of LV-3Ag/SVGmu (5 ×106 TU each antigen).
Mice vaccinated with phosphate-buffered saline (PBS) at all injection points were used as
controls. The mice were challenged 10 days after boost immunization by s.c. injection of
3×106 B16-F10 melanoma cells. When the tumors reached certain size, the mice were
sacrificed and the spleen cells were harvested for further analysis (Figure 3.1a). A strong
tumor-protective immunity was observed in the LV-3Ag immunized group. One mouse
rejected tumor completely and the rest of the animals had significant suppression of tumor
53

size compared with the mice in the control groups (Figure 3.1b), confirming that the LV-
3Ag immunogen was essential for the observed tumor growth-inhibiting immune response.
It has been well established that both cytotoxic CD8 T cells and helper CD4 T
cells are involved in antitumor immunity (Fridman, Pages et al. 2012, Gu-Trantien, Loi et
al. 2013). We next analyzed the infiltration of CD4+ and CD8+ T cells in tumor

Figure 3.1 Protection of mice against B16 melanoma tumor cell challenge after
immunization with the LV-3Ag. (a) Schematic representation of immunization protocol.
Female C57BL/6 mice were immunized at day -28 with VSVG enveloped recombinant
lenti-vectors LV-3Ag (5× 10
6
TU) and boosted at day 0 with SVG enveloped recombinants.
Ten days after boost, mice were challenged subcutaneously with 3× 10
6
B16-F10 cells.
Tumor growth was monitored every other day and the tumor infiltrate T cell was detected
at day 24 after boost immunization. (b) Tumor growth curve of s.c. transplanted B16-F10
cells in C57BL/6 mice vaccinated as described above. Data are presented as mean tumor
volume ± SEM at indicated time points. Picture shows tumor tissue at day 24 after boost
immunization. (c) Representative FACS plots for measuring the population of CD8+ and
CD4+ T cell in the tumor tissue. (d) The percentage of CD8+ and CD4+ tumor infiltrated
T cells were determined at day 24 after boost immunization. (n = 5 mice/group, Error bars
indicate SEM * P<0.05, ** P<0.01)

54

tissues upon immunization with LV-3Ag, tumor cell samples from control and LV-3Ag-
immunized mice were collected, stained and subjected to FACS analysis. As shown in
Figure 3.1c, compared with control mice, the immunization resulted in more CD4+ and
CD8+ T cells infiltration in tumor tissues harvested from LV-3Ag immunized mice.
Further calculation of the T cell numbers in tumor lesions and the ratios of CD8/CD4 T
cells showed that LV-3Ag immunization dramatically increased the number of T cell
infiltrated into tumor lesions and enhanced the ratio of CD8+/CD4+ T cells (Figure 3.1d).
This suggests that both cytotoxic and helper T cells can infiltrate into the local tumor tissue
in response to LV-3Ag immunization.

3.3.2 LV-3Ag immunization potently inhibits tumor growth in a therapeutic model
We also investigated whether LV-3Ag could be potent for inhibiting tumor growth
in a therapeutic model. The immunization was carried out three days after inoculation of
0.5× 106 B16-F10 tumor cells and the tumor volume was measured every other day since
day 10 post-inoculation (Figure 3.2a). The tumor-bearing mice therapeutically vaccinated
with LV-3Ag showed significantly slower tumor growth as compared with control group
(Figure 3.2b). Likewise, we observed a significantly increased number of infiltrated CD4+
and CD8+ T cell as well as ratio of CD8+/CD4+ T cells in tumor harvested from LV-3Ag-
immunized mice, than that of control mice (Figure 3.2c and 3.2d).
Given the observation of LV-3Ag immunization-induced enhancement of tumor
infiltrating lymphocytes (TIL), we further investigated whether the expression of cytokines
with antitumor activities changes in the tumor microenvironment in response to LV-3Ag
immunization. We found that the tumor tissues isolated from LV-3Ag-immunized mice
55

exhibited significantly upregulated level of T-cell activation associated cytolytic cytokines
and enzymes IFNγ, TNF-α, perforin, and granzyme B mRNA in comparison to that from
mice with PBS injection. However, no significant difference in IL-2 mRNA expression
was observed (Figure 3.2e).

Figure 3.2 Therapuetic efficacy of LV-3Ag immunization against B16 melanoma
tumor. (a) Schematic diagram of the experimental protocol for tumor challenge and LV-
3Ag vaccination. B6 mice were s.c. challenged with 2×10
5
of B16-F10 tumor cells and
then immunized at day 3 via f.p. injection of LV-3Ag (10
7
TU). (b) Tumor growth curve
of s.c. transplanted B16-F10 cells in C57BL/6 mice vaccinated as described above. Tumor
growth was measured every other day and mice were sacrificed when the tumor volume in
control group reached around 2000mm
3
. Data are presented as mean tumor volume ±
SEM at indicated time points. (c) Representative FACS plots for measuring the population
of CD8+and CD4+ T cells in the tumor tissue. (d) The percentage of CD8+ and CD4+
infiltrated T cells. (e) The mRNA expression levels of IFN- , TNF- , IL-2, Perforin and
Granzyme B in the tumor tissues as harvested at day 18. Total RNA extracted from tumor
tissues in each group were pooled together, the mRNA expression levels were determined
by real-time RT-PCR. Graph depicts relative levels of mRNA after normalizing to GAPDH
mRNA levels. (n = 5 mice/group, Error bars indicate SEM * P<0.05, ** P<0.01)
56

3.3.3 FAP-PE38 treatment significantly enhances the therapeutic activity of LV-3Ag
immunization
Selective expression of FAP was identified to be associated with tumor stromal cells,
which makes it an ideal target for immunotherapy (Kalluri and Zeisberg 2006, Brennen,
Rosen et al. 2012). We have recently found that FAP-PE38 treatment altered levels of
various growth factors, cytokines, chemokines and matrix metalloproteinases in the tumor
microenvironment (cite our own paper). These findings prompted us to hypothesize that
FAP-PE38 treatment may augment the antitumor activity of LV-3Ag immunization. We
evaluated the efficacy of LV-3Ag immunization combined with FAP-PE38 treatment.
C57BL6 mice were injected s.c. 2× 105 B16-F10 tumor cells on day 0 and received either
no treatment, FAP-PE38 immunotoxin only, vaccine only, or vaccine plus FAP-PE38
treatment (Figure 3.3a). The combination of vaccine and FAP-PE38 treatment had shown
a statistically significant antitumor effect as compared with no treatment (P<0.0001),
vaccine alone (P < 0.001), or FAP-PE38 alone (P < 0.001) at day 20 (Figure 3.3b).
Importantly, tumor progression was delayed in >80% of mice receiving a combination of
vaccine and FAP-PE38 treatment. Further, the survival study showed that the group
receiving combination therapy had a median survival of 31 days (the tumor size of 2000
mm3 was used as a surrogate endpoint of survival), and lived significantly longer than mice
treated with FAP-PE38 alone, vaccine alone, or those in the control group (Figure 3.3c).

57

Figure 3.3 Combined LV-3Ag immunization with FAP-PE38 treatment increase the
anti-tumor activity (a) Representative FACS plots for the population of CD45-FAP+cells
in B16 tumor. (b) Tumor growth curves in B16 bearing mice. B6 mice were s.c. challenged
with 2×10
5
of B16-F10 tumor cells at right flank and then immunized at day 3 via f.p.
injection of LV-3Ag (10
7
TU). The treatment of FAP-PE38 was initiate at day 10 and
totally 4 injections were administrate at the indicated date. Tumor volume was measured
every other day and mice were killed when the tumor volume in control group reached
58

around 2000mm
3
. (n = 10 mice/group) (c) Mouse survival was calculated using the Kaplan-
Meier method. (Error bars indicate SEM * P<0.05, ** P<0.01)

3.3.4 Combined LV-3Ag immunization and FAP-PE38 treatment greatly reduce cell
proliferation and induces apoptosis
We next explored the mechanism underlying significant improved tumor growth
inhibition by combined treatment in vivo. Tumors were excised from the treated mice and
subjected to HE staining, immunohistochemistry staining of Ki-67 for cell proliferation
analysis, and TUNEL assay for cell apoptosis analysis. Immunohistochemical analyses of
tumor tissues revealed that the tumors isolated from mice receiving combined FAP-PE38
and LV-3Ag treatment presented a dramatically decreased tumor cell proliferation rate
(Figure 3.4a and 3.4b). Intratumoral proliferative index decreased by 18.91 % in the FAP-
PE38 treated group as compared to the control group. LV-3Ag immunization resulted in a
28.96 % decrease in intratumoral proliferative activity compared with controls. The
combination of LV-3Ag immunization and FAP-PE38 treatment resulted in a 48.82%
decrease in intratumoral proliferation compared with the control group and each single
treatment group. Intratumoral apoptosis in tumor tissues was examined revealing that the
apoptotic index significantly increased in the combined treatment group as compare to
FAP-PE38 treated group, LV-3Ag immunized group or control group (Figure 3.4a and
3.4b).

59

3.3.5 The combination therapy modulates infiltration of immune cells into tumor
tissues
To understand the mechanisms of the apparent enhanced antitumor activity of combined
therapy of FAP-PE38 and LV-3Ag, we analyzed the effects of single or combined
treatment on tumor-infiltrating immune cells harvested from treated mice (day 20). No
significant changes in tumor-infiltrating macrophages (TAMs) or natural killer (NK) cells
were observed in FAP-PE38 and LV-3A- treated primary tumors compared with the other
therapeutic groups (data not shown). Total CD8+T cell/Treg ratios has been linked to both
cancer progression (Bates, Fox et al. 2006, Bui, Uppaluri et al. 2006) and therapeutic
outcomes in mice and humans (Chen, Zhang et al. 2012, Mandl, Rountree et al. 2012).
We found that combined FAP-PE38 and LV-3Ag treatment, but not any single treatment,
significantly increased CD8/Treg ratios within the tumor, which has also been previously
described as predictive of therapeutical efficacy in the B16 melanoma model (Quezada,
Peggs et al. 2006) (Figure 3.5a). Similarly, we observed significantly increased CD4/Treg
ratios within the tumor in combined FAP-PE38 and LV-3Ag treatment group (Figure
3.5b). We next analyzed the activation state of the tumor-infiltrating T cells among the
different treatment groups by measuring the expression of PD-1 protein, a key co-inhibitory
receptor on tumor-infiltrating T cells. Combination of vaccine and FAP-PE38 treated
group showed a significantly lower percentage of PD-1 expressing CD8+ cells in tumors,
compared with the single LV-3Ag, FAP-PE38 or the control group (Figure 3.5c). This
effect occurred specifically in the CD8 T-cell compartment, as showed by the unchanged
percentage of PD-1 expressing CD4+ cells in tumors upon the treatment (Figure 3.5d).

60

Figure 3.4 Combined LV-3Ag immunization and FAP-PE38 treatment inhibit
tumor cell proliferation and induces apoptosis in vivo. Effects of LV-3Ag and FAP-
PE38 treatment on intratumoral proliferative and apoptotic activity. (a) Representative
images of hematoxylin and eosin (H&E) staining (magnification: x200) and
immunohistochemistry for analysis of cell proliferation marker (Ki-67) as well as the
apoptosis (TUNEL assay) in tumors that were removed from the treated mice in Figure
3.3b (magnification: x400). (b) Quantification of the number of Ki-67-positive
proliferative cells shown in (a). To quantify Ki-67-positive cells, 10 fields were randomly
chosen to count the percentage of Ki-67-positive nuclei area in nuclear staining area. Data
are represented as mean ± SD (n = 3). (c) Quantification of TUNEL positive apoptotic
tumor cell. Images were randomly selected from ten fields in each treated group. Within
one field, area of TUNEL-positive nuclei and area of nuclear staining were calculated. The
data are expressed as % total nuclear area stained by TUNEL in the field. Data are
presented as mean ± SD (n = 3).
61

Figure 3.5 LV-3Ag immunization in combination with depletion of FAP+ stromal cells
enhances tumor infiltrating CTL activity and increase the ratio of Teffs/Treg and
Teffs/MDSCs with in tumor. The population of TILs in B16 tumor tissue (day 20) as
described in Figure 3.3b. Cells were purified from harvested single cell suspension by
Percoll density gradient separation, stained by certain makers and analyzed by flow
cytometry for the composition of various immune subsets. (a), (b) Percentages of CD8+
CTL and CD4+ Teffs expressing PD-1 within CD45
+
TILs. (c), (d) The ratios of CD8+
CTL and CD4+ Teffs to CD4+CD25+FoxP3+ Treg. (e), (f) The ratios of CD8+ CTL and
CD4+ Teffs to Linage
–
CD11b
+
Gr-1
+
MDSCs. Data values show individually analyzed
mice received the indicated therapy and are for and t tests were performed to determine
statistical significance between samples. (n = 5 mice/group, Error bars indicate SEM *
P<0.05, ** P<0.01)

62

In addition to Treg cells, MDSCs, another major components of the immune
suppressive tumor microenvironment, can dampen T-cell activity within tumors that in turn
favors tumor progression (Lindau, Gielen et al. 2013). We, therefore, assessed the effects
of combined therapy on MDSCs. Single LV-3Ag vaccination or FAP-PE3 treatment
slightly increased the ratio of CD8+ T cells to MDSCs within the tumor (Figure 3.5e).
Combination of LV-3Ag vaccine and FAP-PE38 treatment significantly increased
CD8/MDSC ratios compared with single LV-3Ag vaccination, FAP-PE3 treatment or no
treatment. Furthermore, combined therapy also significantly increased the ratio of
CD4/MDSCs in tumors (Figure 3.5f).

3.3.6 The combination therapy alters cytokine profile and fosters a local immune
stimulatory tumor microenvironment
To further substantiate the shifting of tumor microenvironment, we next examined
the genes expression of T-cell activation markers and associated cytolytic cytokines in
tumor-bearing mice by quantitative RT-PCR (Figure 3.6). ICOS, T-cell activation marker,
was significantly increased in the tumors after treatment with LV-3Ag/ FAP-PE38,
compared with FAP-PE38 alone, LV-3Ag alone, or untreated tumors. More
importantly, tumors from combination-treated mice had significantly elevated mRNA
levels of perforin, a protein present in the cytoplasmic granules of CD8+ cytotoxic T
lymphocytes, TRAIL, a stimulator of apoptosis in transformed cells (Anees, Horak et al.
2011) and TNF- , which can induce acute, hypoxic death of both cancer and stromal cells
(Kraman, Bambrough et al. 2010), when compared with those from control, LV-3g-
immunized, or FAP-PE38–treated mice. IL-12p70 heterodimer, composed of IL-12p35
63

and IL-12p40 subunits, is a major Th1-driving cytokine, promoting cell-mediated tumor
immunity. Combination treatment also significantly increased mRNA level of IL-12p35,
but not IL-12p40. IL-2 has been suggested to boost anti-tumor T cell responses by acting

Figure 3.6 LV-3Ag immunization and depletion of FAP+ stromal cells alters the
tumor immune microenvironment. Analysis of mRNA expression levels of ICOS,
Perforin, Granzyme B, IL-12p35, IL-12p40, IL-2, TNF- , TGF-  and TRAIL, from tumor
tissue (day 20) as described in Figure 3B. Five tumors from each group were resected,
homogenized and pooled together. Total RNA was extracted and the mRNA expression
levels were determined by real-time RT-PCR. Graph depicts relative levels of mRNA after
normalizing to GAPDH mRNA levels (mean ± SEM; * P<0.05, ** P<0.01).

as a second costimulatory signal during CTL activation (Jackaman, Bundell et al. 2003).
FAP-PE38–treatment increase IL-2 levels and it is more profound in the combined treated
64

group. IL-2 expression was slightly increased in FAP-PE38–treatment group and such
upregulation is more profound in the combined treatment group. There is no significant
change in all groups for TGF-β expression.

3.4 Discussion
Here, we have used new strategy to improve the immunogenicity of cancer vaccines
by co-delivering a mixture of tumor differentiation antigens gp100, TRP1, TRP2 (LV-
3Ag). LV-3Ag immunization displays great antitumor activity as emphasized by the
almost completely abolished xenograft tumor formation in a prophylactic B16 melanoma
model. Notably, LV-3Ag immunization greatly increases infiltration of tumor-
infiltrating T lymphocytes and the expression of cytolytic proteins and cytokines with
antitumor activities. Furthermore, combination therapy with LV-3Ag and FAP-PE38
demonstrated remarkable antitumor effects in established tumors, a paradigm close to the
clinical situation. Underlying this effect, we observed inhibition of cell proliferation, more
apoptotic cells and significantly increased ratio of CD8+ T cells relative to Tregs and
MDSCs in tumors from the group received combination therapy. Finally, the genes of
cytolytic cytokines and enzymes such as IL-2, IL-12, TNF-  and perforin had higher
expression levels in the LV-3Ag and FAP-PE38 -treated tumors.
The choice of the antigen is one of the critical steps in tumor vaccine design.
However, due to heterogeneous expression of antigens in tumor, most single antigen
vaccine therapy possesses merely a limited therapeutic efficacy. Thus, targeting multiple
distinct tumor-associated antigens in a single vaccination regimen would likely elicit
additive or synergistic polyclonal T cell responses that prevent tumor escape from antigen
65

loss, and, therefore, conferring more effective anti-tumor efficacy. Indeed, LV-3Ag
vaccination prevents a subsequent tumor challenge and also greatly inhibits the growth of
established tumor in mice. Generation of a strong and effective anti-tumor immune
response in tumor-bearing patients, which is the key evaluation of good cancer vaccination,
requires the use of a powerful immunization strategy that can achieve highest efficacy.
Compared to DNA vaccines, which require multiple inoculations or the use of adjuvants
to improve their efficacy, lentiviral vaccines, which can infect both dividing and non-
dividing cells, are considered to be stronger immunogenes and more effective (Ura, Okuda
et al. 2014).
Immunotherapy for solid tumors has shown very promising outcome in preclinical
and early clinical studies. However, the efficacy of immunotherapy in eradicating
established tumors remains limited (Gajewski, Woo et al. 2013, Fotin-Mleczek, Zanzinger
et al. 2014). Our results showed that LV-3g immunization is less effective in inhibiting
growth of established tumor growth in a therapeutic model as compare to their completely
blockage tumor formation in a prophylactic model and such discrepancy is likely due to
the dynamic and complex microenvironment in the tumor. Tumor associated fibroblasts
(TAFs), the most preponderant cell type in the solid tumor, play a critical role in building
immunosuppressive tumor microenvironment to facilitate tumor growth and metastasis
through secreting a range of paracrine factors. For example, CAFs can disrupt IL-2
production of activated T cells and subsequently suppress the proliferation in a contact-
dependent manner (Pinchuk, Saada et al. 2008). A subset of the TAFs was found to
constitutively express programmed death ligands 1 and 2 (PD-L1 and PD-L2), which can
bind to the programmed death 1 receptor (PD-1) on T cells and impair T-cell function
66

(Nazareth, Broderick et al. 2007). Additionally, TAFs has also been suggested to further
bolster the immunosuppressive microenvironment by the recruitment of MDSCs and Tregs
(Kakarla, Song et al. 2012). TGF-β, secreted by TAFs also contributes to the expansion of
naturally occurring Tregs (Nizar, Meyer et al. 2010). These promote us to test if targeting
TAFs has the potential to improve the efficacy of LV-3Ag immunotherapy. Indeed,
combination treatment cooperates to increase the ratio of T cells to both Tregs and MDSCs
and gains significantly improved therapeutic outcome. These findings suggest that FAP-
PE38 ameliorates the immunosuppressive tumor microenvironment, which resulted in the
activation of more CTLs, the increasing release of cytolytic cytokines and, therefore, better
anti-tumor efficacy upon combined treatment.
In conclusion, this study not only provides proof of principle of the use of multiple
tumor antigens for better anti-tumor efficacy, but also highlight the potential of targeting
TAFs to improve current immunotherapy approaches for cancer.

67

Chapter 4
Effective immunization against Mouse Melanoma by engineered
dendritic cell targeting Vesicular Stomatitis Virus and lentivector
Dendritic cells (DCs) are the most powerful antigen-presenting cells (APCs) and
DC–based vaccines are a promising strategy for cancer immunotherapy due to their ability
to activate both antigen-specific T-cell immunity. However, the optimal mode of antigen
delivery and DC activation remains to be determined. We pseudotyped the vesicular
stomatitis virus (VSV) with an engineered sindbis virus glycoprotein mutant (SVGmu),
which is capable of selectively binding to the DC-SIGN protein, to achieve DC-targeting
specificity. This recombinant rVSV specifically transduce DCs and promote their
activation and maturation in vitro. When using mouse melanoma antigen (gp100) as a
model antigen, we found that heterologous DCLV-gp100 prime/ DC-rVSV-Gag boost
immunization elicited gp100-specific CD8 T cell response, therefore slow down the tumor
growth and prolong the medium survival. These findings suggest that DC–specific rVSV
vectors encoding clinically relevant antigens can be an effective prophylactic or therapeutic
vaccine; they also highlight the advantage of heterologous immunization regimen with DC-
directed LV/rVSV to elicit more effective immune response against cancer.

68

4.1 Introduction
Dendritic cells (DCs) are the most potent antigen-presenting cells (APCs) cells,
initiate and maintain both T and B cell immunity. These specialized APCs are distributed
throughout the peripheral tissue to capture pathogens and process them to peptide
epitopes then presented to naï ve T cells. After pathogens encounter, DC cells become
activated and mature into APCs that express major histocompatibility complex molecules
(MHC) and T cell co-stimulatory molecules, including CD40, CD80 and CD86. As the
expression levels of those molecules on DCs greatly affect T-cell priming, manipulating
the DC activation state is commonly used as DC-based vaccine strategies. There is growing
interest in modulating DCs to express antigens or produce immunostimulatory molecules
for therapeutic applications. DCs can be genetically modified to present desired antigens
by using liposomes, gene-gun, or by viral transduction with replication-incompetent viral
vectors. Among those methodologies, the virus-transduction technique is the most
promising approach. For example, transduction DCs with viral vectors that target two DC
surface receptor C-type lectin receptors (CLRs) DEC205 and DC-SIGN (also known as
CD209), which can interact with many viruses, including Sindbis virus, and mediate viral
entry (Zhou, Chen et al. 2006), has been approved to efficiently improve the DC antigen
presentation and induce potent T cell response (Yang, Yang et al. 2008, Morizono, Ku et
al. 2010, Maamary, Array et al. 2011, Tenbusch, Nchinda et al. 2013). Recently, our lab
has generated mutated sindbis virus glycoprotein (SVGmu), a protein unable to interact
with cell-surface heparin sulfate but retaining the ability to interact with DC-SIGN, by
introducing several mutations to wild-type SVG, and lentivectors pseudotyped with
SVGmu can modify both DC-SIGN expressing murine and human DCs in vitro with high
69

specificity and induces a substantial antigen-specific T-cell response and antibody response
in vivo (Yang, Yang et al. 2008).
Vesicular stomatitis virus (VSV) is a nonsegmented negative-strand RNA virus that
has been broadly used as a vaccine platform (Bridle, Boudreau et al. 2009, Geisbert and
Feldmann 2011). As a promising vaccine vector, recombinant VSV can accommodate large
or multiple foreign gene inserts that are expressed at high levels, easily and fast replicate
in vitro with high titer that facilitate the vaccine production, induce strong humoral and
cellular immune responses in vivo (McKenna, McGettigan et al. 2003). Moreover, VSV
does not integrate into the genome of target cell, which may cause genetic transforming or
mutagenesis. The development of a system for recovering recombinant VSVs from plasmid
DNAs by the laboratories of John Rose and Gail Wertz (Lawson, Stillman et al. 1995,
Whelan, Ball et al. 1995) provide a foundation for exploring potential of VSV as a
promising vaccine vector. In the past two decades, recombinant VSV has been developed
as vaccine candidates against many diseases and infections, such as such as influenza
(Roberts, Kretzschmar et al. 1998, Schwartz, Buonocore et al. 2011), ebola (Jones,
Feldmann et al. 2005, Geisbert, Daddario-Dicaprio et al. 2008, Wong, Audet et al. 2014),
measles (Schlereth, Buonocore et al. 2003), tuberculosis (Roediger, Kugathasan et al.
2008), AIDS (Rose, Marx et al. 2001, Egan, Chong et al. 2004, Rabinovich, Powell et al.
2014), melanoma (Diaz, Galivo et al. 2007, Wollmann, Davis et al. 2013) and breast
cancer(Ebert, Harbaran et al. 2005). However, the viral glycoprotein induced inherent
neurotoxicity (Bi, Barna et al. 1995, Johnson, Nasar et al. 2007) has been hampered its
application on human therapy. In this study, we intended to modify DCs via non-
propagating and target-specific VSV approach to both improve the safety, and increase the
70

capability of antigen presenting to achieve a better immune response. We evaluated in mice
a DC-targeting recombinant VSV vector encoding the mouse gp100 tumor associated
antigen (VSVΔG-SVGmu-gp100) as a potential vaccine for inducing an anti-gp100
immune response. We also compared the CD8 T cell immune responses elicited by a
prime/boost vaccine regimen administered with rVSV homologous prime/boost,
heterologous rVSV prime/LV boost and heterologous LV prime/ rVSV boost.

4.2 Materials and Methods
4.2.1 Mice and cell line
6-8 weeks old female BALB/c mice were purchased from Charles River Laboratories
(Wilmington, MA). All mice were maintained in an animal facility at the University of
Southern California in agreement with institute regulations. 293T and B16 cells were
purchased from ATCC (Manassas, VA) and cultured with DMEM (Hyclone, Logan, UT)
with L-glutamine (Hyclone Laboratories, Inc., Omaha, NE) supplemented with 10% FBS
(Sigma-Aldrich, St. Louis, MO). BHK-21 cells were purchased from Kerafast (Boston,
MA) cultured with DMEM supplemented with 5% FBS.
4.2.2 Generation of BHK-SVGmu cell line
The BHK-SVGmu cell lines were generated by stable transduction of BHK-21 cells
with VSVG-pseudotyped lentivirus. The cDNA encoding SVGmu were cloned to lentiviral
backbone plasmid FUW to generate FUW- SVGmu. The lentivectors packaged from 293T
cells were used to transduce BHK-21 cells. The resulting cells (designated as BHK-
SVGmu) were cultured in DMEM supplemented with 5% fetal bovine serum.
71

4.2.3 Virus
rVSV-ΔG-GFP was described previously (Whitt 2010). pVSV-ΔG-gp100 was
cloned by replacing the GFP gene with gp100 gene for the recovery of VSV-ΔG-gp100.
Recombinant VSV was recovered from plasmid DNA as previous described (Lawson,
Stillman et al. 1995, Whitt 2010). Briefly, BHK-21 cells were seeded onto 6 well plates 16
hours prior to transfection then cells were infected at a multiplicity of infection (MOI) of
5 with vTF7-3 vaccina strain. Plasmids encoding the VSV antigenomic RNA and the N,
P, L and G proteins were then transfected into these cells by using a lipofectamine 2000
transfection reagent (Invitrogen, Grand Island, NY) 30 min later. The recovered virus
was amplified with BHK-21 cells that transfected with VSV-G. For pseudotyping VSV-
ΔG with Sindbis virus glycoproteins, BHK-SVGmu cells were infected with recovered
rVSV-ΔG-GFP and rVSV-ΔG-gp100 (VSV-G in trans) at a MOI of 5 and the virus
supernatant was collected 20 hours later. The virus infectivity were tested by a standard
plaque assay. Briefly, BHK-SVGmu cells were incubated with serial dilutions of the virus
supernatant for 1 hour and then were overlaid with DMEN containing 5% FBS and 0.9%
agar to culture for 48 hours to allow the plaque form. Infectious foci were visualized by
counterstaining using a solution of crystal violet (Sigma-Aldrich, St. Louis, MO).
4.2.4 Confocal imaging of SVGmu pseudotyped VSV
To label VSV with organic dyes, purified virus was incubated with 50 nmol of Alexa
Fluor 647 (Invitrogen, Grand Island, NY) for 2 h in 0.1 M sodium bicarbonate buffer
(pH=9.3) at room temperature. After incubation, the unbound dye molecules were removed
via buffer exchange into PBS (pH=7.4), using a Zeba desalting spin column (Thermo
Fisher Scientific). The viruses were then overlaid onto polylysine coated glass-bottom
72

culture dishes and subsequently incubate with the sera from sindbis virus infected mouse
to detect the SVGmu presence on the virus surface. All Fluorescence images were acquired
on a Yokogawa spinning-disk confocal scanner system (Solamere Technology Group, Salt
Lake City, UT) using a Nikon eclipse Ti-E microscope (Nikon, Melville, NY) equipped
with an x60/1.49 Apo TIRF oil objective and a Cascade II: 512 EMCCD camera
(Photometrics, Tucson, AZ).
4.2.5 BMDC culture and transduction
BMDCs were generated following a previously described method (Yang, Yang et al.
2008). Briefly, total BM cells in femurs and tibiae of BALB/c mouse were flushed out with
RPMI 1640 medium using a 10 mL syringe and the red blood cells were lysed with TAC
buffer. Cells were then cultured in RPMI medium containing 10% FBS and GM-CSF (1:20
J558L conditioned medium) for 6 days and the resulting non-adherent cells (mostly
BMDCs) were harvested for further experiments.
4.2.6 Intracellular Cytokine Staining
Splenocytes from immunized or control mice were pooled and incubated with the
gp100 epitope for 6 h at 37 ° C in a 96-well round-bottom plate in complete RPMI medium
(10% FBS, 2 mM L-glutamine, 10 U/ml penicillin, 100 mg/ml streptomycin 10 mM
HEPES, 1 mM sodium pyruvate) and GolgiPlug (0.67 μl/ml BD Biosciences, Franklin
Lakes, NJ) was supplemented in the culture to accumulate intracellular cytokines. Surface
staining was performed by incubating restimulated cells with anti-mouse CD16/CD32
blocking antibody, followed by anti-mouse CD8 and anti-mouse CD4 antibodies. Cells
were then permeabilized in 100 l Cytofix/Cytoperm solution (BD Biosciences) at 4° C
for 10 min, washed with Perm/Wash buffer (BD Biosciences), followed by intracellular
73

staining with anti-mouse IFN-γ, TNF-α and IL-2 at 4° C for 15 min. Data was collected on
a Macsquant cytometer (Miltenyi Biotec Inc., San Diego, CA) and analyzed using FlowJo
software (Tree star Inc. Ashland, OR).
4.2.7 Statistical analysis
Statistical analysis was performed by Student’s t-test (two group comparison) or one-
way Anova followed by Bonferroni non parametric posttest (multi-comparison). P values
less than 0.05 were considered significant. The results were generated by GraphPad Prism
version 5.0 software.

4.3 Results
4.3.1 Pseudotyping of rVSV with Sindbis virus glycoprotein
To investigate if the SVGmu pseudotyped DC-targeting recombinant VSV could be
vaccine platform, we introduced the model antigen GFP and tumor antigen gp100 into the
VSV genomes at the fourth position to replace the VSV-G (Figure 4.1a). We first infected
BHK-21 cells and labeled the new synthesized protein with [
35
S] methionine to assess if
the full-length antigen proteins are being expressed correctly. The cell lysate was
fractionated by SDS-PAGE and the size-expected GFP and gp100 as well as the VSV L,
N, P, and M proteins were detected in VSVΔG-GFP and VSVΔG-gp100 infected cells
(Figure 4.1b). To further confirm the incorporation of sindbis virus glycoproteins onto
VSV particles, we performed the confocal imaging to detect the expression of SVGmu on
the virus surface. SVGmu can only be identified on the surface of virus released from the
BHK-SVGmu cell line, when these virus were stained with serum/antibody generated from
the Sindbis virus infected mouse (Figure 4.1c). To characterize the infectivity of SVGmu-
74

Figure 4.1 rVSV genome, protein expression and pseudotyping. (a) Schematic showing
the rVSV genomes with the replacement VSVG by GFP or Gag gene on the fourth position.
(b) Analysis of protein expression in infected BHK cells. Cells were infected with indicated
recombinant viruses. Six hours post infection, cells were metabolically labeled with [35S]
methionine for 2 hour and then lysed for SDS-PAGE analysis. (c) Confocal image of
SVGmu glycoprotein expression in infected BHK cells. Purified rVSV was labeled with
Alexa Fluor 647 (red) and coated to a poly-lysine containing glass-bottom dish, followed
by rinsing with PBS. The serum/antibody from sindbis virus infected mouse was used to
detect the SVGmu glycoprotein (green). Images were captured using a laser confocal
microscope. The scale bar represents 10μm. (d) The infectivity of recombinant VSVs that
expressing VSVG or SVGmu in trans. A one-step growth curve analysis was performed in
BHK-SVGmu cell line. Data are representatives of three independent experiments.

and VSV-G-bearing recombinant VSV, we next measured the infectious titer on BHK-
SVGmu cell. The BHK-SVGmu cells were infected by serially diluted viral supernatants
that collected at different time points and the virus titer was determined by plaque assay.
75

The titer of the VSV-G-pseudotyped rVSVs was calculated to be approximately 10 ×109
pfu/ml. When SVGmu was used as the envelope glycoprotein, the infectious titer was 50
times lower than VSV-G (Figure 4.1d), indicating that the foreign glycoproteins SVGmu
could substitute for VSV-G in virus assembly despite less efficient infectivity.

4.3.2 VSVΔG-SVGmu-GFP targets to DC-SIGN-expressing cell line and BMDCs
In order to determine the efficacy of SVGmu-pseudotyped rVSVs in DC-SIGN
targeting, we first used a 293T cell line (293T.hDCSIGN) that stably expresses human DC-
SIGN. Parental 293T and 293T.hDCSIGN cells were infected with VSVΔG-VSVG-GFP
and VSVΔG-SVGmu-GFP viruses at the same MOI, then analyzed for GFP expression by
flow cytometry after 24 hours culture. There was no difference between the two cell lines
in their susceptibility to infection by VSV-G–bearing viruses (13% vs 14%). By contrast,
SVGmu pseudotyped viruses infect 293T cells with a lower infectivity ( ～5%) and its
infectivity was significantly enhanced by ～4-fold in the presence of DC-SIGN expression
( ～20%) (Figure 4.2a and 4.2b). The differences of infectious efficiency between DC-
SIGN null and DC-SIGN expressing cell lines suggest that SVGmu-bearing viruses have
a better efficacy in targeting DC-SIGN expressing cells. We then evaluated the infectious
efficiency of these viruses using mouse bone marrow–derived dendritic cells (mBMDCs).
Mouse bone marrow cells were cultured with GM-CSF supplemented medium for 6 days
to generate BMDCs, which were then infected with VSVΔG-VSVG-GFP or VSVΔG-
SVGmu-GFP at the same MOI. The infectivity of VSVΔG-VSVG-GFP virus on CD11c+
cells (7.9%) was only slightly higher than on the both CD11c- cells (4.7%). In contrast,
VSVΔG-SVGmu-GFP virus was significantly (7-fold) more infectious for the CD11c+
76

cells (12.1%) than for CD11c- cells (1.7%) (Figure 4.2c and 4.2d), indicating VSVΔG-
SVGmu-GFP virus has a higher specificity in targeting mBMDCs.

Figure 4.2 SVGmu bearing rVSV can selectively target DC-SIGN and infect dendritic
cells in vitro. (a) Representative flow cytometry plots for analyses of infection efficiency
after 10 hours of infection with VSVΔG-SVGmu-GFP or VSVΔG-VSVG-GFP virus in
293T cells expressing human DC-SIGN (293T.hDCSIGN). The infection efficiency was
measured by analyzing GFP expression and the parental 293T cells lacking the expression
of DC-SIGN were included as controls. (b) The ratio of GFP-positive 293T.hDCSIGN cells
to 293T cells upon VSVΔG-VSVG-GFP or VSVΔG-SVGmu-GFP virus infection.
(c) Representative flow cytometry plots for analysis of infection efficiency of VSVΔG-
VSVG-GFP and VSVΔG-SVGmu-GFP viruses on murine BMDCs. (d) The ratio of GFP-
positive CD11c+ BMDCs to CD11c- cells upon infection with VSVΔG-VSVG-GFP and
VSVΔG-SVGmu-GFP viruses.

4.3.3 VSVΔG-SVGmu-GFP infection promotes the maturation of DCs in vitro
We have previously demonstrated that that SVGmu pseudotyped lentivector could
induce DCs activation and maturation (Yang, Yang et al. 2008). This prompts us to
77

investigate whether DC-targeting VSVΔG-SVGmu-GFP virus can also induce DCs
activation and maturation. We first evaluated the expression of maturation markers on
murine DCs infected with rVSVs. Mouse BMDCs were infected with sucrose gradient
purified VSVΔG-SVGmu-GFP and VSVΔG-VSVG-GFP viruses at multiplicities of
infection (MOI) of 0.01, 0.1, or 1 pfu/cell and then cultured for 24 hours. DCs were stained
with the Abs to CD40, CD80, CD86 and MHC II, followed by FACS analysis on CD11c+-
gated cells. As expected, LPS-treated DCs displayed substantial upregulation of the four
maturation markers (Figure 4.3a). Compared with unstimulated cells, VSVΔG-VSVG-
GFP infected DCs displayed elevated levels of CD86, and a marginal increase in CD40
and CD80 but no up-regulation of MHC class II at all MOI (Figure 4.3a and 4.3b). In
contrast, VSVΔG-SVGmu-GFP enhanced DC maturation marker expression as efficient
as LPS and it did so in a dose-dependent manner. VSVΔG-SVGmu-GFP (at 1 pfu/cell)
infected DCs displayed greatly elevated levels of CD86 (10-fold) and CD40 (6-fold), and
a marginal increase in MHC Class II (2.5-fold) and CD80 (2-fold), suggesting that
VSVΔG-SVGmu-GFP infection can efficiently promote DCs maturation.
Another important property associated with DC maturation is the ability to secrete
inflammatory and regulatory cytokines. We next determined the presence of TNF-α, IL-6,
IL-12 and IL-1β in the supernatants of mouse BMDCs infected with rVSVs or treated with
LPS for 24 hours by ELISA. VSVΔG-VSVG-GFP infected DCs produced low amount of
TNF-α, IL-6, and IL-1β at MOI of 0.01, however, the secretion was not further increased
following infection at an MOI of 0.1 or 1 (Figure 4.3c). In contrast, VSVΔG-SVGmu-GFP
infection result production of TNF-α, IL-6, and IL-1β in a dose-dependent manner with
increased MOI. More importantly, VSVΔG-SVGmu-GFP infected DCs produced
78

Figure 4.3 SVGmu bearing rVSV infection induces BMDCs maturation. (a)
Representative flow cytometry plots of CD40, CD80, CD86 and MHC II expression in
LPS-stimulated or rVSV infected DCs. (b) Kinetic analyses of CD40, CD80, CD86 and
MHC II expression in LPS-stimulated or rVSV infected DCs. The relative expression of
CD40, CD80, CD86 and MHC II was depicted as fold induction of the PBS mock
uninfected control. The infection MOI was indicated (*P < 0.05; **P < 0.01, Error bars
79

indicate mean ± s.d.). (c) Production of TNF-α, IL-6, IL-12 and IL-1β by DCs were
determined by ELISA. The supernatants were collected from the LPS-stimulated or rVSV
infected DCs 24 hours after stimulation or infection. Data are representatives of three
independent experiments.

significant higher levels of cytokines than VSVΔG-VSVG-GFP infected DCs an MOI of
1. Both VSVΔG-SVGmu-GFP and VSVΔG-VSVG-GFP infection stimulated IL-12
production, but such production was not further enhanced with increased MOI.

4.3.4 Heterologous prime/boost with DCLV-gp100 and VSVΔG-SVGmu-gp100
induces CD8+ T cell response in mice
Since the DC-targeting rVSV could effectively infect and activate BMDCs in vitro
and LVs have been demonstrated to be excellent delivery vehicles for antigens for
vaccination purposes to elicit effective immune responses. Our previous studies have
shown that LVs targeted to DC-SIGN were more efficient in transducing CD11c+ DC cells
and generating effective immunity than VSVG psuedotyped LVs (Yang, Yang et al. 2008).
We next studied the utility of DC-targeting rVSV in prime/boost immunization protocol.
Three groups of BALB/c mice received either PBS control, DCLV-gp100 prime followed
by a VSVΔG-SVGmu-gp100 boost, or VSVΔG-SVGmu-gp100 prime followed by a
DCLV-gp100 boost. Five days after the rVSV boost and 10 days after DCLV boost, the
splenocytes were harvested and restimulated with gp100 peptide and IFN-γ-producing
CD8+ T cells were quantified by FACS (Figure 4.4a). From the summarized data, an
enhancement of IFN-γ secretion was seen in the rVSV prime/LV boost group with 3-fold
greater responses than LV prime/rVSV boost group (Figure 4.4b).
80

4.3.5 Heterologous prime/boost with DCLV-gp100 and VSVΔG-SVGmu-gp100
inhibit B16 melanoma growth in vivo
We then investigated whether heterologous prime/boost with DCLV-gp100 and
VSVΔG-SVGmu-gp100 could be effective for inhibiting tumor growth in a therapeutic
model. The mice were inoculated of 0.2× 10
6
B16-F10 tumor cells and immunize with
either DCLV-gp100 (10
7
TU) or VSVΔG-SVGmu-gp100 at the same day. Fourteen days
later, the mice received the heterologous boost immunization, the tumor volume was
measured every other day since day 10 post-inoculation (Figure 4.5a). To our surprise, the
tumor-bearing mice therapeutically immunized with LV prime/rVSV

Figure 4.4 Heterologous DCLV prime/rVSV boost immunization elicits CD8+ T cells
response. (a) Representative FACS plots for measuring IFN- γ-secreting CD8+ T cells.
(b) Statistical comparison of IFN-γ response elicited by the two immununization regimen.
81

boost showed significantly slower tumor growth as compared with control group and rVSV
prime/LV boost group (Figure 4.5b). Moreover, the survival study showed that the group
receiving LV prime/rVSV immunizaiton had a median survival of 31 days (the tumor size
of 2000 mm3 was used as a surrogate endpoint of survival), and lived longer than mice
treated with rVSV prime/LV boost, or those in the control group (Figure 4.5c).

Figure 4.5 Heterologous DCLV prime/rVSV boost immunization inhibit tumor
growth. (a) Schematic representation of immunization and analysis protocol. (b) Tumor
growth curves in B16 bearing mice. B6 mice were s.c. challenged with 2× 10
5
of B16-F10
tumor cells at right flank and immunized via f.p. injection of DCLV-gp100 (10
7
TU) or
VSVΔG-SVGmu-gp100 (10
7
pfu). The boost injection was administrate at the indicated
date. Tumor volume was measured every other day and mice consider be dead when the
tumor volume reached around 2000mm
3
. (n = 3-5 mice/group) (c) Mouse survival was
calculated using the Kaplan-Meier method. (Error bars indicate SEM * P<0.05, ** P<0.01)
82

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Abstract (if available)

Abstract In the past two decades, the selective mechanism-based therapeutics had been developed for cancer treatment with an improved understanding of cancer pathogenesis. Targeted approaches can both inhibit molecular pathways, which are critical for tumorigenesis and tumor progression, and also stimulate host immune response against tumor growth. Recently, many investigations have evaluated the effect of the combined therapy with targeting agents and cytotoxic molecules and found effectively improved clinical outcomes. Herein this thesis, I explored the potential of use targeting immunotoxin based on fibroblast activation protein (FAP), a type II transmembrane cell surface serine protease that highly expressed in tumor-associated fibroblasts (TAFs) in most human epithelial cancers, some soft tissue and bone sarcomas, in cancer therapeutics. I evaluated a novel immune-based approach to specifically targeting FAP-expressing TAFs in a mouse 4T1 metastatic breast cancer model. Upon treatment with FAP-targeting immunotoxin, the levels of various growth factors, cytokines and matrix metalloproteinases were changed in the tumor and the recruitment of tumor infiltrating immune cells in the tumor microenvironment was decreased. In addition, combined treatment with FAP-PE38 and cytotoxic agent paclitaxel effectively inhibited tumor growth in vivo. These findings highlight the potential use of TAFs targeting therapy in cancer treatment. Next, I assessed a strategy of utilizing multi-antigens vaccine by administration of three melanoma-associated antigens (gp100, TRP1, TRP2) in lentiviral vectors (termed LV-3Ag). The LV-3Ag immunization dramatically increased functional T cell infiltration into tumors and generated protective and therapeutic antitumor immunity. I also studied the combination therapy with aFAP-PE38 and LV-3Ag and they exhibited significantly enhanced antitumor effects on tumor growth in established B16 melanoma model. The mechanism likely involves in modulation of immune suppressive tumor microenvironment and, consequently, activation of cytotoxic CD8+ T cells, which are capable of specifically recognizing and destroying tumor cells. In my third study, I attempted to use dendritic cells (DCs) targeting vesicular stomatitis virus VSV), which displays engineered sindbis virus glycoprotein mutant (SVGmu) on its surface to selectively binding to the DC-SIGN protein, to express a mouse melanoma antigen (gp100) in cancer therapy. This recombinant rVSV specifically infected DCs and stimulateed their activation and maturation in vitro. A heterologous DCLV-gp100 prime/rVSV-Gag boost immunization elicited gp100-specific CD8 T cell response, therefore greatly inhibiting the tumor growth and prolong the medium survival. These findings suggest that DC-specific rVSV vectors encoding tumor antigens can be an effective therapeutic agents and also potentiate the advantage of heterologous immunization regimen with lentivirus to achieve more effective immune response against cancer.

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Asset Metadata

Creator Fang, Jinxu (author)

Core Title Engineering immunotoxin and viral vectors for cancer therapy

School Viterbi School of Engineering

Degree Doctor of Philosophy

Degree Program Chemical Engineering

Publication Date 10/20/2017

Defense Date 08/17/2015

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

Tag breast cancer,cancer vaccine,fibroblast activation protein,immunotoxin,OAI-PMH Harvest,tumor microenvironment,tumor-associated fibroblast

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Wang, Pin (committee chair), Malmstadt, Noah (committee member), Pratt, Matthew Robert (committee member)

Creator Email jinxufan@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c40-191996

Unique identifier UC11276487

Identifier etd-FangJinxu-3985.pdf (filename),usctheses-c40-191996 (legacy record id)

Legacy Identifier etd-FangJinxu-3985.pdf

Dmrecord 191996

Document Type Dissertation

Format application/pdf (imt)

Rights Fang, Jinxu

Type texts

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

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

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA