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Discovery of novel small molecules for ovarian cancer treatment

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Discovery of novel small molecules for ovarian cancer treatment

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DISCOVERY OF NOVEL SMALL MOLECULES FOR OVARIAN CANCER
TREATMENT

by

Shili Xu

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
(PHARMACOLOGY AND PHARMACEUTICAL SCIENCES)

May 2013

Copyright 2013 Shili Xu
ii
DEDICATION

For my parents, my in-laws and my wife for their unconditional love and constant
support throughout the course of my research.
iii
ACKNOWLEDGMENTS
I would like to express my forever gratitude to my advisor and my Committee Chair,
Prof. Nouri Neamati for giving me the chance to study and explore the power of small-
molecule compounds for cancer treatment in his lab, and providing me with invaluable
guidance, encouragement, inspiration and constant support during my Ph.D. study at the
University of Southern California. His mentorship significantly shaped my scientific
career and my desire to make cancer a disease of the past.
I am also sincerely grateful to Dr. Roger F. Duncan, Dr. Ebrahim Zandi, Dr. Bangyan
Stiles and Dr. J. Andrew MacKay for their advice and serving as my Qualifying Exam
and Dissertation Committees. Especially, I need to thank Dr. Roger F. Duncan for
teaching me sophisticated experimental skills as well as the unbiased interpretation of
facts.
Special thanks are conveyed to past and current members in Prof. Neamati’s group,
including Dr. Xuefei Cao, Dr. Roppei Yamada, Dr. Fedora Grande, Dr. Hiroyuki Otake,
Dr. Bikash Debnath, Dr. Tino Sanchez, Dr. Srinivas Odde, Dr. Rambabu Gundla, Dr.
Shuzo Tamura, Dr. Yunma Shabaik, Dr. Erik Serrao, Dr. Melissa Millard, Helen Ha,
Kavya Ramkumar, Divya Pathania, Si Li, Yuting Kuang and Saranya Sankar for their
everyday support and inspiring discussions. There were numerous enjoyable and
memorable moments we had together.
My final and most heartfelt acknowledgement must go to my parents, my in-laws and
my wife for their love and encouragement. Without them, all my discoveries and
achievements would never happen.
iv
TABLE OF CONTENTS
ACKNOWLEDGMENTS ....................................................................................... iii
ABSTRACT ........................................................................................................... ix
CHAPTER I: OVARIAN CANCER AND SMALL-MOLECULE INHIBITORS ......... 1
CHAPTER II: PDI IS A PROMISING DRUG TARGET FOR CANCER THERAPY
............................................................................................................................... 4
II-1: INTRODUCTION ........................................................................................ 5
II-2: STRUCTURAL PROPERTIES OF PDI FOR ITS BIOCHEMICAL
ACTIVITIES ........................................................................................................ 8
II-3: THE THIOL-DISULFIDE EXCHANGE REACTIONS OF PDI ................... 12
II-4: PDI IS A POTENTIAL DRUG TARGET FOR CANCER TREATMENT .... 16
II-4-1: PDI Is Highly Expressed in Cancer Tissues ...................................... 17
II-4-2: PDI Is Associated with Clinical Outcomes of Cancer Therapy .......... 20
II-4-3: PDI Supports Tumor Survival and Progression ................................. 22
II-5: APPROACHES TO MEASURING PDI ACTIVITIES AND SCREENING
FOR PDI INHIBITORS ..................................................................................... 24
II-5-1. Reductase Assays ............................................................................. 25
II-5-2: Oxidation Assays ............................................................................... 28
II-5-3: Isomerization Assays ......................................................................... 30
II-5-4: Protein Chaperone Assays ................................................................ 33
II-6: SMALL-MOLECULE INHIBITOR OF PDI ................................................. 36
II-6-1: Synthetic Compounds ........................................................................ 37
II-6-2: Plant Metabolites ............................................................................... 43
II-6-3: Antibiotics ........................................................................................... 46
II-6-4: Hormones ........................................................................................... 50
II-6-5: Xenoestrogens ................................................................................... 53
II-7: CONCLUDING REMARKS ....................................................................... 55
CHAPTER III: DISCOVERY OF AN ORALLY ACTIVE IRREVERSIBLE
INHIBITOR OF PROTEIN DISULFIDE ISOMERASE ......................................... 57
III-1: RESULTS ................................................................................................ 58
III-1-1: Novel PACMAs Demonstrate Cytotoxicity in a Panel of Ovarian
Cancer Cell Lines. ........................................................................................ 58
III-1-2: Structure-Activity Relationships for New PACMAs and Related
Compounds .................................................................................................. 61
III-1-3: Active PACMA Analoes Covalently Bind to Their Cellular Target
Protein in Human Ovarian Cancer Cells ...................................................... 63
III-1-4: Identification of PDI as a Target of PACMAs .................................... 66
III-1-5: Active PACMAs Affect PDI Secondary Structure and Inhibit PDI
Activity .......................................................................................................... 73
III-1-6: PACMA 31 Suppresses Tumor Growth in Human Ovarian Cancer
Mouse Xenografts. ....................................................................................... 78
III-2: DISCUSSION ........................................................................................... 84
v
III-3: EXPERIMENTAL PROCEDURES ........................................................... 87
CHAPTER IV: GP130 IS A PROMISING TARGET FOR CANCER
CHEMOTHERAPY .............................................................................................. 98
IV-1: INTRODUCTION ..................................................................................... 99
IV-2: GP130 STRUCTURE ............................................................................ 101
IV-3: THE EXPRESSION OF GP130 CYTOKINES AND COMPONENTS OF
THE GP130-RELATED RECEPTOR COMPLEXES ..................................... 107
IV-4: GP130 SIGNALING IN PHYSIOLOGICAL PROCESSES .................... 112
IV-5: DYSREGULATED GP130-MEDIATED SIGNALING IN CANCER ........ 117
IV-6: GP130 IN OTHER DISEASES AND ITS THERAPEUTIC IMPLICATIONS
....................................................................................................................... 138
IV-7: CONCLUDING REMARKS .................................................................... 141
CHAPTER V: DISCOVERY OF AN ORALLY ACTIVE INHIBITOR OF GP130 142
V-1: RESULTS ............................................................................................... 143
V-1-1: SC144 Exhibits Anticancer Activity in a Panel of Human Ovarian
Cancer Cell Lines ....................................................................................... 143
V-1-2: SC144 Induces gp130 (Ser782) Phosphorylation and Down-
Regulates gp130 Glycosylation .................................................................. 146
V-1-3: SC144 Decreases gp130 Protein Levels on Cell Surface ............... 148
V-1-4: SC144 Directly Binds gp130 ............................................................ 150
V-1-5: SC144 Suppresses the Activation of Stat3 ...................................... 152
V-1-6: SC144 Decreases the Expression of gp130/Stat3 Target Genes ... 154
V-1-7: SC144 Potently Blocks Cytokine-Triggered gp130 Signaling .......... 156
V-1-8: Inhibition of gp130/Stat3 Leads to Cytotoxicity ................................ 161
V-1-9: SC144 and Its Active Analogs Inhibit gp130/Stat3 Signaling and Show
Cytotoxicity in Other Cancer Types ............................................................ 163
V-1-10: SC144 Suppresses Tumor Growth in Human Ovarian Cancer
Xenografts .................................................................................................. 167
V-2: DISCUSSION ......................................................................................... 174
V-3: MATERIALS AND METHODS ................................................................ 179
CHAPTER VI: FUTURE WORK ........................................................................ 188
VI-1: Is Cancer a Curable Disease? .............................................................. 188
VI-2: Destroy, Control or Reprogram ............................................................. 189
VI-3: Dark Matters in Cancer.......................................................................... 189
VI-4: Effects of PACMAs on PDI .................................................................... 190
VI-5: Other Targets of PACMAs ..................................................................... 190
VI-6: Toxicity Associated with PDI Inhibition .................................................. 191
VI-7: Other Potential Targets of SC144 ......................................................... 191
REFERENCES .................................................................................................. 194

vi
LIST OF TABLES
Table 1. Symptoms of ovarian cancer…………...............…………………………..1
Table 2. Mechanisms of actions of paclitaxel and carboplatin…………...………...2
Table 3. Currently available structures of isolated domains in PDI……………....10
Table 4. Cytotoxicity of select PMCMAs in a panel of human ovarian cancer
cells….………........……..…………………..…………...............…………………...60
Table 5. Active PACMAs affected PDI secondary structure.……….............……73
Table 6. Available crystal and NMR structures of gp130…...………..……….....102
Table 7. Antagonists against cytokine ligands or subunits of gp130 receptor
complexes in clinical trials………………….................................................……140
Table 8. Cytotoxicity of SC144 in a panel of human ovarian cancer cell lines...144
Table 9. Cytotoxicity of stattic in a panel of human ovarian cancer cell lines....162
Table 10. Cytotoxicity of SC144 and its analogs in a panel of cancer cell lines
.………........……..…………………..…………..................................……………164

vii
LIST OF FIGURES
Figure 1. Structural properties of PDI ……….................…………………............11
Figure 2. In vitro and in vivo biochemical reactions involving PDI……................15
Figure 3. PDI is highly expressed in multiple cancer types compared to
respective normal tissues……………...……..............…………............................19
Figure 4. PDI expression is associated with glioblastoma patients’ overall survival
rate. …………...……..............………….......…..........…........................................21
Figure 5. PDI activity assays ………………………………………….....................35
Figure 6. Chemical structures of PDI inhibitors from synthetic compounds….....42
Figure 7. Chemical structures of PDI inhibitors from plant metabolites, antibiotics,
hormones and xenoestrogens…........................…………...................................54
Figure 8. PACMA analogs used in this study ……………………….....................59
Figure 9. BODIPY conjugation of PACMA 31……………………...…....…...........65
Figure 10. PACMA 57 covalently binds to PDI……………………....……............67
Figure 11. Confirmation of PDI as the cellular target of PACMAs……….............69
Figure 12. Determination of PACMA 31 binding site in PDI………………...........71
Figure 13. PACMA 31 covalently binds to Cys397/Cys400 in PDI active site
......................………........................................…………......................................72
Figure 14. Active PACMAs alters PDI secondary structures…………….............74
Figure 15. Active PACMAs inhibit PDI activity…………………..........……...........75
Figure 16. Silencing of PDI inhibits cell growth of OVCAR-8 cells.......................77
Figure 17. PACMA 57 targets in vivo ovarian tumor via binding PDI……...........79
Figure 18. PACMA 31 suppresses tumor growth in a mouse xenograft model of
human OVCAR-8 ovarian cancer…………………………………..........................82
Figure 19. Histochemistry analysis of organs from OVCAR-8 xenograft mice in
the control group and in PACMA 31 treatment groups…………..........................83
Figure 20. Composition of gp130 protein and gp130-associated receptor
complexes. .......................................................................………………………106
Figure 21. Expression levels of gp130 in different tissues………………...........109
Figure 22. Expression levels of gp130 cytokines and other gp130 receptor
complex subunits in different tissues...............................………..…......…........110
Figure 23. gp130-mediated signaling pathways and physiological processes..116
Figure 24. Dysregulation of the expression of gp130 in cancers……................118
Figure 25. SC144 exhibits cytotoxicity in human ovarian cancer cell lines…....145
viii
Figure 26. SC144 induces gp130 phosphorylation and deglycosylation………147
Figure 27. SC144 decreases gp130 cell surface expression………..................149
Figure 28. SC144 directly binds gp130………………………………..........….....151
Figure 29. SC144 inhibits constitutive Stat3 phosphorylation in ovarian cancer
cells....................................................................................................................153
Figure 30. SC144 suppresses the expression of gp130/Stat3 target genes…..155
Figure 31. SC144 inhibits downstream signaling induced by gp130 cytokines.157
Figure 32. SC144 inhibits nuclear translocation of Stat3 induced by gp130
cytokines............................................................................................................159
Figure 33. SC144 inhibits nuclear translocation of Stat1 induced by gp130
cytokines............................................................................................................160
Figure 34. gp130/Stat3 inhibition causes cytotoxicity in human ovarian cancer
cells....................................................................................................................162
Figure 35. Effects of SC144 and its analogs on the abilities of a panel of cancer
cells to form colonies.…………...…………..........…………................................165
Figure 36. Effects of SC144 and its analogs on the activity of the gp130/Stat3
signaling.............................................................................................................166
Figure 37. SC144 suppresses human ovarian cancer xenograft in nude mice.168
Figure 38. SC144 treatment reduced tumor blood vessels in OVCAR-8 xenograft
mice................................................................................................................…170
Figure 39. SC144 treatment induces extensive areas of necrosis in tumor but no
damage in normal tissue.……................……………..........................................171
Figure 40. Effects of SC144 on in vivo ovarian tumor.…...................................173
Figure 41. SC144 has no significant effects on HDAC activity..........................176
Figure 42. A working model for the anticancer mechanism of SC144 in ovarian
cancer................................................................................................................178

ix
ABSTRACT
Ovarian cancer is one of the leading cancers causing women’s death in the United States
mainly due to its late diagnosis and resistance to current clinically used drugs. Therefore,
the development of novel potent drugs for the treatment of ovarian cancer is in urgent
medical need.
Protein disulfide isomerase (PDI), an endoplasmic reticulum (ER) chaperone protein,
catalyzes disulfide bond breakage, formation and rearrangement. The effect of PDI
inhibition on ovarian cancer progression is not yet clear, and there is a dearth of potent,
selective and safe small-molecule inhibitors of PDI. In chapter III, I report a novel class
of propynoic acid carbamoyl methyl amides (PACMAs) that are active against a panel of
human ovarian cancer cell lines. Using fluorescent derivatives, 2D gel electrophoresis,
and mass spectrometry, we established that PACMA 31, one of the most active analogs,
act as irreversible small-molecule inhibitors of PDI. We also demonstrated that PDI is
essential for the survival and proliferation of human ovarian cancer cells. In vivo
PACMA 31 showed tumor targeting ability and significantly suppressed ovarian tumor
growth without causing damage in normal tissues. These irreversible small-molecule PDI
inhibitors represent a new approach for the development of targeted anticancer agents for
ovarian cancer therapy, and can also serve as useful probes for investigating the biology
of PDI-implicated pathways.
On the other hand, despite abundant evidence that ovarian cancer progression is
dependent upon IL-6/Stat3 signaling, the role of glycoprotein 130 (gp130), the signal
transducer of this signaling axis, is unclear in ovarian cancer, and there is a dearth of
small-molecule inhibitors of gp130. In chapter V, I report that gp130 is an attractive drug
x
target in ovarian cancer due to its role in promoting ovarian cancer progression via the
activation of its downstream Stat3 signaling, and identify a small-molecule gp130
inhibitor SC144 with anticancer activity. In vitro, SC144 exhibits potency in human
ovarian cancer cells without cytotoxicity in normal epithelial cells. SC144 binds gp130,
induces gp130 phosphorylation (S782) and deglycosylation, abrogates Stat3
phosphorylation and nuclear translocation, and thus inhibits the expression of
downstream target genes. In addition, SC144 shows potent inhibition of gp130 ligand-
triggered signaling. In vivo, SC144 suppresses tumor growth with oral bioavailability in a
mouse xenograft model of human ovarian cancer without causing damage to normal
tissues.
My discovery of PDI and gp130 as potential drug targets for ovarian cancer treatment,
and the identification of PACMA 31 and SC144 as inhibitors of PDI and gp130,
respectively, not only promote the development of targeted therapy, but also bring new
hope for cancer patients.

1

CHAPTER I: OVARIAN CANCER AND SMALL-
MOLECULE INHIBITORS
Ovarian cancer is the fourth-leading cause of death in women with gynecological
diseases in the United States. There are 22,280 estimated new cases and
15,500 estimated deaths in 2012. Ovarian cancer has been regarded as the
“silent killer” because the disease usually shows no obvious symptoms in its
early phase. As the cancer grows, symptoms become apparent (Table 1).
However, these symptoms are not specific to the disease, and are more likely
due to causes other than ovarian cancer, resulting in frequent delay in diagnosis
of ovarian cancer. In fact, about 70% of the ovarian cancer cases are diagnosed
at the late and distant stage, and therefore are poorly treatable (Siegel et al.,
2012).
Table 1. Symptoms of ovarian cancer
Common symptoms Less common symptoms
Pressure or pain in the abdomen, pelvis, back,
or legs
Shortness of breath
A swollen or bloated abdomen Feeling the need to urinate often
Nausea, indigestion, gas, constipation, or
diarrhea
Unusual vaginal bleeding (heavy
periods, or bleeding after menopause)
Feeling very tired all the time

While most of ovarian cancers occur sporadically with unclear causes,
inherited BRCA1 and BRCA2 mutations account for 10 to 15 percent of ovarian
2
cancers in the United States (Campeau et al., 2008). Collaborative reanalysis of
data from a large-sample study has shown that oral contraceptives significantly
reduced the risk of ovarian cancer, and conferred protection for years to even
decades after oral contraceptive use had ceased (Beral et al., 2008). Therefore,
oral contraceptives seem to be an effective preventative approach against
ovarian cancer.
Currently, the standard treatment for ovarian cancer is defined by the delivery
of cytotoxic chemotherapeutic agents paclitaxel and carboplatin following
aggressive surgical cytoreduction. The chemical structures and mechanisms of
these chemotherapeutic agents are shown in Table 2. Although they are usually
effective at the beginning, almost all ovarian cancer patients develop drug
resistance after prolonged treatment with these standard agents, resulting in
ovarian cancer relapse and eventually the death of patients (Petty et al., 1998).
In this context, there is an urgent need for breakthrough drugs with effective
therapeutic impact on ovarian cancer.
Table 2. Mechanisms of actions of paclitaxel and carboplatin
Drug Chemical structure Mechanism of action
Paclitaxel

Paclitaxel binds β tubulin, stabilizes the
microtubule polymer, prevents microtubule from
disassembly, and thus blocks mitosis and
induces apoptosis of proliferating cells.
Carboplatin

Like cisplatin, carboplatin crosslinks DNA and
interferes with cell division by mitosis. The
damaged DNA further activates apoptosis.
Unlike cisplatin, carboplatin may have other
biological mechanisms.

O NH O
O
OH
HO
O O
O
OH
O
O
O
O
O
Pt
O
O
NH
3
NH
3
3
Small-molecule drugs and protein-based biological drugs (biologics) are the
two classes of therapeutic agents. Small-molecule drugs make up over 90
percent of the drugs on the market today. My research focuses on the
development of novel small-molecule inhibitors (average molecular weight < 500)
against proteins essential for ovarian cancer development. Compared to
biologics, small molecules have the following advantages: easy synthesis, high
stability, low cost, orally bioavailability, and ability to target both extracellular and
intracellular molecules. Generally there are two classes of small-molecule drugs:
reversible and irreversible agents. Most small-molecule drugs are reversible
drugs, which interact with their targets via non-covalent binding; whereas
irreversible drugs derive part of their affinity by forming covalent bonds with their
targets (Singh et al., 2011). Herein, I report my successful development of an
irreversible inhibitor of protein disulfide isomerase (PDI) and a reversible inhibitor
of glycoprotein 130 (gp130) for the treatment of ovarian cancer.

4
CHAPTER II: PDI IS A PROMISING DRUG TARGET FOR
CANCER THERAPY
Accumulating evidence about the involvement of PDI in multiple diseases has led
to discovery of a few potential small-molecule inhibitors. Recent studies have
indicated PDI as a potential chemotherapeutic target for several different cancer
types, suggesting that currently available PDI inhibitors may exhibit potency for
these cancer types. In this chapter, I will introduce current knowledge about the
relationship between PDI and cancer, not only based on previously published
results, but also on our extensive exploration of available data sets from gene
expression studies. I will also introduce strategies for discovery of PDI inhibitors,
and previously documented inhibitors of PDI.

5
II-‐1:
INTRODUCTION

PDI is a 57 kDa dithiol-disulfide oxidoreductase chaperone. Mainly located in
endoplasmic reticulum (ER), PDI is one of the most abundant soluble ER
proteins, and accounts for up to 0.8% of total cellular protein (Ferrari and Soling,
1999). PDI is the first protein-folding catalyst, independently discovered by two
research groups in 1963 (Goldberger et al., 1963; Venetianer and Straub, 1963).
The group of Brunó Straub, who later became the president of Hungary,
discovered PDI from pigeon and chicken pancreas for its ability to oxidize
reduced ribonuclease (Venetianer and Straub, 1963). During the same period,
PDI was discovered in rat liver homogenate to activate ribonuclease by the group
of Christian B. Anfinsen (Goldberger et al., 1963), who later shared the 1972
Nobel Prize in Chemistry for his work on ribonuclease. A decade later its official
name, PDI was used for the first time (Hawkins and Freedman, 1975).
Subsequently, at least 19 other members have been discovered, forming the PDI
protein family (Figure 1A). Different aspects of the biochemistry of the PDI family
proteins have been previously reviewed (Hatahet and Ruddock, 2009; Kozlov et
al., 2010).
In the ER, PDI plays a central role as a reductase, an oxidase, an isomerase
and a chaperone. PDI mediates oxidative protein folding via its chaperone
protein activity as well as its ability to catalyze disulfide bond oxidation
(formation), reduction (breakage) and isomerization (rearrangement) in all
nonnative protein and peptide substrates. In addition, PDI mediates peptide
loading to MHC I (Peaper and Cresswell, 2008), and regulates NAD(P)H oxidase
6
activity (Janiszewski et al., 2005) in the ER. PDI is also a subunit of proly-4
hydroxylase (an essential enzyme for the synthesis of collagens) (Koivu et al.,
1987) and microsomal triglyceride transfer protein (a central enzyme for the
assembly of apoB-containing lipoproteins) (Wetterau et al., 1991). To date, no
PDI global knockout mouse has been reported. Considering the essential role of
PDI, such an absence implies that PDI knockout strains may not be viable.
Dysregulation of PDI expression and/or enzymatic activity is associated with a
series of diseases (Benham, 2012), such as neurodegenerative (Hoffstrom et al.,
2010; Hoozemans et al., 2007; Uehara et al., 2006; Unterberger et al., 2006) and
cardiovascular diseases (Laurindo et al., 2008; Severino et al., 2007; Shibata et
al., 2001; Sun et al., 2007). Despite locating primarily in the ER as a soluble
oxidoreductase, PDI is also present on the extracellular side of the plasma
membrane (Donoghue et al., 2000; Jiang et al., 1999). Although how PDI is
secreted or translocated to the cell surface remains unclear, some evidence
suggests that it is attached to the cell membrane via electrostatic charges
(Terada et al., 1995). PDI functions as a reductase (Bi et al., 2011) as well as an
isomerase (Willems et al., 2010) on the cell surface. Cell surface PDI regulates
multiple important biological processes, including coagulation (Popescu et al.,
2010), injury response (Reinhardt et al., 2008), platelet activation (Essex and Li,
1999; Essex et al., 2001; Lahav et al., 2003) and thrombus formation (Cho et al.,
2008), T-cell migration (Bi et al., 2011), glioma cell migration (Goplen et al.,
2006), gamete fusion (Jain et al., 2007), and nitric oxide internalization from
extracellular S-nitrosothiols (Ramachandran et al., 2001). Importantly, cell
7
surface PDI facilitates viral infection (Stolf et al., 2011). An example is its
involvement in HIV-1 fusogenic events. Cell surface PDI catalyzes the reduction
of at least two disulfide bonds in gp120, an HIV-1 envelope glycoprotein,
resulting in a major conformational change in gp120 that enhances its binding
with the co-receptors CXCR4 and CCR5 (Barbouche et al., 2003; Gallina et al.,
2002; Ryser and Fluckiger, 2005). Besides ER and cell surface, PDI has also
been reported at other subcellular locations, including cytoplasm, mitochondria
and nucleus (Rigobello et al., 2001; Turano et al., 2002). However, these
observations are not conclusive and the biological functions of PDI at these
distinct locations remain unclear.
Increasing knowledge on the involvement of PDI in multiple diseases has led
to development of potential small-molecule inhibitors as well as robust activity
assays for screening PDI inhibitors. Recent studies have implicated PDI as a
novel and promising chemotherapeutic target for several different types of
cancer. Herein, I discuss current knowledge on the relationship between PDI and
cancer, various assays used for the discovery of PDI inhibitors, and currently
available PDI inhibitors.

8
II-‐2:
STRUCTURAL
PROPERTIES
OF
PDI
FOR
ITS
BIOCHEMICAL
ACTIVITIES

Encoded by the P4HB gene, PDI has a multi-domain structure (Gruber et al.,
2006). The full-length PDI contains 508 amino acids, whereas the mature form
lacking the signal sequence of the first 17 amino acids in the N-terminus,
contains 491 amino acids. PDI has four distinct domains, a, b, b’, and a’, with a
highly acidic C-terminal extension c and a b’-a’ linker x (Figure 1B). A classic
KDEL ER-retrieval signal lies at the C-terminal of c, making PDI mainly an ER-
resident protein. Although the structure of the full-length human PDI has not yet
been resolved, structures of isolated domains are available (Table 3) (Denisov et
al., 2009; Kemmink et al., 1996; Kemmink et al., 1999; Nguyen et al., 2008;
Wang et al., 2012). These structures not only shed light on the structure-activity
relationship in PDI, but also provide important tools for the design and discovery
of PDI inhibitors.
The a and a’ domains are active domains, sharing 33.6% identity with each
other (Figure 1C). Each of the two domains contains an identical active site Cys-
Gly-His-Cys (Figure 1B). PDI’s reductase, oxidase, and isomerase activities rely
on the thiol groups of these active-site cysteines (Ellgaard and Ruddock, 2005;
Hatahet and Ruddock, 2007). The a and a’ domains operate independently of
one another, as disruption of active site cysteines in either domain abolished
50% of the catalytic activity of PDI, and disruption of cysteines in both domains
completely abolished PDI’s oxidoreductase activity (Vuori et al., 1992). In each
thioredoxin-like domain, however, the two active-site cysteines contribute
differently to PDI’s enzymatic functions. Being at the N-terminus of the α2 helix,
9
the N-terminal active-site cysteine positions on the surface of PDI with its thiol
group accessible for redox reactions, whereas the C-terminal active-site cysteine
has limited solvent exposure (Figure 1D). In addition, the pKa of the two
cysteines are substantially different. The N-terminal cysteine has a pKa in the
range of 4.5 to 5.6 (Kortemme et al., 1996; Ruddock et al., 1996), whereas the
pKa of the C-terminal cysteine was calculated to be 12.8 (Lappi et al., 2004).
However, it has been indicated that during the biochemical reaction involving PDI
there is a conformational change within the a domains that causes the pKa of the
C-terminal cysteine to shift from 12.8 to 6.1 (Karala et al., 2010; Lappi et al.,
2004). In the initial step of a reaction, the N-terminal cysteine forms a transient
disulfide bond with a cysteine residue in a substrate, resulting in a heterodimer.
Subsequently the C-terminal cysteine attacks the N-terminal cysteine to release
the substrate, an event called the “escape pathway” (Walker and Gilbert, 1997).
Compared to the a and a’ domains, the b and b’ domains share substantially
less identity (16.5%) in the sequence with each other (Figure 1E). Each of the
two b domains exhibits a thioredoxin-like fold, β-α-β-α-β-α-β-β-α (Kemmink et al.,
1997), whereas they lack the active sites. Although all PDI domains contribute to
the binding with misfolded proteins, the b’ domain has been shown to provide the
principal substrate-binding site of PDI, with high affinity but board specificity
(Klappa et al., 1998). Via hydrophobic interaction (Karala and Ruddock, 2010),
the b’ domain is essential and sufficient for small peptide binding, whereas in the
case of binding large peptides it is essential but not sufficient (Klappa et al.,
1998). This substrate-binding site is also necessary for the interaction of PDI with
10
the α-subunite of prolyl-4-hydroxylase. It has been reported that the minimum
sequence requirement for PDI to function as a subunit of prolyl-4-hydroxylase is
fulfilled by the b’a’ construct, whereas the a and b domains enhance such an
assembly (Pirneskoski et al., 2001). In addition, the b’ domain is essential for the
chaperone activity of PDI that can be inhibited by a peptide substrate for the b’
domain (Quan et al., 1995).

Table 3. Currently available structures of isolated domains in PDI
PDB ID Structure Domain Method Year Ref
3UEM

b-b’-x-a’ X-ray (2.29 Å) 2011 (Wang et al., 2012)
2K18

b-b’ NMR 2008 (Denisov et al., 2009)
3BJ5

b’-x X-ray (2.2 Å) 2007 (Nguyen et al., 2008)
1X5C

a’ NMR 2005 N/A
2BJX

b NMR 1998 (Kemmink et al., 1999)
1MEK

a NMR 1996 (Kemmink et al., 1996)
NA: not applicable.

11

Figure 1. Structural properties of PDI. (A) Protein members of the PDI family. Amino
acid sequences of the active sites are shown. (B) Domain architecture of PDI. (C)
Sequence comparison of the a and a’ domains. Sequence comparison was performed
using STRETCHER (EMBOSS, Pasteur: http://mobyle.pasteur.fr). The active site CGHC
is highlighted. (D) Accessibility of the N-terminal and the C-terminal cysteines in the PDI
active site. Structural information was obtained from 1X5C, and analyzed using Chimera
1.7 (UCSF). (E) Sequence comparison of the b and b’ domains.

12
II-‐3:
THE
THIOL-‐DISULFIDE
EXCHANGE
REACTIONS
OF
PDI

It is well established that PDI is able to act as an oxidase, a reductase and an
isomerase, depending on the redox state of PDI’s active-site cysteines and the
properties of the substrate protein or peptide. In an oxidation reaction, a
substrate dithiol is oxidized to a disulfide in parallel with the reduction of the
active-site disulfide in PDI to the dithiol state. Subsequently, oxidants such as
glutathione disulfide (GSSG) act as terminal electron acceptors to oxidize PDI’s
dithiol back to the disulfide state to complete the catalytic cycle (Figure 2A). In a
reduction reaction, a substrate disulfide is reduced to the dithiol state with the
concomitant formation of an active-site disulfide in PDI. To fulfill the catalytic
cycle, reductants such as GSH, NADPH, and DTT serve as terminal electron
donors to reduce PDI’s disulfide back to its dithiol state (Figure 2B). In an
isomerization reaction, where no additional redox reagents are required, PDI
catalyzes a shift of the disulfide-bond position among the substrate cysteines,
without a net change in the number of disulfide bonds in the substrate or in the
redox state of the active site of PDI (Figure 2C).
In normal cells, and under physiological conditions, PDI activity is tightly
regulated. When PDI catalyzes formation of disulfide bonds in nascent proteins
to mediate oxidative protein folding, the active-site disulfides in PDI are reduced
that have to be efficiently oxidized to reactivate PDI. Several biochemical
reactions are known to oxidize the active sites of PDI (Figure 2D). In the past,
GSSG was considered to be a major regulator in oxidizing the active sites of PDI.
However, this consensus was modified by the discovery of an ER flavor-oxidase
13
Ero1 (endoplasmic reticulum oxidoreductin 1) that plays a central in catalyzing
PDI oxidation (Appenzeller-Herzog et al., 2010; Mezghrani et al., 2001).
Presented as two isoforms (Ero1α and Ero1β) in mammalian cells (Cabibbo et
al., 2000; Pagani et al., 2000), Ero1 preferentially interacts and reacts with
reduced PDI, especially the a’ domain, oxidizing PDI active-site dithiols to
disulfides (Wang et al., 2009b). However, as part of this reaction, Ero1 uses
molecular oxygen as an electron acceptor. As a result, Ero1 produces one
molecule of hydrogen peroxide (H
2
O
2
) when it introduces one disulfide bond in
PDI (Gross et al., 2006). Due to such a potential lethal side reaction and also the
required existence of reduced PDI in the ER for disulfide idomerization, the
activity of Ero1 must be strictly regulated. Oxidation of PDI’s active-site disulfide
is also contributed by other mechanisms, such as H
2
O
2
, Prdx4, DHA, and vitamin
K, and has been recently reviewed in depth (Hatahet and Ruddock, 2009;
Laurindo et al., 2012). In addition, S-glutathionylation of the active-site cysteins in
PDI has been reported to impair PDI activity (Townsend et al., 2009).
Efficient regulation of PDI activity is critical for almost all cellular functions and
even cell survival. Disulfide bond formation occurs in about 30% of proteins, and
is essential for the their biogenesis. Because PDI serves as one of the most
abundant and essential enzymes for protein folding via catalyzing disulfide bond
formation and isomerization, dysfunction of PDI results in rapid accumulation of
unfolded and misfolded proteins in the ER lumen. With hydrophobic amino acid
side chains exposed to the surface, these proteins form insoluble aggregates and
trigger unfolded protein response (UPR) and ER stress (Hotamisligil, 2010;
14
Schroder and Kaufman, 2005). Subsequently three signal-transducing proteins,
PERK, IRE1 and ATF6 positioned on the ER membrane and act as ER stress
sensors, are activated to modulate the UPR. PERK activation immediately leads
to global translational attenuation through direct phosphorylation of elF2, the
regulating initiator of mRNA translation, resulting in a decrease of protein influx
into the ER lumen (Harding et al., 2000). In addition, phospho-eIF2α also up-
regulates the translation of ATF4, increasing its protein abundance (Harding et
al., 2000). Activated IRE1, a site-specific endonuclease, stabilizes XBP-1 mRNA
by direct removal of a small intron, leading to upregulation of XBP-1 protein
levels (Yoshida et al., 2001). ATF6, when triggered by UPR, translocates to the
Golgi apparatus for cleavage to yield an active fragment, ATF6 p50 (Schindler
and Schekman, 2009). ATF6 p50, XBP-1 and ATF4 further enter nucleus, bind to
ERSE promoters, and activate the expression of proteins involved in UPR
regulation. However, when the imbalance of cellular homeostasis exceeds a
cell’s ability to restore, cell death occurs.
15

Figure 2. In vitro and in vivo biochemical reactions involving PDI. PDI catalyzes (A)
oxidation, (B) reduction and (C) isomerization reactions in vitro. In the oxidation reaction, GSSG
represents the terminal electron acceptors, and in the reduction reaction, GSH represents the
terminal electron donors. No extra electron acceptor or donor is needed for the isomerization
reaction that is initiated by the reduced form of PDI. (D) In ER, PDI mainly catalyzes oxidation
and isomerization reactions, mediating disulfide bond formation and rearrangement for oxidative
protein folding. While catalyzing disulfide bond formation in a substrate, PDI’s active-site
cysteines are reduced and efficiently re-oxidized. Impairment of PDI activity leads to
accumulation of unfolded and misfolded proteins, causing UPR and ER stress.
16
II-‐4:
PDI
IS
A
POTENTIAL
DRUG
TARGET
FOR
CANCER
TREATMENT

Although PDI has been extensively studied during the past few decades, its role
in cancer progression is not well established. Therefore, limited knowledge about
the relationship between PDI and cancer is currently available. However, the
published data strongly suggest that PDI is significantly associated with cancer
progression, and is a potential drug target for cancer treatment.
17
II-4-1: PDI Is Highly Expressed in Cancer Tissues
Gene expression microarray studies provide an important tool for assessing PDI
levels in different cancer types. By analyzing published microarray data sets, we
compared PDI expression in different cancer types with that in normal tissues.
We found that PDI expression is significantly up-regulated (P < 10
-5
, fold change
> 2) in brain and CNS cancers (Figure 3A) (Bredel et al., 2005; 2008; Gutmann et
al., 2002; Rickman et al., 2001; Shai et al., 2003; Sun et al., 2006), lymphoma
(Figure 3B) (Basso et al., 2005; Compagno et al., 2009; Piccaluga et al., 2007),
kidney (Figure 3C) (Beroukhim et al., 2009; Jones et al., 2005; Yusenko et al.,
2009), prostate (Figure 3D) (Singh et al., 2002; Welsh et al., 2001), lung (Figure
3E) (Beer et al., 2002), ovarian (Figure 3F) (Bonome et al., 2008), and male
germ cell tumors (Figure 3G) (Korkola et al., 2006). Upregulation of PDI in select
cancer types has also been confirmed by several proteome analyses. PDI protein
levels have been shown to increase in patient tissues of prostate
adenocarcinoma compared to benign prostate hyperplasia (BPH), and were
significantly higher in prostate adenocarcinoma tissues of Gleason score 7
compared to those of Gleason score 5 (Alaiya et al., 2011). Studies using 2-DE
in combination with MALDI-TOF show that PDI was significantly over-expressed
in the infiltrating ductal carcinoma of both the female (Chahed et al., 2005) and
the male breast (Chahed et al., 2008) compared to the respective adjacent non-
neoplastic breast tissues. Another study reported that PDI was one of the most
up-regulated proteins in breast tumor interstitial fluids, and thus could serve as a
potential serological marker for early detection of breast cancer (Gromov et al.,
18
2010). Upregulation of PDI protein abundance also correlates with cancer
metastasis and invasion. Increased PDI protein levels were observed in axillary
lymph node metastatic breast tumor compared to primary breast tumor
(Thongwatchara et al., 2011). PDI was also found to be strongly expressed on
migrating glioma cells in an in vitro migration assay, and on invasive glioma cells
in both xenografts and at the invasive front of human glioblastomas (Goplen et
al., 2006). Together, these data suggest that PDI is highly expressed in cancer
tissues, and that PDI can potentially be used as a diagnostic marker for select
cancer types.
19

Figure 3. PDI is highly expressed in multiple cancer types compared to respective normal
tissues. Cancer types analyzed include (A) brain (Bredel et al., 2005; 2008; Gutmann et al.,
2002; Rickman et al., 2001; Shai et al., 2003; Sun et al., 2006), (B) lymphoma (Basso et al., 2005;
Compagno et al., 2009; Piccaluga et al., 2007), (C) kidney (Beroukhim et al., 2009; Jones et al.,
2005; Yusenko et al., 2009), (D) prostate (Singh et al., 2002; Welsh et al., 2001), (E) lung (Beer
et al., 2002), (F) ovarian (Bonome et al., 2008), and (G) male germ cell tumors (Korkola et al.,
2006). Data sets were obtained from Oncomine
TM
(Compendia Bioscience, Ann Arbor, MI), and
analyzed using Prism 5 (GraphPad Software, Inc). Student t-test was used for statistical analysis.
Box: 25% - 75%. Whiskers: Min and Max.
20
II-4-2: PDI Is Associated with Clinical Outcomes of Cancer Therapy
Chemotherapeutic resistance is a major concern in clinical cancer treatment. It
has been shown that PDI is involved in anticancer drug resistance. Compared to
aplidin-sensitive Hela cells, aplidin-resistant Hela (Hela-R) cells showed
significantly higher levels of PDI protein, and inhibition of PDI by bacitracin
sensitized Hela-R cells to aplidin (Gonzalez-Santiago et al., 2007). These data
suggest that combination therapies with PDI inhibitors and traditional anticancer
agents may overcome drug resistance in other cancer types as well, and
probably may even achieve synergetic effects. In addition, by exploring published
microarray data sets, we also found that lower PDI expression is significantly
associated with higher overall survival rate of cancer patients, for example, in
glioblastoma (Figure 4) (2008; Shai et al., 2003). Such information suggests
cancer cell-associated PDI level may be a potential prognostic marker.
Particularly, because PDI can be secreted into extracellular tumor environment,
serum PDI levels may reflect cellular PDI in a tumor, and makes it easier for
clinical detection.
21

Figure 4. PDI expression is associated with glioblastoma patients’ overall survival
rate (2008; Shai et al., 2003). Data sets were obtained from Oncomine
TM
(Compendia
Bioscience, Ann Arbor, MI), and analyzed using Prism 5 (GraphPad Software, Inc).
Kaplan-Meier survival analysis method was used to generate survival curves. **, P <
0.01; ***, P < 0.0001.

22
II-4-3: PDI Supports Tumor Survival and Progression
Increasing evidence from functional studies has indicated that PDI plays an
important role in supporting survival and progression of multiple cancers. In
melanoma, inhibition of PDI activity using bacitracin enhanced apoptosis
triggered by fenretinide or velcade (Lovat et al., 2008). A PDI inhibitor, bacitracin
or an anti-PDI mAb inhibited in vitro migration and invasion of human glioma cells
(Goplen et al., 2006). Silencing of PDI alone has also been reported to be
enough to induce apoptosis in cultured MCF-7 (breast cancer), SH-SY5Y
(neuroblastoma) and HeLa (cervical cancer) cells (Hashida et al., 2011).
Recently, we have shown that silencing PDI in human ovarian cancer cells
resulted in substantial cytotoxicity, as measured by the MTT and the colony
formation assays (Xu et al., 2012). Additionally, PACMA 31, a novel irreversible
PDI inhibitor, exhibited significant anticancer activity in both in vitro and in vivo
ovarian cancer models (Xu et al., 2012). Although it is unclear what PDI-
implicated cellular pathways are involved in the context of cancer, we
hypothesize that ER stress contributes to the cytotoxicity induced by PDI
silencing or inhibition. A hallmark of cancer is its ability to maintain growth and
proliferation (Hanahan and Weinberg, 2000, 2011), requiring sufficient supply of
proteins for cellular construction and functions, intercellular communication,
extracellular matrix destruction, and angiogenesis stimulation. Therefore,
compared to normal cells, cancer cells express proteins more efficiently by
turning on different transcriptional (e.g. c-Myc, Jun) and translational (e.g. eIF2α,
eIF4E) machineries (Darnell, 2002; Silvera et al., 2010). As an important enzyme
23
and a chaperone for oxidative protein folding, PDI is up-regulated in order to
control excess protein production in cancer cells. Therefore, once PDI is
inhibited, the newly synthesized proteins without correct disulfide bond formation
and proper folding rapidly accumulate, triggering ER stress and eventually
causing cancer cell death. Based on this premise, PDI could serve as a
promising drug target for cancer treatment. Moreover, the role of cell surface PDI
in cancer progression is unclear. Therefore, inhibition of PDI may interrupt other
essential cancer pathways contributing to the cytotoxicity of PDI inhibition.
24
II-‐5:
APPROACHES
TO
MEASURING
PDI
ACTIVITIES
AND
SCREENING
FOR

PDI
INHIBITORS

During the past decades, a series of assays have been established for
measuring PDI’s enzymatic activities as well as screening for inhibitors. Here,
we discuss known PDI activity assays based on the different activities of PDI
(reductase, isomerase, oxidase, and chaperone). Because many of these assays
are widely used to screen for PDI inhibitors, better understanding of their
mechanisms lays a solid foundation for the development of specific and potent
inhibitors of PDI.
25
II-5-1. Reductase Assays
II-‐5-‐1-‐1:
Insulin
Turbidity
Assay

Insulin comprises of two polypeptide chains, the A and B chains that are linked
together by two disulfide bonds (Holmgren, 1979). As a reductase, PDI disrupts
the two bonds between the A and B chains. Loss of the disulfide bonds results in
the B chain aggregation while the A chain remains soluble in the solution (Figure
5A). The turbidity generated by the B chain can therefore serve as a reporter to
measure PDI’s reductase activity, and can be continuously monitored by the
absorbance at 540 ~ 650 nm using a spectrophotometer. Commonly used
electron donors for this reaction include GSH (Morjana and Gilbert, 1991) and
DTT (Holmgren, 1979; Khan et al., 2011). Such reaction has been made to be
continuously measured by coupling the formation of GSSG to NADPH oxidation
by using glutathione reductase (Gilbert, 1998; Morjana and Gilbert, 1991). In
addition, by using a stop reagent to convert into an end point assay, the insulin
turbidity assay can be modified to act as an automated and a robust system for
high throughput screening (HTS) for PDI inhibitors (Smith et al., 2004). An
effective stop reagent is H
2
O
2
because it terminates the reaction without
modifying the turbidity signal. H
2
O
2
oxidizes DTT and thus aborts the enzymatic
reaction. The relatively simple and efficient method makes the insulin turbidity
assay a commonly utilized assay in PDI studies. It is also an efficient assay for
HTS of PDI inhibitors in that all the reagents it requires are relatively available
and cost effective. The sensitivity of this assay is in the micromolar range.
26
However, it also has drawbacks. This assay measures only the reductive
properties of PDI, which is not its primary physiological function. Also, it is not a
sensitive method for quantification of PDI, and the long lag phase does not
provide an estimate of the true initial rates (Hatahet and Ruddock, 2009). For
screening or testing PDI inhibitors, the insulin turbidity assay may provide true
negative results if a tested compound is easily reduced by DTT or GSH and thus
loses its inhibitory activity.
II-‐5-‐1-‐2:
Insulin
Degradation
Assay

Utilizing
125
I radio-labeled insulin (Besanger et al., 2012), the insulin degradation
assay is a modified version of the insulin turbidometric assay (Figure 5B).
Radioisotope (
125
I) is linked to the B chain of insulin through a chelator. PDI
catalyzes the reduction of disulfide bonds, causing the precipitation of
125
I-labeled
B chain. The reaction is terminated by trichloroacetic acid, and the remaining
acid-soluble radiation intensity in the supernatant provides an estimate of PDI’s
reductase activity (Carmichael et al., 1977; Mandel et al., 1993). However, the
involvement of radioactive materials limits this assay from broad applications to
some extent. In addition, it may underestimate the PDI reductase activity when
the system is contaminated by proteases that could add extra soluble radioactive
fragments in the supernatant. It is also inconvenient to apply this assay for HTS
due to the multiple-step reactions.
27
II-‐5-‐1-‐3:

Fluorimetric
Assay

A fluorimetric assay was developed to studied the PDI reductase activity by
measuring an increase in the fluorescence intensity of the fluorescent probes
(Figure 5C). When two identical fluorescent molecules are in close proximity,
intermolecular interaction quenches the fluorescence, a phenomenon called
fluorescence self quenching. Based on this mechanism, the fluorimetric assay is
used for measuring PDI’s reductase activity. PDI catalyzes the reduction of a
fluorescent probe (e.g. di-(o-aminobenzoyl)-GSSG (diabz-GSSG), Dieosin-
GSSG (Di-E-GSSG)), where a disulfide bond links two identical fluorescent
moieties ((o-aminobenzoyl)-GSH (abz-GSH), eosin-GSSG (E-GSH)). As a result,
the two fluorescent moieties are separated, causing a substantial increase in
fluorescence intensity (Raturi et al., 2005) (Tomazzolli et al., 2006). This is a
sensitive assay capable of measuring PDI’s reductase activity at a PDI
concentration as low as 2.5 nM (Raturi et al., 2005). It is also a quantitative- and
kinetic-friendly assay. The assay exhibits a rapid increase in fluorescence
immediately upon addition of PDI, and hence the true initial rates can be
acquired. The assay is adaptable for HTS of PDI inhibitors. One shortcoming of
this assay is the high cost of the chemicals and the synthesis of fluorescent
probes.
28
II-5-2: Oxidation Assays
II-‐5-‐2-‐1:

Ribonuclease
Oxidation
Assay

Ribonuclease (RNase) digests RNA by catalyzing hydrolysis of the
phosphodiester bonds. In this assay, PDI catalyzes oxidative renaturation of
RNase from its inactive reduced form (Figure 5D) (Hong and Soong, 2008; Lyles
and Gilbert, 1991). Active oxidized RNase subsequently catalyzes the hydrolysis
of cCMP (cyclic cytidine monophosphate) into CMP (cytidine monophosphate).
The amount of CMP can be monitored by the absorbance at 296 nm wavelength,
and the rate of cCMP hydrolysis indicates the oxidase activity of PDI. Although
convenient, this assay should be used with a concern that the product (CMP) can
competitively inhibit PDI and alter the reaction rate. In addition, the calculation of
active RNase concentration depends on the K
m
value for cCMP and K
i
value for
CMP. These parameters can vary depending on the pH and salt concentration.
Other versions of this oxidation assay are also used in PDI research where
RNase and CMP are replaced by other enzymes and their substrates. However,
these alternative versions are less commonly used. For example, PDI catalyzes
the oxidation of inactive reduced lysozyme to its active oxidized form that can
digest the suspension of Micrococcus lysodeikticus cell wall (Khan et al., 2011).
Reduced, denatured bovine pancreatic trypsin inhibitor (RBPTI) was also
reported for testing PDI’s oxidase activity (LaMantia and Lennarz, 1993). In this
PDI catalyzed reaction, RBPTI is oxidized to its active form that can be measured
by virtue of its ability to inhibit trypsin.
29
II-‐5-‐2-‐2:
Peptide
Oxidation
Assay

This assay uses a decapeptide NRCSQGSCWN consisting of two cysteine
residues separated by a linker region, with a fluorescent group (tryptophan)
adjacent to one cysteine and a protonatble group (arginine) adjacent to the other
cysteine (Figure 5E) (Ruddock et al., 1996). These two groups are brought
together when PDI catalyzes the formation of the disulfide bond between the two
cysteines, resulting in fluorescence quenching. This fluorescence spectroscopy
allows rapid and reproducible determination of PDI’s oxidase activity.
30
II-5-3: Isomerization Assays
II-‐5-‐3-‐1:
Scrambled
RNase
(sRNase)
Assay

The sRNase assay is the first PDI activity assay ever published (Ibbetson and
Freedman, 1976). In this assay, sRNase is prepared by subjecting RNase to
denaturing conditions such that it is in a fully oxidized form with randomly formed
disulfide bonds and hence, is incapable of hydrolytically cleaving RNA (Gilbert,
1998; Hillson et al., 1984). Acting as an isomerase, PDI catalyzes the exchange
of inter- and intra-molecular disulfides in sRNase, leading to the regain of the
native disulfide pairing as well as enzymatic activity in RNase (Figure 5F). Based
on this mechanism, the activity of PDI is measured by scrutinizing RNase activity
towards RNA. Regarding the fact that PDI is an essential isomerases in a cell,
this sensitive and relatively easy-to-perform assay proves more useful for
mechanistic studies. sRNase can be prepared as a stock solution and stored for
a long duration (Gilbert, 1998). Because the samples are placed in close
proximity to the photomultiplier, the assay can be employed to analyze turbid
samples. Additionally, it can test crude whole cell homogenates as well as
homogenous enzymes (Hillson et al., 1984). However, because this assay
involves withdrawing aliquots at regular intervals, such a labor-intensive process
makes it inconvenient for HTS of PDI inhibitors. In addition, the assay measures
the change in A260 relative to A280, and as such, those small molecules that
have absorption at 280 nm wavelength interfere with the assay (Smith et al.,
2004). It is also important to note that the substrate sRNase is a complex
31
mixture of species with intramolecular and intermolecular disulfides, so the
activity may exhibit batch-to-batch variation. This assay assesses the substrate
degradation rather than directly measuring PDI’s isomerase activity (Gilbert,
1998). The RNA hydrolysis may be catalyzed by an intermediate instead of a
final native RNase product from the isomerization reaction resulting in
miscalculating the isomerase activity of PDI.
II-‐5-‐3-‐2:
Bovine
Pancreatic
Trypsin
Inhibitor
Refolding
Assay

Bovine pancreatic trypsin inhibitor (BPTI) is a single chain polypeptide that
suppresses trypsin in protein digestion. In its native conformation, the protein
folds upon itself and is held together by three disulfide bonds (3S) (Creighton and
Goldenberg, 1984). In this assay, BPTI is trapped in non-native two-disulfide-
bond (2S) states, and PDI acts as an isomerase to catalyze disulfide
rearrangement in BPTI to refold it to its native conformation (Figure 5G)
(Karala
et al., 2007; Karala and Ruddock, 2010). At set timepoints, the reaction is
quenched by the addition of iodoacetamide that reacts with free thiol groups and
adds 57 Da to the protein mass. The isomerase activity of PDI can be assessed
by using ESI-MS to analyze the amount of refolded BPTI (3S) and its folding
intermediates (1S and 2S). This assay is time consuming as refolded BPTI and
its folding intermediates need to be purified for the ESI-MS analysis. In addition,
when this assay is used for testing PDI inhibitors that interfere with the MS
analysis, additional steps are required to remove the compounds after the
reaction (Karala and Ruddock, 2010). Therefore, this assay is not amenable for
HTS. In addition, different BPTI protein species (1S, 2S and 3S) may influence
32
their detection by ESI-MS. The results from this assay are semi-quantitative
(Karala and Ruddock, 2010).
33
II-5-4: Protein Chaperone Assays
II-‐5-‐4-‐1:
Protein
Aggregation
Assay

Rhodanese consists of a single polypeptide chain folded into two domains of
equal size. Guanidine hydrochloride (Gdn-HCl)-denatured rhodanese tends to
aggregate during self refolding process in the absence of protein chaperones.
Based on this property, refolding of denatured rhodanese can be used for
studying the chaperone activity of PDI that can be measured
spectrophotometrically at 320 nm (Figure 5H) (Karala and Ruddock, 2010). The
absence of disulfide bonds in rhodanese makes it suitable for testing the
chaperone activity of PDI independent of its isomerase activity (Song and Wang,
1995). Other proteins have also been used in the protein aggregation assay to
study PDI’s chaperone activity, such as D-glyceraldehyde-3-phosphate
dehydrogenase (GAPDH, another protein containing no disulfide bonds) (Cai et
al., 1994). In addition, citrate synthase and luciferase have been used in the
protein aggregation assay to study the chaperone activity of DsbG, a protein
disulfide isomerase present in the periplasm of Escherichia coli (Shao et al.,
2000). Besides chemical denaturation using Gdn-HCl, thermal denaturation is
also utilized for the protein aggregation assay. Substrate proteins used for
thermal denaturation include alcohol dehydrogenase (ADH) (Primm and Gilbert,
2001) and citrate synthase (Horibe et al., 2001).
34
II-‐5-‐4-‐2:
Green
Fluorescent
Protein
(GFP)
Assay

In addition to preventing protein aggregation, PDI’s chaperone activity also
mediates substrate proteins to regain their activities. In this GFP assay, refolding
of acid denatured GFP is promoted by PDI, resulting in an increase in
fluorescence that can be monitored in real time (Figure 5I) (Mares et al., 2011).
GFP serves as a model substrate to study the chaperone activity of PDI because
not only does it lack disulfide bonds but also upon acid denaturation (pH 1.5) it
exhibits a low fluorescence intensity compared to the active structure.
35

Figure 5. PDI activity assays. (A) Insulin turbidity assay. PDI catalyzes reduction of disulfide
bonds between insulin A and B chains, causing aggregation of B chain. (B) Insulin degradation
assay. PDI catalyzes reduction of disulfide bonds between A chain and
125
I-labeded B chain in
insulin, causing B chain aggression and a decrease in radiation intensity in the supernatant. (C)
Fluorimetric assay. PDI catalyzes the reduction of the disulfide bond in di-(o-aminobenzoyl)-
GSSG (diabz-GSSG), and releases two (o-aminobenzoyl)-GSH (abz-GSH) molecules with an
increase of fluorescent activity. (D) RNase oxidation assay. PDI catalyzes oxidation of reduced
RNase to its active form that hydrolyzes cCMP into CMP, with an increase in A960. (E) Peptide
oxidation assay. PDI catalyzes formation of an intramolecular disulfide bond in NRCSQGSCWN
and thus, brings Trp and Arg in close proximity, resulting in quenching of Trp fluorescence by Arg.
(F) sRNase assay. PDI catalyzes refolding of sRNase to its active form that digests RNA, leading
to a change in A260/A280. (G) BPTI refolding assay. PDI catalyzes the refolding of BPTI from its
non-native 2S form to the native 3S form. The change is subsequently measured using ESI-MS.
(H) Protein aggregation assay. PDI mediates denatured rhodanese refolding, preventing folding
rhodanese from aggregation. (I) GFP assay. PDI catalyzes the folding of non-fluorescent
denatured GFP to its native form that regains fluorescent activity.
36
II-‐6:
SMALL-‐MOLECULE
INHIBITOR
OF
PDI

Although PDI has been intensively studied in the past decades, no selective PDI
inhibitors have emerged for clinical use. Among the limited number of PDI
inhibitors, most are neither potent nor selective, and show significant off-target
toxicity. However, increasing knowledge on the role of PDI in different diseases
will ultimately lead to the discovery of potent inhibitors. Recent discoveries of
synthetic small molecules PDI inhibitors such as PACMA 31 and 16F16 have
provided necessary tools to further understand the role of PDI in human
diseases. In this section, we will provide a comprehensive review of lead PDI
inhibitors published to date. It is important to note that all purported PDI inhibitors
require further validations to be considered a bona fide and selective for this
target. Further studies will shed more light on the usefulness of these compounds
for further development. Chemical structures of select lead compounds are
shown in Figure 6 and Figure 7.
37
II-6-1: Synthetic Compounds
II-‐6-‐1-‐1:
Propynoic
Acid
Carbamoyl
Methyl
Amides

Propynoic acid carbamoyl methyl amides (PACMAs) are a class of novel small
molecules with significant cytotoxicity towards a board range of cancer cells
(Yamada et al., 2011). I have recently reported a representative compound,
PACMA 31 (1) as an irreversible inhibitor of PDI (Chapter III) (Xu et al., 2012). In
the insulin turbidity assay, compound 1 showed an IC
50
value of 10 µM in
inhibition of PDI’s reductase activity. Analyzed using mass spectrometry, the
terminal propynoic group of compound 1 covalently reacted with the thiol groups
of PDI’s active-site cysteines. The binding of compound 1 to PDI also alters its
secondary protein structure. However, compound 1 shows no significant binding
to other cysteine-containing proteins, such as BSA and Grp78. By inhibiting PDI,
compound 1 exhibited cytotoxicity towards a panel of cultured human ovarian
cancer cells, including OVCAR-3, OVCAR-8, and NCI/ADR-RES (paclitaxel- and
doxorubicin-resistant). Compound 1 showed tumor targeting ability in a mouse
xenograft model bearing human ovarian tumor, and suppressed tumor growth via
i.p. injection or oral administration, without significant toxicity towards normal
tissues. There is also a marked correlation between PDI inhibitory activity and
cytotoxicity in ovarian cancer cells among all PACMA analogs (S. Xu, S.
Saranya, and N. Neamati, unpublished data). SAR analysis showed that the
terminal propynoic moity is essential for PACMAs’ PDI inhibition and cytotoxicity
toward cancer cells, although modifications in other functional groups also to
38
different extents alter their activity. A fluorescent analog, PACMA 57 (compound
2), synthesized via conjugation of a fluorescent molecule BODIPY to compound
1, showed similar properties to the lead compound 1 (Xu et al., 2012). Therefore
this fluorescent analog (2) serves as a useful tool to further study the in vitro and
in vivo properties of PACMAs, and to expand our knowledge of PDI-implicated
cellular pathways.
II-‐6-‐1-‐2:
16F16
(3)

16F16 (3) was identified in a HTS of 68,887 compounds for the ability to
suppress apoptosis in an in vitro P12 cell based model of Huntington's disease
(Hoffstrom et al., 2010), in which apoptosis is induced by polyglutamine (polyQ)
and mediated by PDI. By using Huisgen cycloaddition chemistry (or “click
chemistry”) and MS, PDI and one of its isoforms ERp57 (also known as PDIA3)
were identified as cellular targets of compound 3. This irreversible inhibitor
showed an IC
50
of 63 µM in inhibition of PDI’s reductase activity using a
fluorimetric assay. Within 3 - 12 µM range, compound 3 showed a dose-
dependent rescue of polyQ-induced apoptosis that correlated with PDI inhibition.
At concentrations > 12 µM, compound 3 showed cytotoxicity due to PDI inhibition
as well as potential off-target effects. SAR analysis showed that similar to
compound 1 the chloroacetyl moiety was critical to its activity in that replacement
of the chlorine with a methyl group caused a substantial drop in compound 3’s
ability to suppress polyQ-induced cell death and to inhibit the enzymatic activity
of PDI. In addition, its ester moiety could tolerate small modifications such as a
replacement of methyl with ethyl, whereas significant changes such as an
39
incorporation of biotin or fluorescein affinity tags caused a complete loss of
activity. Compound 3 was also tested for its protective effects in a corticostriatal
brain slice model of neurotoxicity induced by mutant huntingtin gene (HTT) exon
1 (HTT-N90Q73), where compound 3 showed dose-dependent rescue of
neurotoxicity in medium spiny neurons (MSNs) in the striatal region of the brain
slices. Compound 3 may bind to PDI’s active-site cysteines in a similar manner
with compounds 1 and 2. It is likely that compounds 1-3 may bind to other
proteins, especially free cysteine-containing proteins.
II-‐6-‐1-‐3:
Arsenical
Compounds

Phenylarsine oxide (PAO, 4) is known to crosslink vicinal sulfhydryl groups (Frost
and Lane, 1985), and form coordination bonds through its As
+3
with the vicinal
thiols of the CXXC motif of proteins such as PDI (Kalef and Gitler, 1994). It
induced rapid shedding of L-selectin from isolated neutrophils, a process
negatively regulated by PDI (Bennett et al., 2000). It also inhibited PDI-catalyzed
reductive release of acid soluble [
125
I] tyramine-SH from surface bound [
125
I]
tyramine-SS-poly(D-lysine), with an IC
50
of 5.8 µM. It is effective prior to or during
HIV-1 infection but not after the infection progressed in P4, PM1, H9, 1G5 and
macrophage depleted peripheral blood monocytic cells. aPAO (5), a para-amino
derivate of PAO, prevents virus entry into cells (Gallina et al., 2002). However,
this class of PDI inhibitors possess low specificities for PDI as they can react with
other proteins containing the CXXC motif. For example, compound 15 has been
reported to inhibit protein tyrosine phosphatase (PTPase) (MacRobbie, 2002)
40
and Rho GTPase (Gerhard et al., 2003). In fact, it is a widely used PTPase-
specific inhibitor.
II-‐6-‐1-‐4:
Sulfhydryl
Reagents

A series of sulfhydryl reagents were reported to inhibit PDI’s catalytic activity.
They react with the free thiol groups in PDI and therefore act as irreversible
inhibitors. Generally, they exhibit relatively low specificities for PDI.
DTNB [5,5'-dithiobis(2-nitro benzoic acid), 6], known as Ellman’s reagent, is a
membrane impermeable sulfhydryl blocker. It came to be recognized for its ability
to inhibit the activation of diphtheria toxin, a process involving disulfide bond
cleavage mediated by cell surface PDI (Ryser et al., 1994; Ryser et al., 1991).
The inhibition of diphtheria toxin activation by compound 6 was similar to the
inhibition of PDI by bacitracin or anti-PDI antibodies (Mandel et al., 1993). In the
insulin degradation assay, compound 6 inhibited the reductase activity of PDI
with an IC
50
of 100 µM and showed complete inhibition at 1 mM (Ryser et al.,
1994). It also prevented HIV-1 infection of H9 cells in a dose-dependent manner,
with an IC
50
of 0.3 mM. These similar IC
50
values suggest that compound 6
blocked HIV-1 infection via PDI inhibition. Similar to compound 6, pCMBS (7) is
another membrane impermeable sulfhydryl reported to bind to the thiol groups in
PDI and prevent diphtheria toxin activation (Feener et al., 1990; Ryser et al.,
1991).
Alkylators and unsaturated aldehydes were reported to inhibit the reductase
activity of PDI in the insulin turbidity assay at low physiological pHs (Liu and Sok,
2004). Among the compounds tested, iodoacetamide (8) was found to be the
41
most potent (IC
50
= 8 µM) and maximum inhibition was observed at pH 6. A
similar result was obtained by using the NADPH/Insulin assay. N-ethylmaleimide
(NEM; 9) also showed an IC
50
value similar to that of compound 8. The IC
50
value
for acrolein (10) at pH 6.3 was found to be 10 µM. In addition, thiomuscimol (11)
and cystamine (12) were also reported to inhibit PDI’s reductase activity in the
fluorimetric assay, with IC
50
values of 23 and 66 µM, respectively (Hoffstrom et
al., 2010). SAR analyses showed that replacement of the sulphur by oxygen in
compound 11 or reduction of the intramolecular disulfide in compound 12
completely abolished their inhibitory activities, confirming that they act as
irreversible inhibitors. Like compound 3, these compounds (11 and 12) were also
able to rescue polyQ-induced apoptosis in a cell-based assay.
42

Figure 6. Chemical structures of PDI inhibitors from synthetic compounds.

43
II-6-2: Plant Metabolites
II-‐6-‐2-‐1:
Juniferdin
(13)
and
Its
Analogs

Juniferdin is a sesquiterpenoid originally isolated from the plant Ferula juniperina.
This natural product was identified as a hit in a high throughput insulin turbidity
screening of 10,000 RIKEN Natural Product Depository (NPDepo) compounds
for PDI inhibitors as anti-HIV-1 agents (Khan et al., 2011). An IC
50
value of 0.156
µM in the insulin turbidometric assay makes juniferdin the most potent inhibitor of
PDI’s reductase activity among the 10,000 compounds tested. Based on this hit
compound, a series of analogs were synthesized. Although all the derivatives
showed lower inhibitory activity on PDI than juniferdin, they provided useful
information for SAR. The sesquiterpene ring turned out to be essential for the
activity of juniferdin, because replacement with a 1-octyl completely abolished its
inhibitory activity, whereas analogs with other ring structures, such as cyclooctyl,
cyclododecyl or (1R)-menthyl, could retain modest activity. Modifications on the
sesquiterpene ring also affected the activity of juniferdin. While the 9,10-
monoepoxide 1:1 stereoisomers showed similar inhibition to juniferdin, the
2,3,9,10-diepoxide or the 4,9,10-trihydroxy showed no inhibition, and the 2,3-
monoepoxide and the stereoisomeric 4,9,10-trihydroxy derivative exhibited 15-
and 11-fold lower PDI inhibition, respectively. However, compound 14 carrying a
9,10-monoepoxide showed similar activity with juniferdin, with an IC
50
value of
0.167 µM. In addition, the p-hydroxybenzoate group is also important.
Replacement with p-methoxybenzoate completely abolished activity, whereas
44
lack of p-hydroxyl or replacement with p-fluorobenzoate or p-acetylbenzoate led
to 30-, 6-, and 4-fold lower inhibition. Juniferdin and compound 14 were also
shown to be specific inhibitors of the reductase activity of PDI, as they had no
significant inhibition of PDI’s oxidase activity in oxidation assays using RNase or
lysozyme. Further more, it had negligible inhibition of other PDI family reductases
ERp57 and ERp72 that share the same active site CGHC. These results suggest
that 13 and 14 do not bind to the active site in PDI; instead, they may bind to
residues or domains specific for PDI’s reductase activity. Although both 13 and
14 inhibited PDI-catalyzed reduction of gp120 in vitro, only juniferdin showed
pronounced cytotoxicity in HeLa, HepG2, HT1080 and K562 cell lines.
Considering that PDI silencing using siRNA, shRNA, or PDI inhibitors (e.g.
compound 1, 2, 3) resulted in cytotoxicity (Hashida et al., 2011; Hoffstrom et al.,
2010; Park et al., 2006; Xu et al., 2012), it is possible that compound 14 may
have poor stability.
II-‐6-‐2-‐2:
Quercetin-‐3-‐Rutinoside
(15)

Quercetin-3-rutinoside is also known as rutin, a natural product belonging to the
flavonol family. It is a plant polyphenolic compound widely consumed in daily
foods, such as buckwheat, berries, tea, and vegetables. A high throughput insulin
turbidity screening of a library of 4,900 compounds identified rutin as a lead
inhibitor of PDI’s reductase activity, with an IC
50
value of 6.1 µM (Jasuja et al.,
2012). Rutin does not covalently bind to PDI, as it showed reversible inhibition of
PDI’s reductase activity in a fluorimetric assay using Di-E-GSSG as a probe. The
K
d
value of rutin binding to PDI was 2.8 µM. SAR analysis indicates that the
45
sugar moiety is essential for the activity of the compound. Replacement of the
rutinose with galactose, glucose or glucuronic acid resulted in analogs with
similar PDI inhibitory activities, showing IC
50
values of 5.9, 7.1 and 5.9 µM,
respectively. Analogs without the sugar moiety, such as quercetin, tamarixetin,
isorhamnetin or diosmetin, completely lost inhibitory activity. In addition, rutin
acts as a relatively specific inhibitor of PDI. While it inhibited PDI by 60% at 30
µM, only negligible inhibition (< 10%) was observed for other oxidoreductases
sharing the active site CGHC of PDI, including ERp5, ERp57, ERp72, thioredoxin
and thioredoxin reductase. Cellular assays further showed that rutin prevented
aggregation of human and mouse platelets and endothelial cell-mediated fibrin
generation in human endothelial cells. In vivo, rutin blocked thrombus formation
by inhibiting PDI. While genetic deletion of PDI is toxic to cells (Hashida et al.,
2011; Hoffstrom et al., 2010; Park et al., 2006; Xu et al., 2012), incubation of rutin
at 100 µM with cultured endothelial cells for over 72 hours showed no toxicity,
indicating that it may have poor cell permeability and only target extracellular
PDI. In fact, rutin was tested in two clinical trials. NCT00003365 protocol
evaluating the effect of rutin on preventing the development of colon cancer was
terminated. The results of protocol NCT01254006 examining the effect of rutin in
combination with forskolin, vitamins B1 and B2 have not been released.
46
II-6-3: Antibiotics
II-‐6-‐3-‐1:
Bacitracin

A cyclic dodecapeptide antibiotic bacitracin was reported in 1981 to be the first
PDI Inhibitor (Roth, 1981). Its inhibition of the reductase activity of PDI was
demonstrated by the insulin degradation assay, with an IC
50
of 90 µM. Since
then, bacitracin has been widely used to study the biochemistry of PDI as well as
PDI’s role in various cellular events, including melanoma cell death (Lovat et al.,
2008), glioma cell invasion (Goplen et al., 2006), virus entry (Jain et al., 2007;
Markovic et al., 2004; Ryser et al., 1994), platelet adhesion (Essex and Li, 1999;
Essex et al., 2001; Lahav et al., 2003) and thrombus formation (Cho et al., 2008),
gamete fusion (Jain et al., 2007), Cu/Zn superoxide dismutase aggregation in
motor neurons (Atkin et al., 2006), stroke protection (Descamps et al., 2009), the
vitamin K cycle (Wajih et al., 2007), the regulation of NFκB (Higuchi et al., 2004)
and NAD(P)H oxidase (Janiszewski et al., 2005), the reductive activation of
cholera (Orlandi, 1997) and diphtheria (Mandel et al., 1993) toxins, the shedding
of human thyrotropin receptor ectodomain (Couet et al., 1996). It also served as
a standard control in screening compounds for the ability to inhibit the reductive
activity of PDI (Khan et al., 2011). In fact, natural bacitracin is produced by
certain strains of Bacillus licheniformis and Bacillus subtilis as a mixture of over
22 structurally related peptides (Govaerts et al., 2003). Among them, bacitracin A
(16) is the major analog (Karala and Ruddock, 2010), and B, F, H are also of
relatively high abundance in commercial bacitracin mixtures (Dickerhof et al.,
47
2011). A recent study reported that the major bacitracin analogs, A, B, F and H
had different inhibition of PDI’s reductase activity in the insulin turbidity assay,
with IC
50
values of 590, 1050, 20 and 40 µM, respectively (Dickerhof et al.,
2011). MALDI-TOF/TOF MS demonstrated that bacitracin bound PDI with
disulfide bond formation between an open thiol form of the bacitracin thiazoline
ring and Cys314/345 in the substrate-binding b’ domain of PDI (Dickerhof et al.,
2011). Another study confirmed that bacitracin had no inhibition of the reductase
activity of the isolated catalytic a domain of PDI in the insulin turbidity assay
(Karala and Ruddock, 2010). Bacitracin is nonspecific for PDI. It binds and
inhibits other proteins with or without PDI activity. For example, bacitracin
inhibited the PDI activity of fibronectin (FN), leading to a reduction in FN’s
association into an insoluble matrix (Weston et al., 2001). At 1 mM, bacitracin
completely inhibited the oxidase activity of FN in the RNase oxidation assay but
only 25% of that of PDI. A recent study extensively examined bacitracin different
PDI activity assays (Karala and Ruddock, 2010), showing that bacitracin had no
significant inhibition of PDI’s oxidase activity in the peptide oxidation assay, or
the isomerase activity in the BPTI refolding assay. In the protein aggregation
assay, bacitracin prevented refolding rhodanese from aggregation in a PDI-
independent manner without a significant effect on PDI’s chaperone activity,
whereas it inhibited the chaperone activity of BiP, an ER-resident molecular
chaperone (Karala and Ruddock, 2010). However, another study showed that
bacitracin (15 µM) inhibited the chaperone activity of PDI without having a
substantial effect on substrate aggregation in the absence of PDI in the protein
48
aggregation assay (Primm and Gilbert, 2001). In the insulin turbidity assay
bacitracin inhibited the reductase activity of PDI as well as Escherichia coli DsbC
in a dose-dependent manner in the milimolar range via competition of substrate
binding (Karala and Ruddock, 2010). Hence, there is a need to re-evaluate the in
vivo effects of bacitracin and its relationship with PDI. The off-target effects of
bacitracin should be considered in its future use for PDI studies. Although widely
used in research on PDI-implicated diseases for decades, bacitracin failed to
enter clinical trials primarily because of its week cell permeability (Godin and
Touitou, 2004) and its nephrotoxicity (Wang et al., 2008a).
II-‐6-‐3-‐2:
Ribostamycin
(17)

Produced by Streptomyces ribosidificus, ribostamycin is an aminoglycoside
antibiotic effective against both gram-positive and gram-negative strains. The
affinity column chromatography of proteins in bovine liver resulted in the
identification of PDI as the main binding protein target of 17 (Horibe et al., 2001).
Surface plasmon resonance determines the K
d
of PDI to ribostamycin to be 319
µM, which is over 100 times higher than that of rutin. Ribostamycin inhibited
PDI’s chaperone activity in a dose-dependent manner in protein aggregation
assays with rhodanese, citrate synthase and GAPDH. It required at least a 100:1
molar ratio of ribostamycin to PDI to sufficiently inhibit the chaperone activity of
PDI. However, ribostamycin had no effect on the isomerase activity of PDI,
indicating that it does not bind to the active site of PDI. Ribostamycin is the first
reported inhibitor of the chaperone activity of PDI. It is important to note that PDI
is not the only protein ribostamycin binds to. For example, it was also
49
documented to target the 16S ribosomal RNA to cause mistranslation (Moazed
and Noller, 1987).
II-‐6-‐3-‐3:
Other
Antibiotics

Based on the discovery of ribostamycin as a PDI chaperone inhibitor, a panel of
other antibiotics were also screen for their abilities to inhibit the chaperone
activity of PDI (Horibe et al., 2002). Several other antibiotics were found to bind
PDI, including vancomycin (K
d
= 206 µM), sisomicin (K
d
= 392 µM), neomycin (K
d

= 872 µM), gentamicin (K
d
= 904 µM), kanamycin (K
d
= 1.05 mM), and
streptomycin (K
d
= 1.25 mM). They all showed inhibition of the chaperone activity
of PDI. Particularly, vancomycin and sisomycin sufficiently inhibited the
chaperone activity at a 100:1 molar ratio of antibiotic to PDI. It is still unclear
whether these compounds bind and inhibit PDI in vivo. Also, it remains unknown
whether or not PDI inhibition contributes to their antibiotic action. However, the
advantage of using these antibiotics for PDI inhibition is that these are relatively
well-recognized compounds and their toxicity has already been assessed (Davis,
1987).
50
II-6-4: Hormones
II-‐6-‐4-‐1:
Estrogens

At 1 µM concentration, several estrogens have been shown to inhibit PDI’s
reductase activity by over 30% in the insulin degradation assay, including estrone
(E
1
, 18, 56%), 17β-estradiol (E
2
, 19, 55%), diethylstilbestrol (DES, 45%), and
estriol (E
3
, 38%). In addition, E
1
and E
2
were also tested in the sRNase assay
and showed inhibition of PDI’s isomerase activity (Tsibris et al., 1989). However,
no significant inhibition of PDI’s chaperone activity was observed in the protein
aggregation assay (Primm and Gilbert, 2001). In fact, amino acid sequence
segments in PDI have significant similarity with the estrogen binding domain in
estrogen receptor (ER), but not with the steroid domains of the progesterone and
glucocorticoid receptors or with thioredoxin (Tsibris et al., 1989). The ALIGN
scores are 6.3 and 5.4 for the 120-163/182-230 PDI segments compared to the
350-392/304-349 ER segments. Another study further confirmed that PDI had
one E
2
binding site, and the K
d
of E
2
binding to PDI was 2.1 ± 0.5 µM (Primm and
Gilbert, 2001). This E
2
binding site is distinct from the peptide/protein binding
sites and the bacitracin binding site. In addition, E
2
is also a potent inhibitor of
somatostatin binding to PDI, whereas 17α-E2 and DES are the next potent
inhibitors (Hiroi et al., 2006).
II-6-4-2: Thyroid
Hormones

3,3’,5-triiodo-L-thyronine is also known as triiodothyronine or T
3
(20). It is a
thyroid hormone that has important roles in numerous biological processes, such
51
as cell growth, development and differentiation, energy metabolism, and
regulation of body temperature and heart rate (Ichikawa and Hashizume, 1991).
As T
3
carries out multiple activities, it was proposed to bind to a different target
apart from the previously known target, the nuclear receptor c-erbA. By using an
affinity labeling reagent N-bromoacetyl-3,3’,5-[
125
I]triiodo-L-thyronine
(BrAc[
125
I]T
3
), a 55-kDa polypeptide was identified as a major T
3
-binding protein,
called T
3
BP, in various cell types (Cheng, 1983; Horiuchi et al., 1982). Sequence
analysis of T
3
BP cDNAs determined T
3
BP to actually be PDI (Yamauchi et al.,
1987). It was further indicated that at equilibrium T
3
bound to two independent
sites in PDI (Guthapfel et al., 1996; Primm and Gilbert, 2001). One study
conducted by Guthapfel et al showed that while the first binding site exhibited
high affinity and could be saturated at near physiological T
3
concentrations with a
K
d
of 21 nM, it had a remarkably low B
max
(1.8 mmol T
3
/mol PDI monomer),
implying T
3
binding is mainly nonspecific; the second binding site had low affinity
and was unsaturated up to 100 µM (Guthapfel et al., 1996). Later Primm et al
suggested that regarding the low B
max
, the previously published high binding
affinity was due to an impurity rather than PDI itself, and reported that the two T
3

binding sites in PDI had comparable affinity, with K
d
values of 4.3 ± 1.4 µM
(Primm and Gilbert, 2001). One of the two binding sites also bound bis-ANS (a
hydrophobic probe), and the other overlapped the E2 site. T3 can completely
displace E
2
, with half-maximal inhibition at 8.5 ± 3.2 µM (Primm and Gilbert,
2001). In addition to T
3
, a competition assay showed that PDI also bound a wide
variety of T
3
analogs, including D-T
3
, 3,3’,5-triiodothyropropionate, 3,3’,5-
52
triiodothyroacetate, 3,5-diiodo-L-tyrosine, L-thyronine, and 3,5-diiodo-L-thyronine.
Primm et al reported they did not observe significant inhibition of PDI’s catalytic
activity in mediating RNase refolding, nor any effects on the chaperone activity
(Primm and Gilbert, 2001). In contrast, Guthapfel et al showed that T
3
inhibited
the activity of PDI in mediating RNase refolding under similar conditions, with an
inhibition constant K
i
of 1.3 ± 0.5 µM, and its analogs D-T
3
and L-thyronine were
more potent inhibitors (Guthapfel et al., 1996). The ability of T
3
to inhibit PDI’s
catalytic activity was further supported by Hirol et al (Hiroi et al., 2006), showing
35% inhibition at 20 µM and the half-maximual inhibition at 3.49 µM. It was
suggested that since T
3
was found in nanomolar concentration in the cells but
had a high K
i
for PDI, the inhibition of PDI redox activity could not be counted as
a physiological action (Guthapfel et al., 1996). To date, the in vivo inhibitory
effect of T
3
on PDI remains elusive. It would be interesting to examine whether T
3

binds and inhibits PDI in vivo, and whether it contributes to the physiological
functions of T
3
. AT
3
(21), an N-acetylated form of T
3
, shows marked PDI
inhibition. It inhibited PDI-catalyzed reductive cleavage of the cell surface bound
[
125
I]tyramine-SS-poly(D-lysine), with an IC
50
of 70 µM (Gallina et al., 2002). It
also showed significant inhibition of HIV-1 entry into host cells.
Regarding the high abundance of PDI, the binding ability of PDI to hormones
suggests that PDI serves as a high capacity hormone reservoir of the ER.
53
II-6-5: Xenoestrogens
Xenoestrogens are chemical compounds that imitate estrogen. Bisphenol A [2,2-
bis-(4-hydroxyphenyl) propane; BPA; 22] is a synthetic versatile industrial
monomer for plastic products, and is considered as an environmental hazard
(Okada et al., 2008; Rubin, 2011). As a hormone disrupting chemical, it strongly
binds to estrogen-related receptor γ (ERR-γ) with a K
d
of 5.5 nM and preserves
its basal constitutive activity (Matsushima et al., 2007) For the purpose of
defining its biological aspects, BPA binding proteins were screened in rat brain,
resulting in identifying PDI as a major binding protein (Hiroi et al., 2006). The K
d

of BPA binding to recombinant rat PDI was 22.6 ± 6.6 µM, and that for
recombinant human PDI was 17.51 ± 3.93 µM. Although BPA has a lower affinity
to the hormone-binding site on PDI than those of E
2
(K
d
= 2.1 ± 0.5 µM (Primm
and Gilbert, 2001)) and T
3
(K
d
= 4.3 ± 1.4 µM (Primm and Gilbert, 2001)), BPA
acted as a competitive inhibitor in PDI-binding to E
2
and T
3
, which may partially
explain its adverse biological effects. In the RNase oxidation assay, BPA
inhibited the oxidase activity of PDI in a dose-dependent manner and reached a
plateau at 20 µM, with a maximum 24% inhibition. The half-maximal inhibition
was 3.72 µM. In addition, an analog of BPA, tetrachlorobisphenol A (23) was
reported to be the most potent inhibitor of T
3
binding to PDI with an IC
50
value of
0.2 µM, 100-fold lower than that of BPA (Imaoka, 2011). Further studies identified
the binding site of BPA to PDI located within the a and b’ domains, and indicated
the b’ domain contributed to inhibition of catalytic activity of PDI by BPA
(Hashimoto et al., 2012). In addition, mutation of H258 within the b’ domain
54
completely prevented BPA binding to PDI and mutations of Q245 and N300
within the b’ domain also decreased the binding affinity, indicating that these
residues are important sites for their interaction.

Figure 7. Chemical structures of PDI inhibitors from plant metabolites, antibiotics,
hormones, and xenoestrogens.

55
II-‐7:
CONCLUDING
REMARKS

Extensive studies on PDI in the past decades have elaborated on many aspects
of PDI, including the structure and domain architecture, biochemical redox
reactions, cellular pathways, physiological roles and involvement in multiple
diseases. Accumulated data demonstrate the important role of PDI in regulating
a vast variety of biochemical events essential for cell homeostasis. Dysregulation
of PDI's gene expression, post-translational modification, or enzymatic activity
results in the development of various human diseases. However, it was only until
recently that PDI's role in cancer became more recognized. Because PDI is
highly expressed in select cancer types, supports tumor growth and progression,
and is associated with clinical outcomes, it is a potential drug target for cancer
therapy.
To date, PDI has not yet been clinically used as a drug target for any diseases.
One reason is that the exact PDI pathways and the causes for their pathological
dysregulation remain unclear in many PDI-associated diseases. Hence there
arises a need to characterize the structural and functional aspects of PDI in the
context of diseases. One the other hand, more importantly, it is because there is
a dearth of selective and potent PDI inhibitors. Bacitracin, the first PDI inhibitor,
failed to enter clinical trials because of its off-target toxicity and weak cell
permeability. Similar reasons limited many other PDI inhibitors from clinical
studies. The development of robust PDI activity assays has led to the recent
discoveries of a series of novel PDI inhibitors, such as the irreversible inhibitors
56
PACMA 31 and 16F16, and the reversible inhibitors juniferdin and rutin. These
novel PDI inhibitors have proven potent in disease models of cancer,
Huntington's disease, HIV-1 infection and thrombosis. They not only provide
useful tools for further exploring the biology of PDI, but also serve as leads for
further optimization that is required for selecting viable candidate for clinical
studies.
57
CHAPTER III: DISCOVERY OF AN ORALLY ACTIVE
IRREVERSIBLE INHIBITOR OF PROTEIN DISULFIDE
ISOMERASE

Previously, my lab reported that a class of propynoic acid carbamoyl methyl
amides (PACMAs) showed a broad spectrum of cytotoxicity in a panel of human
cancer cell lines, with relatively selective potency in ovarian cancer cells resistant
to doxorubicin and paclitaxel (Yamada et al., 2011). In this study, with a series of
newly designed and synthesized PACMA derivatives, I established that these
novel small molecules are potent irreversible PDI inhibitors with marked
cytotoxicity in human ovarian cancer cells. Among them, PACMA 31, exhibited
in vivo activity with oral bioavailability in a mouse xenograft model of human
ovarian cancer. To my knowledge, PACMA 31 is the first orally active small-
molecule PDI inhibitor with desirable pharmacological properties for cancer
treatment. Most importantly, this study provided strong evidence that PDI is a
druggable target for cancer therapy and opens a new area of research to develop
new treatments with a novel mechanism of action.
58
III-‐1:
RESULTS

III-1-1: Novel PACMAs Demonstrate Cytotoxicity in a Panel of Ovarian
Cancer Cell Lines.
In order to establish more informative structure-activity relationships and gain key
insights on the likely protein targets of PACMAs in cancer, a series of new
PACMA derivatives were designed and synthesized (Figure 8). The new
compounds were tested in human ovarian cancer cell lines OVCAR-8, NCI/ADR-
RES, HEY and OVCAR-3. Many of these compounds exhibited anticancer
potency with IC
50
values less than or equal to 10 µM (Table 4). It is important to
note that the NCI/ADR-RES cell line shares a large number of karyotypic
abnormalities with OVCAR-8 (Roschke et al., 2003) but expresses high levels of
MDR1 and P-glycoprotein (Alvarez et al., 1995), which makes it resistant to
multiple anticancer drugs in clinical use, including paclitaxel and doxorubicin. In
addition, the human ovarian cancer cell line HEY is naturally resistant to cisplatin
(CDDP). Similar to the PACMA compounds reported previously (Yamada et al.,
2011), a number of these novel analogs exhibited potent cytotoxicity towards
NCI/ADR-RES and HEY cells, implicating their potential ability to overcome the
current drug resistance issue in ovarian cancer therapy.
59

Figure 8. PACMA analogs used in this study. (A) General method for the synthesis of
PACMAs and analogs. (B) Structures of compounds evaluated in the study.
60
Table 4. Cytotoxicity of select PACMAs in a panel of human ovarian cancer cells
Compound
IC
50
(µM)
a

OVCAR-8 NCI/ADR-RES HEY OVCAR-3
1 (Yamada et al., 2011) 2.9 ± 1.7 0.3 ± 0.1 1.2 ± 1.2 2.0 ± 1.0
3 (Yamada et al., 2011) 2.5 ± 1.7 0.2 ± 0.1 0.5 ± 0.2 1.2 ± 0.7
4 (Yamada et al., 2011) 2.7 ± 1.6 1.0 ± 0.3 1.0 ± 0.3 2.0 ± 0.8
9 (Yamada et al., 2011) 7.8 ± 1.1 0.9 ± 0.9 1.1 ± 0.5 7.0
20 2.2 0.52 0.31 3.1
21 1.5 0.32 0.33 2.5
22 1.5 2.0 0.72 0.25
23 5.0 0.45 NT
b
0.6
24 1.5 2.5 NT 2.5
25 0.2 1.3 NT NT
26 10 1.8 NT 3
27 > 10 6.3 NT > 10
28 0.52 0.02 NT 0.23
29 0.5 ± 0.3 NT NT NT
30 2.2 0.14 NT 0.3
31 0.9 ± 0.1 1.4 ± 0.5 NT 0.32
32 6.0 8.0 NT > 10
33 5.0 > 10 0.32 NT
34 0.4 ± 0.2 NT NT NT
35 3.4 ± 0.4 NT NT NT
36 10.4 ± 1.6 12.8 ± 3.8 NT NT
37 2.2 0.33 NT 0.42
38 2.2 0.32 NT 1.5
39 2.2 0.32 NT 0.53
40 1.2 2.5 NT 2.5
41 4.3 2.2 NT 2.5
42 1.0 0.43 NT 0.43
43 > 10 > 10 NT NT
44 3.1 8.5 NT 2.3
45 1.5 9.0 NT 2.3
46 > 10 > 10 NT NT
47 > 10 > 10 > 20 > 10
48 > 10 > 10 > 20 > 10
49 > 10 > 10 > 20 > 10
50 7.0 7.3 7.0 7.3
51 > 10 > 10 >10 > 10
52 5.5 2.2 2.3 6.3
53 > 10 > 10 > 10 > 10
54 > 10 > 10 > 10 > 10
55 > 10 > 10 > 10 > 10
56 > 10 > 10 > 10 NT
a
IC 50 is defined as drug concentration causing a 50% decrease in cell population.
b
NT, not-tested.
61
III-1-2: Structure-Activity Relationships for New PACMAs and Related
Compounds
The cytotoxicity data for the new compounds revealed a number of interesting
findings regarding the role of the various parts of these molecules. PACMAs 20-
24, having variations of the phenyl substituents of the most potent compounds in
our initial study (4 and 9) (Yamada et al., 2011), resulted in only modest changes
in activity suggesting that substituent R
2
of general structure E (Figure 8A) may
not be critical. Interestingly, 26-28, that lack the R
2
substituent retained activity,
but this varied significantly. 26 and 27 were inactive against OVCAR-8 with
reduced activity in the other cell lines, while PACMA compound 28 was among
the most potent tested. Varying substituent R
3
of general structure F (Figure 8A)
led to more potent compounds, such as 29-31 or led to reduced activity in 32 and
33, implying that R
3
may be involved in binding to the target protein. By varying
both the R
3
and R
2
substituents, activity can be improved (e.g. 34), while
changes in the R
1
substituent can lead to the opposite effect (35 and 36).
However, some changes in the N-substituent R
1
were well tolerated without
significant change in activity as in compounds 3 (Yamada et al., 2011), 37-40.
Interestingly, significant changes in activity were noted among the carboxyl
analogs 41-43, suggesting that a more hydrophobic group is preferable over a
more hydrophilic one. Overall, the various modifications of the R
1
,

R
2
, and R
3
substituents led to some significant variations in potency, implying that these
compounds exert their activity via some type of interaction with their target that
involves molecular recognition and selective binding. Introducing a linker of type
62
X as in general structure F (Figure 8A) resulted in some reduction in activity (e.g.
44, 45), but the compounds remained active. However, the most unique
structural feature of the active PACMA compounds is their propynoic acid amide
unit, and its modification or replacement generally leads to inactive compounds.
For example, the replacement of the alkyne moiety in PACMA 1 (Yamada et al.,
2011) with hydrogen, alkyl or vinyl groups (e.g. 46-49) led to total loss of activity.
Notably, the lack of activity of compound 49, despite the fact that it contains the
electrophilic acrylamide moiety, implies the unique reactivity features of the
propynoic acid amide group in the PACMA compounds. Thus, while we
anticipated that the electrophilic acetylenic group of PACMAs might serve as an
electrophile for possible irreversible covalent binding to a nucleophilic moiety in
the target protein, these results suggest additional selectivity and reactivity
characteristics resulting from the unique electronic and geometric features of this
moiety. The only other electrophilic group that showed some modest activity was
the α-bromo amide group of compounds 50 and 52, while other electrophilic
groups were entirely inactive, including less reactive alkyl bromides (e.g. 51),
acrylamides (e.g. 53), and electrophilic aromatic amides (e.g. 54). Interestingly,
substituted alkynyl derivatives (e.g. 55, 56) were also inactive, further
demonstrating the unique reactivity features of the propynoic acid amide groups
of PACMAs.
63
III-1-3: Active PACMA Analogs Covalently Bind to Their Cellular
Target Protein in Human Ovarian Cancer Cells
Based on the electron-deficient nature of the propynoic acid amide moiety, and
as confirmed by the structure-activity analysis, we anticipated that the active
PACMAs would be able to react irreversibly with certain nucleophilic groups such
as the thiol groups of cysteine side chains to form covalent adducts. This
property can be used to identify the protein target responsible for their activity
and selectivity. To test this hypothesis, we first conjugated one of the most active
analogs, 31, to the fluorescent dye BODIPY, resulting in 57. We also synthesized
58, a close analog of 57 lacking the propargyl group and expected to be inactive,
as well as the BODIPY compound 59 with acylated linker that can serve as the
control (Figure 9A). The ability of 31, 57, 58 and 59 to inhibit ovarian cancer cell
growth was compared. PACMAs 31 and 57 exhibited similar potency (Figure 9B),
indicating that the conjugation of BODIPY to 31 did not affect 31’s cytotoxic
activity. No considerable cytotoxicity was observed with 58 or 59, demonstrating
that the electrophilic alkyne is essential for potency, and that the BODIPY moiety
does not contribute to cytotoxicity. The fluorescent properties of 57 (ex: 490 nm,
em: 537 nm) were determined by Fluorolog (Figure 9C). In addition, 57 and 58
displayed comparable fluorescent activity, and were slightly less fluorescent than
59 (Figure 9D), suggesting that the conjugations quenched the BODIPY’s
fluorescence only to a small extent.
To examine whether the active analog 57 covalently binds to its target protein,
we treated OVCAR-8 cells with 57, 58, 59 or equal amount of DMSO. Cells were
64
lysed after treatment, and subjected to SDS-PAGE. A fluorescent band (~ 57 kDa)
was only observed in the lane with 57-treated samples (Figure 9E). The
interaction of 57 with its cellular protein target of ~57 KDa is covalent as it was
preserved under the denaturing conditions of the SDS-PAGE.
65

Figure 9. BODIPY conjugation of PACMA 31. (A) Structures of 31, 57, 58 and 59. (B)
Cytotoxicity of 31, 57, 58 and 59 in OVCAR-8 cells was measured by the MTT assay
after 72 hr treatment. BAR, SEM. (C) Identification of excitation (upper panel, 496 nm)
and emission peaks (lower panel, 537 nm) for 57. (D) Fluorescent activities of 57, 58
and 59 (λ
ex
: 492 nm, λ
em
: 535 nm). (E) 57 covalently binds to specific cellular proteins.
Whole cell lysates of OVCAR-8 cells treated with 57, 58 and 59 at 2 µM for 30 min were
subjected to SDS-PAGE, followed by fluorescence scan (left) and SYPRO Ruby staining
(right). Arrow indicates a fluorescent band in 57-treated cells. One of three
representative experiments is shown.
66
III-1-4: Identification of PDI as a Target of PACMAs
In order to identify the 57 kDa protein, we performed two-dimensional gel
electrophoresis (2DGE) with whole cell lysates from 57-treated OVCAR-8 cells
(Figure 10A). Using mass spectrometry, PDI was identified as a protein target of
PACMA 57 (Figure 10B).
67

Figure 10. PACMA 57 covalently binds to PDI. (A) Identification of PDI as the cellular
protein target for 57. OVCAR-8 cells were incubated with 2 µM of 57 for 30 min. Whole
cell lysates were prepared as described for 2DGE, analyzed by IEF/SDS/PAGE,
scanned for BODIPY fluorescence (λ
ex
: 488 nm, λ
em
: 526 nm), and silver stained. The
fluorescently tagged protein spot (arrow, ~ 57 kDa) was excised from the gel and
analyzed by mass spectrometry. (B) PDI was identified. Identified sequences covered
21.65% of the full length PDI protein sequence.
68
To confirm PDI as the target, we treated OVCAR-8 cells with compounds 57-
59 or DMSO. Whole cell lysates were subjected to immunoprecipitation with anti-
PDI antibody. A strong fluorescent band of ~ 57 kDa was detected only in the
lane with PACMA 57-treated samples (Figure 11A), indicating that PACMA 57
covalently bound to PDI. In addition, subcellular colocalization of PDI and
PACMA 57 was determined in OVCAR-8 cells using confocal microscopy (Figure
11B). To evaluate whether the parent PACMA 31 binds to the same site in PDI
as its fluorescent analog 57, we performed a competition assay using purified
PDI protein. PACMA 31 pretreatment blocked PDI protein from binding PACMA
57 (Figure 11C), demonstrating that the conjugation with BODIPY moiety does
not change PACMA 31’s target site. PACMA 57 binds to PDI protein in a time-
dependent manner (Figure 11D). The presence of DTT considerably increased
this interaction (comparing lane 9 to lane 2), indicating that PACMA 57 targets
cysteine residues. Additionally, the presence of urea also substantially increased
this interaction, indicating that not all the free sulfhydryl groups are accessible to
PACMA 57 in the PDI protein in native conformation. We further evaluated the
selectivity of the active analogs for PDI. After incubating PACMA 57 with an
equal amount of PDI, BSA, or GRP78 core domain (another important molecular
chaperone within the ER), we demonstrated that PACMA 57 selectively binds to
PDI, whereas no detectable fluorescence was observed from PACMA 57-treated
BSA or GRP-78 core domain (Figure 11E). Together, these results indicate that
the active PACMAs selectively target and covalently bind to PDI.
69

Figure 11. Confirmation of PDI as the cellular target of PACMAs. (A) Immunoprecipitated
cellular PDI was covalently bound by 57. OVCAR-8 cells were incubated with 57, 58 and 59 at 2
µM for 30 min. Whole cell lysates were subjected to immunoprecipitation using monoclonal anti-
PDI antibody. Precipitated proteins were analyzed by SDS/PAGE and scanned for BODIPY
fluorescence (upper panel), followed by Western blotting with anti-PDI antibody (lower panel). (B)
Subcellular co-localization of PDI and 57. OVCAR-8 cells were treated with 2 µM of 57 for 30 min,
followed by fixation and permeabilization. PDI was stained with anti-PDI mAb. Subcellular
localization of PDI and 57 was analyzed using confocal fluorescence microscopy. Red, PDI.
Green, 57. Yellow, merge. One of five representative microscope fields is shown. (C) Competition
between 57 and 31 on PDI. 100 ng/µL of recombinant PDI protein was incubated with 100 µM of
31 or DMSO in sodium phosphate buffer (pH 7.0) for 1 hr at 37°C, followed by 1 hr incubation
with 20 µM of 57. Solutions were subsequently mixed with 5✕ SDS sample buffer and analyzed
by SDS/PAGE, fluorescence scanning (upper panel) and Coomassie Blue stain (lower panel). (D)
Kinetics study of covalent interaction between PDI and PACMA 57. 100 ng/µL recombinant PDI
was incubated with 20 µM PACMA 57, 8 M Urea, or 100 mM DTT for indicated time at 37 °C,
followed by analysis using SDS/PAGE, fluorescence scanning (upper panel) and Coomassie Blue
staining (lower panel). One of two representative experiments is shown. (E) 57 selectively bound
to PDI. 20 µM of 57 was incubated with 100 ng/µL recombinant PDI (lane 1, with 100 ng/µL BSA
as a carrier protein), BSA (lane 2), or the core domain of GRP78 (lane 3) for 30 min at 37 °C,
followed by analysis using SDS/PAGE, fluorescence scan (upper panel) and Coomassie Blue
stain (lower panel).

70
To identify the precise cysteine residues in PDI that form covalent bonds with
31, on-line LC-Orbitrap CID and ETD MS/MS were used (workflow for sample
preparation and data analysis is shown in Figure 12A). Addition of 31 led to a
defined mass change of one peptide derived from the digestion of recombinant
PDI protein that was directly measured by high resolution MS with high
confidence (Figure 12B). This peptide fragment contains PDI’s active site
cysteines: C(397)GHC(400). The mass shift suggests that recombinant PDI was
modified by 31 at either Cys397 or Cys400. Interestingly, Cys397 and Cys400
were not modified simultaneously, presumably due to steric hindrance caused by
the binding of a PACMA 31 molecule. CID and ETD fragmentation MS/MS were
further utilized to localize the modification site with single amino acid resolution.
Detection of precursor ions at high resolution and a nearly complete series of
fragmentation ions from both CID (Figure 12C) and ETD (Figure 12D) allowed
the accurate sequencing and assignment of the modification site to
Cys397/Cys400. Integrating all the CID and ETD results, Protein Discoverer 1.3
automatically assigned potential modification sites at either Cys397 or Cys400
with high confidence (Figure 12E and F).
71

Figure 12. Determination of PACMA 31 binding site in PDI. (A) Workflow for sample
preparation and data analysis. B, High resolution mass spectra acquired in Orbitrap for
accurate mass shift identification of tryptic peptides of PDI before and after 31 binding.
Based on the selection of precursor ions of 762.9855 (M
+3
/3), further (C) CID and (D)
ETD fragmentation MS/MS were used for accurate and consistent sequencing and
localization of 31 binding site. (E) The Protein Identification Details view from Proteome
Discoverer 3.1 shows identified PDI sequence (green with high confidence) and (F)
potential PACMA 31 binding sites. c: cysteine modification with PACMA 31; C:
carbamidomethylation of cysteine.
72
When 31 was docked on Cys397 of PDI using PDB structure 3UEM we
obtained a fitness score of 40.73 forming a covalent bond between the terminal
carbon atom of 31’s propynoic moiety and the sulfur atom of Cys397. We also
observed two additional π−π interactions: one between 31’s phenyl ring in R
1
and
PDI’s Trp396, and the other between 31’s thienyl ring (R
2
) and PDI’s Phe304
(Figure 13A). When 31 was docked on Cys400 of PDI we obtained a fitness
score of 33.53. PACMA 31 bound PDI through a covalent bond with Cys400 on
the opposite side of the Cys397 site (Figure 13B). In addition, 31’s amide (-NH)
formed a hydrogen bond with an oxygen within Pro395. There is also an extra π-
cation interaction between 31’s phenyl ring in R
1
and a side chain nitrogen of
Lys401.

Figure 13. PACMA 31 covalently binds to Cys397/Cys400 in PDI active site.
Covalent docking of 31 (red) against PDI (blue, PDB: 3UEM) with a covalent bond
between the terminal carbon atom of 31’s propynoic moiety and the sulfur atom of (A)
Cys397 or (B) Cys400. GOLD fitness values are 40.73 and 33.53 for Cys397 and
Cys400, respectively.
73
III-1-5: Active PACMAs Affect PDI Secondary Structure and Inhibit PDI
Activity
Circular dichroism (CD) spectroscopy was performed to examine whether
covalent binding of the active PACMAs to PDI would affect its secondary
structure. Based on the CD data analysis using the K2D2 web server
(http://www.ogic.ca/projects/k2d2/) (Perez-Iratxeta and Andrade-Navarro, 2008),
PACMAs 57 and 31 affected the secondary structure of PDI, whereas no
substantial difference was observed between the spectra of recombinant PDI
treated with vehicle control DMSO and the inactive analog 56 (Table 5, Figure
14A). No substantial changes were observed in the secondary structure of the
control protein BSA treated with 31, 56, or 57 (Table 5, Figure 14B), indicating
that covalent binding of active PACMAs to PDI affects its secondary structure.

Table 5. Active PACMAs affected PDI secondary structure
% Secondary
structures
DMSO 31 57 56
PDI
α 42.60 47.81 36.77 42.60
β 9.77 8.18 12.31 9.77
BSA
α 84.27 84.27 84.27 84.27
β 1.24 1.24 1.24 1.24
74

Figure 14. Active PACMAs alters PDI secondary structures. Representative circular
dichroic spectra of (A) recombinant PDI and (B) BSA after treatments with 56, 31, and
57 for 1 h at 37°C.

75
Changes in protein structure are usually associated with variations in protein
activity. We therefore examined the effect of PACMAs on PDI’s activity with or
using the insulin aggregation assay, a well-established assay for evaluating PDI’s
activity (Khan et al., 2011). PACMA 31 significantly inhibited PDI’s activity in a
dose- and time-dependent manner, producing complete inhibition at 100 µM
(Figure 15A). Direct comparison of 31 and phenylarsine oxide (PAO, a previously
reported small-molecule PDI inhibitor (Root et al., 2004)) (Figure 15B)
demonstrated that 31 (IC
50
of 10 µM) is a more potent PDI inhibitor than PAO
(IC
50
of 85 µM). On the other hand, 56 was inactive as expected (Figure 15C).
These results demonstrate that covalent binding of the active PACMAs to PDI
inhibits its enzymatic activity.

Figure 15. Active PACMAs inhibit PDI activity. (A) 31 significantly inhibited PDI’s
activity in a dose-dependent manner in the insulin aggregation assay. Curves were
generated from mean values. BAR, SEM. *, P < 0.05; **, P < 0.01; ***, P < 0.001. (B)
Comparison of the inhibitory activity of 31 and PAO. (C) 56 did not exhibit significant
effects on PDI’s enzymatic activity. Experiments were performed in triplicate.

76
Although PDI has been known to play an important role in cancer progression
(Fonseca et al., 2009; Goplen et al., 2006; Lovat et al., 2008; Townsend et al.,
2009), it may be cancer and cell type specific. Therefore, we evaluated the
viability of human ovarian cancer cells by silencing PDI. PDI siRNA substantially
down-regulated PDI expression in OVCAR-8 cells between 24 and 96 h (Figure
16A) that was consistent with the significant inhibition of OVCAR-8 cell growth in
the MTT assay (Figure 16B). Additionally PDI knockdown significantly inhibited
colony formation by OVCAR-8 cells (Figure 16C). Consistently, PACMA 31
significantly inhibited colony formation in OVCAR-8 cells in a dose-dependent
manner (Figure 16D). These results indicate that PDI knockdown is sufficient to
cause considerable cytotoxicity and may account for the cytotoxicity caused by
the active PACMAs.
77

Figure 16. Silencing of PDI inhibits cell growth of OVCAR-8 cells. (A)
Representative Western blot of 24-96 h silencing of PDI in OVCAR-8 cells. (B) PDI
siRNA showed significant cytotoxicity as measured by MTT assay. Histogram shows
mean values of growth inhibition (%). (C) Silencing of PDI significantly inhibited colony
formation in OVCAR-8 cells. Histogram shows mean number of colonies. (D) 24 h
treatment of 31 significantly inhibited OVCAR-8 colonies at indicated doses. Histogram
shows mean number of colonies. BAR, SEM. **, P < 0.01; ***, P < 0.001.
78
III-1-6: PACMA 31 Suppresses Tumor Growth in Human Ovarian
Cancer Mouse Xenografts.
To evaluate the tumor targeting ability of active PACMAs in vivo, we tested 57-59
in a mouse xenograft model of human OVCAR-8 ovarian cancer. After 3 day
continuous i.p. administration, tumor, liver and brain tissues were collected for
frozen sections that were analyzed using fluorescent microscopy. PACMA 57-
treated tumor sections exhibited strong fluorescence intensity, whereas no
fluorescence was detected in 58 or 59-treated tumor sections (Figure 17A).
Consistent with our in vitro data, one fluorescent band at ~57 kD was observed
only in the 57-treated tumor sample (Figure 17B), confirming that PDI is
covalently modified by 57 in the tumors. Since liver is the major organ for drug
metabolism, we also examined the fluorescent intensity in the liver of 57-59-
treated animals by fluorescence microscopy utilizing the same settings and
conditions used to examine the tumor sections. Compared with 57-treated tumor
sections, the liver sections of 57-treated animals exhibited substantially lower
fluorescent intensity (Figure 17A). In comparison, liver sections from 58-treated
animals exhibited higher fluorescent intensity than liver sections from 57-treated
animals. No fluorescence emission was detected in 59-treated liver sections as
expected. In addition, no fluorescence emission was detected in brain sections
with treatments of active or inactive compounds. Taken together, these data
demonstrate that the active PACMAs selectively target and accumulate in
ovarian tumors in vivo, and do not cross the blood-brain barrier.
79

Figure 17. PACMA 57 targets in vivo ovarian tumor via binding PDI. (A) Tumor
targeting ability of PACMA 57. Mice were treated with 57, 58 and 59 at 10 mg/kg for 3
days with two injections per day. 6 h after the last injection, tumor, liver, and brain
tissues were collected and frozen sections were prepared for fluorescent microscopy
analysis. (B) PACMA 57 binds PDI in in vivo ovarian tumor. Homogenized tumor
samples from the 57- and 58-treated mice were analyzed using Gel fluorescence and
Coomassie blue stain. A ~57 kDa fluorescent band in the 57-treated tumor is indicated
by a red arrow.

80
To determine the in vivo efficacy of 31, we tested its effect on established
tumors from OVCAR-8 cells via i.p. or p.o. administration (Figure 18A). 31 was
given i.p. at 20 mg/kg/day for the first three weeks, with 5-day on and 2-day off
treatment cycles. The dose was escalated to 40 mg/kg/day for the next 7 days.
After 30-day i.p. treatment, the mouse xenografts were left untreated for
additional 32 days. A second xenograft study was conducted to evaluate p.o.
administration of 31. Treatment was initiated with a dose of 20 mg/kg/day and
was gradually increased by 20 mg/kg/day with each dose for 3 days before it was
orally dosed at 200 mg/kg per day for additional 32 days. The total duration of the
oral treatment was 62 days. Compared with the control group, i.p. or p.o.
administration of 31 significantly inhibited tumor growth by 85% (from 796.6 mm
3

to 117.0 mm
3
, P = 0.009) and 65% (from 796.6 mm
3
to 280.1 mm
3
, P = 0.015) at
day 62, respectively (Figure 18 B). Thus, 31 not only suppresses tumor growth in
vivo, but also is orally bioavailable. One mouse was found dead on day 30 in the
i.p. administration group, but no obvious abnormalities were observed in other
31-treated mice. The tumors in the i.p. treatment group did not aggressively grow
after treatment was stopped on day 30. This may be at least partially due to 31’s
prolonged duration of drug action, a common feature of irreversible inhibitors.
Compared with mice in the control group, no substantial body weight loss was
detected in i.p. or p.o. groups during the study (Figure 18C), indicating that 31 did
not exert severe adverse effects on the mice at its effective anticancer dose. All
mice were dissected at the end of the study (Day 62). H&E staining of tumor
sections showed extensive areas of necrosis in both 31-treatment groups (i.p.
81
and p.o.), compared to the control group (Figure 18D). In addition, no detectable
abnormalities were observed in the organs examined, including liver, kidney,
spleen, heart, lung, and pancreas (Figure 19), further demonstrating the safety of
PACMA 31.
82

Figure 18. PACMA 31 suppresses tumor growth in a mouse xenograft model of human
OVCAR-8 ovarian cancer. (A) Treatment schedules of PACMA 31 for mouse xenografts with
human OVCAR-8 tumor. I.p. administration of PACMA 31 (red areas) was dosed i.p. at 20
mg/kg/day for the first three weeks, with 5-day on and 2-day off treatment cycles. The dose was
escalated to 40 mg/kg/day for the next 7 days. After 30-day i.p. treatment, the mouse xenografts
were left untreated for additional 32 days. A second xenograft study was conducted to evaluate
p.o. administration of PACMA 31 (yellow areas). Treatment was initiated with a dose of 20
mg/kg/day and was gradually increased by 20 mg/kg/day with each dose for 3 days before it was
orally dosed at 200 mg/kg per day for additional 32 days. The total duration of the oral treatment
was 62 days. (B) Growth curves of s.c. tumors in mice treated with 31 through i.p. (red; n = 4) or
per os administration (yellow; n =4) or treated with vehicle (blue; n = 5). Results are presented as
mean tumor volume (BAR, SEM). (Inset) Comparison of tumor volumes between control and 31
i.p. (**, P < 0.01) or 31 per os treatment group (*, P < 0.05) on day 62 (BAR, SEM). (C)
Comparison of body weight of control mice and 31-treated mice. (D) PACMA 31 treatment
induced extensive areas of necrosis in OVCAR-8 tumors. Representative images of H&E-stained
tumor sections from control (Left) and 31-treated (Center, i.p.; Right, per os) mice are shown.
Arrows indicate areas of necrosis.
83
Figure 19. Histochemistry analysis of organs from OVCAR-8 xenograft mice in the control
group and in PACMA 31 treatment groups.
84
III-‐2:
DISCUSSION

In this study, we designed and synthesized a series of PACMAs that showed in
vitro and in vivo anticancer activity in human ovarian cancer by targeting PDI. As
a prototype of the ER protein disulfide isomerase family, PDI catalyzes the
formation, cleavage and rearrangement of disulfide bonds and facilitates
oxidative protein folding by acting as a molecular chaperone (Noiva, 1999).
Therefore, PDI inhibition causes accumulation of unfolded or misfolded proteins
that leads to ER stress and the unfolded protein response (UPR) and results in
cell death (Lovat et al., 2008). Compared with normal tissues, PDI is
overexpressed in ovarian tumors (Bonome et al., 2008). In addition, estrogen has
been reported to increase PDI expression (Ejima et al., 1999), to bind to PDI
(Primm and Gilbert, 2001), and to promote ovarian cancer progression (Spillman
et al., 2010). Based on these evidences, PDI is a novel and promising drug target
for ovarian cancer therapy.
We demonstrated that inhibition of PDI by either small molecule compounds or
PDI siRNA resulted in substantial cytotoxicity in human ovarian cancer cells.
Previously, it was shown that PDI knockdown results in apoptosis in human
breast cancer MCF-7 cells and cell death in human neuroblastoma SH-SY5Y
cells (Hashida et al., 2011). Moreover, a PDI inhibitor, bacitracin, abrogated
survival responses to ER stress and enhanced apoptosis caused by ER stress
inducing agents, fenretinide and velcade, in human melanoma CHL-1, A375 and
WM266-4 cells (Lovat et al., 2008). In addition, both bacitracin and a PDI
monoclonal antibody can inhibit migration and invasion of glioblastoma cells
85
(Goplen et al., 2006). However, PDI’s role in apoptosis might be cell type- and
tissue-dependent. It was documented that PDI knockdown by PDI siRNA has no
significant effects on the viability of human cervical cancer Hela cells (Hashida et
al., 2011). Additionally, PDI inhibitors were shown to suppress apoptosis induced
by misfolded proteins in PC12 cells, although toxicity was observed with
complete inhibition of PDI at high concentrations of inhibitors and knockdown
with shRNA (Hoffstrom et al., 2010). Therefore, PDI could be exploited as an
important target for developing drugs against different cancers.
Irreversible drugs have proved to be successful therapies for various
indications (Singh et al., 2011). Irreversible inhibitors often contain reactive
functional groups such as aldehydes, haloalkanes, alkenes, Michael acceptors,
phenyl sulfonates, or fluorophosphonates, that usually covalently modify an
enzyme’s serine, cysteine, threonine or tyrosine residues with amino acid side
chains containing nucleophiles such as hydroxyl or thiol groups (Lundblad, 2004).
To-date, a large number of irreversible inhibitors have been reported to exhibit
anticancer activities. Alpha-difluoromethylornithine (DFMO), an irreversible
inhibitor of ornithine decarboxylase, is under clinical trials for chemoprevention of
skin, cervical, colorectal, esophageal, and prostate cancers (see
http://www.ClinicalTrials.gov/). Several covalent EGFR inhibitors such as HKI-
375, HKI-272, EKB569, BIBW2992 and PF299804(Janne et al., 2011) targeting a
unique cysteine 797 residue located at the lip of the EGFR ATP binding site are
currently being tested in clinical trials for a variety of cancers (Zhou et al., 2011a).
Similar to the binding of irreversible inhibitors, post-translational modifications of
86
PDI also suppress its enzymatic activity. Nitrosative stress-induced S-
glutathionylation of PDI inhibits its enzymatic activity, leading to the activation of
UPR and cell death in human ovarian cancer SKOV3 cells (Townsend et al.,
2009). Additionally, NO-induced S-nitrosylation of PDI down-regulates its
enzymatic activity, resulting in the accumulation of polyubiquitinated proteins and
the activation of UPR in neurodegenerative disorders (Uehara et al., 2006).
In summary, by using 2DGE coupled with mass spectrometry, we identified
and demonstrated PDI as a cellular protein target for PACMAs. We also showed
that PDI knockdown in human ovarian cancer cells was cytotoxic, and our
irreversible PDI inhibitors exhibited both in vitro and in vivo anticancer activity in
human ovarian cancer models with tumor targeting ability and no substantial
toxicity to normal tissues. Moreover, PACMAs were effective on human ovarian
cancer cell lines resistant to conventional chemotherapy. Resistance to first line
therapy occurs in all ovarian cancer patients and is a major cause of mortality.
Therefore, development of effective and safe PDI inhibitors as anticancer agents
may overcome the current treatment failure in ovarian cancer therapy.
87
III-‐3:
EXPERIMENTAL
PROCEDURES

Cell Culture
OVCAR-8 and OVCAR-3 cells (National Cancer Institute) were maintained in
RPMI-1640 supplemented with 10% heat-inactivated FBS (Gemini-Bioproducts).
NCI/ADR-RES cells, resistant to paclitaxel and doxorubicine (National Cancer
Institute), were maintained in RPMI-1640 supplemented with 10% heat-
inactivated FBS and 5 mM L-glutamine. HEY cells, naturally resistant to cisplatin,
were kindly provided as a gift by Dr. Louis Dubeau (University of Southern
California) and maintained in DMEM supplemented with 10% heat-inactivated
FBS and 5 mM L-glutamine. Cells were grown as monolayer at 37°C in a
humidified atmosphere of 5% CO
2
. For subculture and counting, cells were
washed with PBS without calcium or magnesium, incubated with 0.25% trypsin-
EDTA solution (Mediatech, Inc.) for 5-10 min, resuspended with culture medium
and centrifuged. All experiments were performed using cells in exponential
growth. Cells were routinely checked for Mycoplasma contamination by using
PlasmoTest (InvivoGen).
Preparation of Whole Cell Lysate
Attached OVCAR-8 cells were washed with ice-cold PBS, and lysed in cold lysis
buffer (20 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, pH 7.5)
with 1x protease and phosphatase inhibitors. Whole cell lysates were centrifuged
for 10 min at 13,000 rpm at 4°C, and then supernatant was collected.
88
Protein Identification by Two-Dimensional Isoelectric Focusing/SDS/PAGE
And Mass Spectrometry Sequence Analysis
Whole cell lysate from OVCAR-8 cells was subjected to two-dimensional
IEF/SDS/PAGE analysis, as described by O’Farrell (O'Farrell, 1975), with
modifications as described by Duncan and Hershey (Duncan and Hershey, 1984)
to promote spot focusing. 2D gels were scanned for BODIPY fluorescence (λ
ex
:
488 nm, λ
em
: 526 nm) using a Typhoon 8610 Imager and subjected to silver
stain. The fluorescently tagged target (~ 57 kDa) was excised from the gel and
the protein was subjected to tryptic in-gel digest. Tryptic peptides were isolated
from the gel, and subjected to sequencing and subsequent protein identification
by tandem LC/MS/MS using an LTQ XL (Thermo Fischer) linear ion trap mass
spectrometer as described (Wang et al., 2008b).
Immunoprecipitation
200 µL whole cell lysate from OVCAR-8 cells was incubated with anti-PDI (RL90)
mAb (1: 200, Stressgen, ALX-804-012-R100, isotype: mouse) and 20 µL protein
A/G PLUS-agarose beads (Santa Cruz) with gentle rocking for 1 hr at 4°C. Beads
were washed with cell lysis buffer 5 times and boiled with 20 µL 2✕ SDS sample
buffer (125 mM Tris-HCL, pH 6.8, 4% SDS, 20% glycerol, 100 µM DTT, 0.02%
bromophenol blue) for 5 min, followed by centrifugation for 1 min at 14,000 g.
Supernatant was analyzed by SDS/PAGE, fluorescence scanning, and Western
blotting.
89
Western Blotting
Protein concentration of whole cell lysate was determined by BCA protein assay
kit (Thermo Scientific). Proteins were resolved in 10% SDS-PAGE and
electrotransferred to Immun-Blot PVDF membrane (Bio-Rad). After blocking with
5% milk in TBST, membranes were probed with monoclonal anti-PDI antibody
(Cell Signaling, #3501, isotype: rabbit), subsequently with horseradish
peroxidase-conjugated secondary antibody, and developed using Dura Extended
Duration Substrate (Thermo Scientific). Immunoreactive proteins were visualized
with Chemi-Doc System (Bio-Rad).
Fluorescence Confocal Microscopy
OVCAR-8 cells were grown on poly-L-lysine (Sigma-Aldrich) coated glass
coverslips and treated with indicated compounds for 1 h. Cells were fixed with
3.7% formaldehyde in PBS for 15 min at room temperature and permeabilized for
10 min with ice-cold 100% methanol at -20°C. After blocking with 5% goat serum,
slides were incubated with anti-PDI antibody (Stressgen, 1:200) overnight at 4°C,
followed by incubation with Cy5-linked goat anti-mouse secondary antibody
(1:1000) at room temperature for 1 hr. Images were collected on a Leica TCS SP
confocal system with 100x magnification.
PDI Expression and Purification
The expression vector of recombinant human PDI protein with an N-terminal His-
tag was a kind gift from Dr. Lloyd W. Ruddock (University of Oulu, Finland). PDI
expression and purification was performed as described (Lappi et al., 2004).
90
Briefly, protein production was carried out in E. coli strain BL21 (DE3) pLysS
grown in LB medium at 37°C, 200 rpm and induced at an A
600
of 0.3 for 3 h with
1 mM IPTG. Cells were pelleted by centrifugation (6500 rpm for 15 min). The
pellet was resuspended in 0.03 volume of buffer A (20 mM sodium phosphate
(pH 7.3)) and lysed by sonication. The cell debris was removed by centrifugation
(16,000 g for 30 min). The clarified supernatant was loaded onto a 1-mL-bed-
volume Ni-nitrilotriacetic acid (NTA) histidine-binding column (QIAGEN, Inc.)
equilibrated with 10-column volumes of buffer A. After loading, the column was
washed with 10-column volumes of 20 mM sodium phosphate, 50 mM imidazole,
0.5 M NaCl (pH 7.3) and then with 10-column volumes of buffer A. His-tagged
proteins were eluted using 20 mM sodium phosphate, 50 mM EDTA (pH 7.3).
The eluent was dialyzed in 100 mM sodium phosphate buffer pH 7.0 with 2 mM
EDTA.
siRNA Transfection
Subconfluent OVCAR-8 cells were transfected in 96-well plates with 80 nM PDI
or control siRNA according to the manufacturer’s protocol (Santa Cruz
Biotechnology). Protein expression was determined by immunoblotting 24, 48, 72
and 96 h after siRNA transfection. Cell viability was evaluated using a 3-(4,5-
dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay (Carmichael
et al., 1987) 48, 72, and 96 h after siRNA addition.
91
Growth Inhibition Assay
Cells were seeded in 96-well microtiter plates, allowed to attach overnight prior to
the indicated treatments. After 72 h, cells were incubated with 0.3 mg/ml MTT
(Amresco) for additional 3 h at 37°C. After removal of the supernatant, DMSO
was added to the wells and the absorbance was read at 570 nm. All assays
were performed in triplicate. Percentage of cell growth inhibition was expressed
as: (1-A/C) ×100% (A and C were the absorbance values from experimental and
control cells, respectively). Inhibitory concentration 50% (IC
50
) values were
determined for each drug from a plot of log (drug concentration) versus
percentage of cell growth inhibition. Standard deviation (SD) or standard error of
mean (SEM) was calculated based on the IC
50
values obtained from at least
three independent experiments.
Determination of PACMA 31 Binding Site on PDI Using LC/MS/MS Analysis.
As described in workflow (Fig. S6A), recombinant PDI was incubated for 3 h at
37°C with PACMA 31. To remove unreacted PACMA 31 and eliminate potential
artificial PACMA 31 modifications after reduction with DTT, sample was desalted
using Zeba Spin Desalting Columns (Thermo Scientific). Desalted sample was
denatured, reduced with DTT, alkylated with iodoacetamide, and digested with
trypsin as described previously (Zhou et al., 2011b). Negative control
experiments were conducted in the same manner using all reagents except the
PACMA 31 treatment.
92
Samples were analyzed using an LC/MS system consisting of an Eksigent
NanoLC Ultra 2D (Dublin, CA) and Thermo Fisher Scientific LTQ Orbitrap XL
(San Jose, CA). Briefly, peptides were separated in a 10 cm column (75 µm inner
diameter) packed in-house with 5 µm C18 beads on a Eksigent NanoLC Ultra 2D
system using a binary gradient of buffer A (0.1 % formic acid) and buffer B (0.1%
formic acid and 80% ACN). The peptides were loaded directly without any
trapping column with buffer A at a flow rate of 300 nL/min. Elution was carried out
at a flow rate of 250 nL/min, with a linear gradient from 10% to 35% buffer B in
95 min followed by 50% B for 15 min. At the end of the gradient, the column was
washed with 90% B and equilibrated with 5% B for 10 min. The eluted peptides
were sprayed into the LTQ Orbitrap XL. The source was operated at 2.1-2.25 kV,
with no sheath gas flow, with the ion transfer tube at 250°C. MS spectra in the
range of m/z 350–2000 were acquired in the Orbitrap at an FWHM resolution of
60,000 after accumulation to an AGC target value of 500,000 in the linear ion
trap with 1 microscan.
For peptide sequencing and modification site localization, the same precursors
were selected for fragmentation by CID or ETD, and fragment ions were
analyzed in the liner ion trap. The five most abundant precursor ions were
selected for fragmentation by CID and ETD with an isolation width of 3 Th. The
instrument was operated in data-dependent acquisition mode, whereby five CID
followed by five ETD data-dependent MS/MS scans succeeded the high
resolution MS scan. For all sequencing events, dynamic exclusion was enabled
93
to minimize repeated sequencing. Peaks selected for fragmentation more than
once within 60 s were excluded from selection (10 ppm window).
Proteome Discoverer 1.3 (Thermo Fisher Scientific) was used for protein
identification using Sequest algorithms. The following criteria were followed. For
MS/MS spectra, the variable modifications were PACMA 31 modification of
cysteine, carbamidomethylation of cysteine and oxidation of methionine with a
maximum of four modifications. Searches were conducted against Uniprot or in-
house customer database with PDI sequence. Up to two missed cleavages were
allowed for protease digestion and peptides had to be fully tryptic. MS1 tolerance
was 10 ppm and MS2 tolerance was set at 0.8 Da. Peptides reported via search
engine were accepted only if they met the false discovery rate of 1%. There was
no fixed cutoff score threshold, but instead spectra were accepted until the 1%
FDR rate was reached. Only peptides with a minimum of six amino acid lengths
were considered for identification. We also validated the identifications by manual
inspection of the mass spectra.
Molecular Docking
Molecular docking of PACMA 31 on PDI was performed in a two-step procedure
using a PDI crystal structure (PDB ID: 3UEM). PDI structure and PACMA 31
were prepared using Protein Preparation (Wang et al., 2009c) and LigPrep
(Jones et al., 2011) wizards, respectively, before PACMA 31 was docked at
Cys397 or Cys400 using Induced-Fit Docking (IFD) module (Schrodinger, LLC)
(Sherman et al., 2006). The software automatically generated a binding site
94
(based on the size and shape of PACMA 31) comprising amino-acid residues
surrounding Cys397 or Cys400. In the next step, covalent docking was
performed on the relaxed binding site using Genetic Optimization for Ligand
Docking (GOLD, Cambridge Crystallographic Data Center (CCDC)) (Siegel et al.,
2012). The covalent docking site was defined as residues from IFD (5 Å
surrounding the docked ligand) with a sphere of 15 Å radius and atoms involved
in covalent linking. Docking studies were performed using the standard default
settings with 100 GA runs on each molecule. For each GA run, a maximum of
100,000 operations were performed on a set of 5 groups with a population of 100
individuals. The annealing parameters were used as default cutoff values of 3.0
Å for hydrogen bonds and 4.0 Å for van der Waals interactions. When the top
three solutions attained root-mean-square deviation (rmsd) values within 1.5 Å,
docking was terminated. The docking interactions between PDI and PACMA 31
were quantified by Gold-Score (a scoring function used by the software) and a
dimensionless fitness value (a function of the intra- and intermolecular hydrogen
bonding interaction energy, van der Waals energy, and ligand torsion energy)
(Jones and Willett, 1995; Jones et al., 1997).
Spectroscopic Analysis of PDI
The effect of covalent interaction with PACMAs on PDI secondary structure was
examined by J-815 circular dichroic spectrometer using a rectangular 1 mm
cuvette. PDI protein (18 µM) was incubated with indicated PACMAs (200 µM) or
DMSO for 2 h, followed by dialysis in distilled water for 1 h at room temperature.
Native and modified PDI samples at the same concentration were maintained at
95
25°C while scanned in the far-UV region (190-260 nm) in three replicates. The
CD output was analyzed for secondary structures using K2D2 web server
(www.ogic.ca/projects/k2d2/) (Perez-Iratxeta and Andrade-Navarro, 2008).
Measurement of PDI Activity
PDI activity was assayed by measuring the PDI-catalyzed reduction of insulin in
the presence of DTT, thus measuring the aggregation of reduced insulin B chains
at 620 nm as described previously (Khan et al., 2011). Briefly, recombinant PDI
protein (0.4 µM) was incubated with indicated compounds at 37°C for 1 h in
sodium phosphate buffer (100 mM sodium phosphate, 2 mM EDTA, and 8 µM
DTT, pH 7.0). After this incubation, the modified recombinant PDI protein was
added to the reaction mixture consisting of DTT (500 µM) and bovine insulin (130
µM, Sigma). The reduction reaction was catalyzed by PDI at room temperature,
and the resulting aggregation of reduced insulin B chains was measured at 620
nm.
Mice
Athymic mice (Charles River Laboratories, Wilmington, MA) were fed ad libitum
and kept in air-conditioned rooms at 20 ± 2°C with a 12 h light-dark period.
Animal care and manipulation were in agreement with the USC institutional
guidelines and with the Guidelines for the Care and Use of Laboratory Animals.
96
In Vivo Tumor Xenograft Studies
OVCAR-8 cells in logarithmic growth phase from cell culture were implanted in
athymic mice (5 × 10
5
cells in 100 µL of PBS/mouse) under aseptic conditions.
Tumor growth was assessed by biweekly measurement of tumor diameters with
a Vernier caliper (length × width). Tumor volume was calculated according to the
formula: Tumor volume (mm
3
) = D × d
2
/2, where D and d are the longest and
shortest diameters, respectively. For i.p. administration, tumors were allowed to
grow to an average volume of 50 mm
3
. Mice were then randomly assigned into 3
groups: vehicle control (n=5), i.p. treatment with 31 (n = 4, 20 mg/kg/day for the
first three weeks, with 5-day on and 2-day off treatment cycles. Dose was
escalated to 40 mg/kg/day for the next 7 days.) and per os treatment of 31 (n = 4,
initial dose of 20 mg/kg/day was gradually increased by 20 mg/kg/day with each
dose for 3 days before it was orally dosed at 200 mg/kg per day for additional 32
days increasing dose from 20 mg/kg to 200 mg/kg). Treatment schedules are
described in Figure 18A. Treatment of each animal was based on individual body
weight. The body weights and tumor volumes in each group were measured
twice a week. The percentage of tumor growth inhibition was calculated as T/C%
= 100 × (mean tumor volume of treated group)/(mean tumor volume of control
group).
97
Histochemical Analysis
Upon autopsy, tumors, kidneys, livers, pancreas, spleens, hearts, and lungs were
removed, fixed in formalin, embedded in paraffin, and sectioned. Sections (4 µm)
were stained with H&E for histologic examination.
Statistical Analysis
The Student’s test was used for statistical analysis and P-value determination in
SPSS 16.0 software (SPSS Inc.) and Prism 5 (GraphPad Software, Inc).
Differences were considered statistically significant at P < 0.05.
98
CHAPTER IV: GP130 IS A PROMISING TARGET FOR
CANCER CHEMOTHERAPY
Ubiquitously expressed in the human body, gp130 is a shared subunit of receptor
complexes for at least nine cytokines (IL-6, OSM, LIF, IL-11, CNTF, CLC, IL-27,
CT-1 and NP) that mediate highly diverse biological processes. Dysregulation of
gp130 expression, activation or associated signaling pathways is related to the
development of a variety of human diseases, including cancer. However, the
roles of gp130 in the progression of many cancer types are still unclear, and
gp130-mediated signaling cascades triggered by different chemokine ligands can
result in distinct effects even on the same cancer type. Therefore, understanding
how gp130-mediated signaling pathways interface with various human cancers
may provide important insights into the development of novel gp130-targeting
agents with improved efficacy and reduced off-target toxicity for cancer therapy.
Herein, we review what is currently known regarding the relationship between
gp130 and human cancers.
99
IV-‐1:
INTRODUCTION

Cancer cells acquire multiple unique biological capabilities called the hallmarks of
cancer (Hanahan and Weinberg, 2000, 2011). Cancer progression can be driven
by the aberrant regulation of cellular signaling networks. Among them,
glycoprotein 130 (gp130) is one of the important molecules sensing and
transducing oncogenic signals. As a consequence, gp130 mediates the
development of multiple hallmarks of cancer, including cell-death resistance,
sustaining proliferation, angiogenesis, invasion and metastasis.
gp130 (also called IL6ST, IL6-beta or CD130) is a 130 kDa transmembrane
protein originally discovered on the surface of activated human T lymphocytes in
1978 (Andersson et al., 1978). It was after over a decade later that Taga et al
identified gp130 as a subunit of the IL-6 receptor complex (Taga et al., 1989) and
Hibi et al successfully cloned and expressed gp130 that substantially fostered
studies on gp130 (Hibi et al., 1990). Research during the past 20 years has
shown that gp130 serves as a shared signal transducing subunit of the receptor
complexes for at least nine human cytokines: interleukin-6 (IL-6) (Taga et al.,
1989), interleukin-11 (IL-11) (Yin et al., 1993), interleukin-27 (IL-27) (Pflanz et al.,
2004), leukaemia inhibitory factor (LIF) (Ip et al., 1992), ciliary neurotrophic factor
(CNTF) (Ip et al., 1992), oncostatin M (OSM) (Gearing et al., 1992),
cardiotrophin-1 (CT-1) (Pennica et al., 1995), cardiotrophin-like cytokine
(CLC/CLCF-1) (Elson et al., 2000), and neuropoietin (NP) (Derouet et al., 2004).
In addition, two viral homologs of IL-6 (vIL-6) from HHV-8 and the Rhesus
macaque rhadinovirus are also known to interact with human gp130 and elicit
100
cellular responses similar to cellular IL-6 (Kaleeba et al., 1999; Osborne et al.,
1999). Therefore, gp130 appears to be capable of recognizing various cytokine
structures in chemically unique fashions for each ligand.
In this chapter, I will review the biological properties of gp130-related cytokine
receptor complexes, their involvements in cancer progression, and approaches
to regulate their signaling for cancer therapy.
101
IV-‐2:
GP130
STRUCTURE

Several crystal and NMR structures of gp130 protein have been determined,
which provide substantial insights on its macromolecular topology and ligand
recognition mechanisms at the atomic level. The structural information deposited
in PDB databank is listed in Table 6. In addition, lower-resolution cryo-electron
microscopy (cryo-EM) studies have presented the architecture and dynamics of
the IL-6/IL-6R/gp130 and IL-11/IL-11R/gp130 (EMDB ID: 1223) receptor
complexes (Matadeen et al., 2007; Skiniotis et al., 2005), further extending our
knowledge of the gp130-containing receptor complexes.

102
Table 6. Available crystal and NMR structures of gp130

PDB
ID
Structure
Image
Method Resolution (Å) Sources Notes
3L5H

x-ray
diffraction
3.60
Homo
sapiens
Full ectodomain
(domains 1-6)
(Xu et al., 2010).
3L5J

x-ray
diffraction
3.04
Homo
sapiens
Fniii domains
(domains 4-6)
(Xu et al., 2010).
3L5I

x-ray
diffraction
1.90
Homo
sapiens
Fniii domains
(domains 4-6)
(Xu et al., 2010).
1BJ8

solution
NMR
The 50 Ca atoms in
the b-stands: 0.81. All
Ca atoms: 1.59. All
heavy atoms: 2.31.
Homo
sapiens
The third N-terminal
domain (D3) (Kernebeck
et al., 1999).
1PVH

x-ray
diffraction
2.50
Homo
sapiens
LIF (169 aa)/gp130-D2D3
complex (Boulanger et al.,
2003a).
1P9M

x-ray
diffraction
3.65
Homo
sapiens
The hexameric IL-6/IL-
6Ra /gp130-D1D2D3
hexamer (Boulanger et
al., 2003b).
1I1R

x-ray
diffraction
2.40
Homo
sapiens,
HHV-8
The HHV-8 IL-6/gp130-
D1D2D3 tetramer.
Domains 1, 2, 3 of the
extracellular domain were
expressed in the
presence of tunicamycin,
an inhibitor of N-linked
glycosylation (Chow et al.,
2001).
1BQU

x-ray
diffraction
2.00
Homo
sapiens
A cytokine-binding region
of gp130 (Bravo et al.,
1998).
103
Human gp130 protein contains a 597-amino acid residue ectodomain, a 22-
amino acid residue transmembrane domain and a 277-amino acid residue
cytoplasmic domain (Figure 20A) (Wang et al., 2009c). The available structures
show that the ectodomain of gp130 comprises six contiguous-sandwich domains
(D1 – D6). The first N-terminal domain (D1) is an Ig-like domain (IgD), which is
followed by one cytokine-binding homology region (CHR) module (D2 and D3).
Functional and structural studies of these membrane-distal domains (D1-D3)
have indicated that both IgD and CHR are responsible for ligand engagement
and necessary for the full activation of gp130-associated receptor complexes
(Chow et al., 2001; Horsten et al., 1995; Huyton et al., 2007; Skiniotis et al., 2005;
Skiniotis et al., 2008). In contrast to the membrane distal domains, the functions
of the three membrane-proximal fibronectin III-like (FnIII) domains (D4-D6) are
not clear. It was shown that individual deletion of the FnIII domains did not affect
the ligand binding capability of gp130 with IL-6 but abolished the receptor
response to IL-6 and IL-11, suggesting that the FnIII domains are involved in
receptor activation (Kurth et al., 2000). Electron microscopic (EM) studies
suggest that cytokine binding brings the FnIII domain “legs” to the correct
proximity and orientation on the cell surface that may instigate JAK trans-
phosphorylation in the cytoplasmic domains (Skiniotis et al., 2005). In addition to
the transmenbrane gp130 protein, at least three different forms of human soluble
gp130 (sgp130) have also been reported (50, 90 and 110 kDa, respectively)
(Narazaki et al., 1993; Tanaka et al., 2000; Zhang et al., 1998) that lack the
transmembrane and cytoplasmic domains but retain the extracellular ligand-
104
binding domains. Although the physiological role of sgp130 has not been further
determined, it was suggested to act as a natural inhibitor of gp130 signaling
(Mitsuyama et al., 2006; Richards et al., 2006).
Accumulating evidence from crystallographic, electron microscopic, and
biochemical studies has elucidated the role of receptor composition for cytokines
signaling via gp130. Based on the currently available data, models of gp130-
associated receptor complexes with respective cytokines are shown in Figure
20B. In the classical IL-6 signaling, IL-6 binds to its receptor complex composed
of homodimerized gp130 with a nonsignaling α-receptor IL-6R in a symmetric
hexameric complex (Boulanger et al., 2003b; Skiniotis et al., 2005), whereas
during trans-signaling, IL-6 interacts with a soluble form of IL-6 receptor (sIL-6R)
that is derived from alternative splicing or proteolytic cleavage of IL-6R, and
subsequently triggers the activation of gp130 signaling (Knupfer and Preiss,
2008). IL-6 trans-signaling severs as an alternative mechanism to trigger IL-6
related pathways in IL-6R-deficient cells. Another important difference between
the classical IL-6 pathway and the trans-signaling pathway is that only trans-
signaling can be inhibited by sgp130, which binds to IL-6/sIL-6R complex, without
significant affinity for IL-6 or IL-6R alone (Jostock et al., 2001). Unlike human IL-
6, viral IL-6 (vIL-6) forms a stable tetrameric complex with gp130 (vIL-
6:gp130:gp130:vIL-6) independent of but enhanced by IL-6R (Hu and Nicholas,
2006), which is probably due to hydrophobic enhancement of vIL-6’s receptor
binding epitopes (Boulanger et al., 2004). IL-11 signaling requires the association
of an α-receptor IL-11R with homo-dimerized gp130, where IL-11R is responsible
105
for high-affinity ligand binding and gp130 is required for signal transduction
(Barton et al., 2000; Matadeen et al., 2007; Nandurkar et al., 1996). LIF weakly
interacts with LIF receptor (LIFR), leading to the hetero-dimerization of gp130
with LIFR to enhance ligand-binding and signaling activation (Gearing et al.,
1991; Heinrich et al., 1998). CT-1 also utilizes the gp130/LIFR heterocomplex for
signaling through directly binding to LIFR but not gp130 (Pennica et al., 1995).
OSM signaling can be mediated by the gp130/LIFR or gp130/OSMR (an OSM-
specific receptor component) heterocomplexes (Gearing et al., 1992; Liu et al.,
1992; Murakami-Mori et al., 1995; Thoma et al., 1994). As a heterodimer
consisting of p28 and EBI3, IL-27 requires the association of gp130 and WSX-1
for receptor complex formation and signal transduction (Pflanz et al., 2004). The
binding of CNTF, CLC or NP to CNTFR, which is anchored to the cell surface
through a glycosylphosphatidylinositol (GPI) linker, results in the recruitment and
hetero-dimerization of gp130 and LIFR for signaling activation (Derouet et al.,
2004; Elson et al., 2000). As a result, differential association with cytokine
specific α-receptor subunits renders gp130 capable of mediating various cellular
events triggered by such diverse cytokines.
106

Figure 20. Composition of gp130 protein and gp130-associated receptor
complexes. (A) Comparison of structural domains of gp130 and sgp130. Both gp130
and sgp130 have IgD and CHR domains, which are involved in ligand binding. In
addition, gp130 protein has a FnIII domain, a transmembrane domain and a cytoplasmic
domain. (B) Composition of gp130-associated receptor complexes and cytokine ligands
recognized by respective receptor complexes.

107
IV-‐3:
THE
EXPRESSION
OF
GP130
CYTOKINES
AND
COMPONENTS
OF
THE

GP130-‐RELATED
RECEPTOR
COMPLEXES

gp130 mRNA is ubiquitously expressed in the human body, exhibiting highest
levels in saphenous vein, pericardium, ovary, omental adipose tissue,
peritoneum, spleen lymph nodes and trigeminal ganglia (Figure 21). However,
the cell surface-located gp130, which is the active and glycosylated form of
gp130, is not expressed in all cell types. For example, gp130 protein in bone
marrow-derived mast cells (BMMCs) is not present on the cell surface but
located intracellularly (Traum et al., 2011). Unlike gp130, other subunits and
ligands of the gp130 receptor complexes display highly restricted expression
patterns in tissues (Figure 22). IL-6 is mainly expressed in smooth muscle and B
lymphocyte, and IL-6R is present at relatively high levels in liver, neutrophil and
leukocyte. OSM is highly expressed in immune and digestive systems, and
OSMR is expressed in multiple systems, such as cardiovascular, urogenital, and
musculoskeletal systems. LIF is highly expressed in smooth muscle, superior
cervical ganglion, and atrioventricular node, and LIFR is highly expressed in
globus pallidus, substantia nigra reticulate, substantia nigra pars compacta in
nervous system. CT-1, another cytokine ligand of the gp130/LIFR receptor
complex, is widely expressed in many systems, such as nervous, cardiovascular,
musculoskeletal, and digestive systems. IL-11 is highly expressed in
atrioventricular node, cilary ganglion and superior cervical ganglion, and IL-11R
is highly expressed in T lymphocyte, heart atrium and aorta. Although CNTF and
CNTFR are coordinately expressed in nervous systems, CNTF is substantially
108
highly expressed in olfactory bulb, where CNTFR level is relatively low. CLC,
another cytokine ligand of the gp130/CNTFR receptor complex, is highly
expressed in immune system. Interestingly, the two subunits of IL-27 (p28 and
EBI3) do not show a correlated tissue expression. p28 is mainly expressed in
digestive system, whereas EBI3 is selectively and highly expressed in placenta.
Their specific α-receptor subunit, IL-27R, is mainly expressed in immune system.
So far, no biodistribution information about NP is available. Based on these data,
although gp130 serves as a common subunit of multiple receptor complexes, the
activity of individual gp130 receptor complexes in specific tissues or organs is not
only determined by the local abundance of the gp130 protein, but also by that of
other receptor complex subunits, the cytokines and even the downstream
signaling components. These combinations of different ratios further result in
distinct signaling activation and thus differential regulation of biological processes
in a tissue-specific manner.
109

Figure 21. Expression levels of gp130 in different tissues. Images were generated
using NEXTBIO (www.nextbio.com).
110

Figure 22. Expression levels of gp130 cytokines and other gp130 receptor
complex subunits in different tissues. Images were generated using NEXTBIO
(www.nextbio.com).
111
The cellular level of gp130 is tightly regulated by a variety of cellular processes
at transcriptional and post-translational levels. gp130 signaling can promote its
own transcription mediated by the binding of activated Stat1 and Stat3 homo-
and heterodimers to a STAT-binding element within the gp130 promoter (O'Brien
and Manolagas, 1997), demonstrating a positive autoregulatory loop of gp130
signaling. At the post-transational level, N-linked glycosylation was required for
stabilizing gp130 (Waetzig et al., 2010). Although the downregulation of gp130
has not been well understood, it has been reported that phosphorylation of
Ser782 induced by LIF, EGF, or PMA results in the internalization and
degradation of gp130 (Gibson et al., 2000). Ser782 can be directly
phosphorylated by MAPK-activated protein kinase 2 (MK2) (Radtke et al., 2010),
calmodulin-dependent protein kinase type II (CaMKII) and possibly CaMKIV
(Gibson et al., 2005), whereas okadaic acid (OA), a potent protein phosphatase
type 2A (PP2A) inhibitor, promotes this phosphorylation as well as the
degradation of gp130 (Mitsuhashi et al., 2005). In addition, a rapid translocation
of gp130 from cell surface to endosomal compartments is induced by IL-6,
followed by c-Cbl/SHP2 complex-dependent ubiquitination and lysosomal
degradation, whereas in the absence of IL-6 stimulation gp130 is degraded
through the proteasome-dependent pathway (Tanaka et al., 2008). Moreover,
CD95L-activated caspase 3 and 8 can cleave gp130 within amino acid residues
800-806 (DHVDGGD) of the cytoplasmic tail (Graf et al., 2008). It was also
documented that the internalization of gp130 did not require activation of the
Jak/STAT pathway (Thiel et al., 1998).
112
IV-‐4:
GP130
SIGNALING
IN
PHYSIOLOGICAL
PROCESSES

gp130 signaling is initiated when autocrine or paracrine cytokines induce the
dimerization of gp130 and an α-receptor subunit to form the receptor complex,
leading to the activation of downstream signaling cascades, such as the
JAK/Stat3, the PI3K/Akt, and the Ras/Raf/Mek/Erk1/2 MAPK pathways (Figure
23). The JAK/Stat3 pathway is tightly associated with gp130 activation. Following
the formation of the gp130 receptor complex, Janus kinase (JAK) family kinases
are recruited to cytoplasmic domain, where they are phosphorylated and
activated. JAK family kinases subsequently phosphorylate certain tyrosine
residues located in the cytoplasmic domains of the receptor complex to create
docking sites for cytosolic Stat proteins, mainly Stat3, via the SH2 domain. Once
recruited, Stat3 protein is phosphorylated at Tyr705 residues and is converted to
active forms. Activated Stat3 undergoes homodimerization (eg, Stat3:Stat3) or
heterodimerization (eg, Stat1:Stat3) and translocates into the nucleus, where it
regulates its target gene expression, resulting in cell growth and proliferation
(cyclin D, p21
WAF1/CIP2
, CDC25A, c-Myc, and survivin), survival (survivin, Bcl-2,
and Bcl-XL), angiogenesis (VEGF) and cell migration (MMP-2, MMP-7, and
MMP-9) (Debnath et al., 2012; Yu et al., 2009). Meanwhile, ligand-dependent
gp130 phosphorylation of Tyr759 triggers the engagement of the tyrosine
phosphatase SHP2 and activates the downstream PI3K/Akt and
Ras/Raf//Mek/Erk1/2 signaling pathways. Activation of phosphatidylinositol 3-
kinases (PI3Ks) leads to the phosphorylation and activation of Akt, which
regulates the activities of several target proteins, and therefore mediates cell
113
growth, proliferation, and survival via various cellular mechanisms (Vivanco and
Sawyers, 2002). In addition, SHP2 recruits GRB2/SOS complex that further
activates the small G-protein Ras. Activated Ras binds to Raf, resulting in the
subsequent phosphorylation and activation of Raf (MAPKKK), Mek (MAPKK),
and Erk1/2 (MAPK). Activated Erk1/2 has a variety of cytoplasmic and nuclear
protein substrates mediating many biological processes (Kolch et al., 2002). In
normal cells, and under physiological conditions, the activation of gp130-related
signaling is under tight control. Activated gp130 undergoes internalization,
polyubiquitination, and lysosomal degradation, resulting in the downregulation of
cell surface-bound gp130 (Tanaka et al., 2008). Downstream signaling pathways
of gp130 also have feedback regularion. For example, the JAK/Stat3 signaling
can be inactivated via multiple mechanisms. Nuclear protein-tyrosine
phosphatases such as TC-PTP and TC45 dephosphorylated Stat3 (Yamamoto et
al., 2002)
,
(Muromoto et al., 2008), causing nuclear Stat3 to shuttle back to the
cytoplasm (Herrmann et al., 2007). SHP1/2 dephosphorylate and inactivate JAK,
Stat3 and the receptor (Eiken et al., 2001; Freed et al., 2003). In addition, active
Stat3 binds to the promoter regions of SOCS genes, the products of which
compose another negative feedback mechanism regulating Stat3 and its
upstream kinases (Butcher et al., 2005; Naka et al., 1997; Stephanou et al.,
1998). SOCS1 and SOCS3 interact with the kinase domain of various JAK
proteins or the phosphotyrosine residues in the intracellular domain of the
receptor, hence suppressing Stat3 activation (Maeda et al., 2000; Weiss et al.,
2003; Wijelath et al., 1997). Protein inhibitor of activated Stat3 (PIAS3)
114
specifically blocks Stat3 dimers binding to DNA, thus abolishing the transcription
of Stat3 target genes (Junicho et al., 2000). Recently, the protein tyrosine
phosphatase (PTP) receptor T (PTPRT) has been reported to specifically
dephosphorylate Stat3 at Tyr705 and thereby regulate the expression of Stat3
target genes as well as the subcellular localization of Stat3 protein (Zhang et al.,
2007). Due to the fact that these feedback regulatory mechanisms are also tissue
specific, they further contribute to the diverse biological functions of gp130-
mediated signaling.
Based on our previous discussion, the various expression patterns of gp130
cytokines and receptor subunits, the different compositions of gp130-associated
receptor complexes, and the diverse regulations of gp130 expression and
signaling activation result in the multiple biological roles of gp130. Therefore,
gp130 mediates a broad range of cytokine-dependent physiological processes,
not limited to but including neuron protection (CNTF, CLC), aging (IL-6, CNTF),
development (IL-6, IL-11, OSM, CT-1, CNTF, CLC), acute phase response (IL-6,
OSM), immune response (IL-6, IL-11, OSM, IL-27, CLC), nerve response (IL-6,
OSM, CNTF), heart protection (IL-6, IL-11, CT-1), angiogenesis (IL-6, IL-11), liver
regeneration (IL-6), and metabolism (IL-6, CNTF) (Figure 22). It has been
reported that targeted deletion of the gp130 gene in mice results in lethality in
uteri or immediately after birth due to myocardial and hematological disorders
(Yoshida et al., 1996), which demonstrates the fundamental biological role of
gp130. Mouse models with conditional mutations or deletions in gp130-related
cytokines, gp130-associated receptor subunits, or molecules in gp130’s
115
downstream signaling pathways have been successfully developed (Fasnacht
and Muller, 2008). These mouse models are widely used in gp130-related
studies that further extend the knowledge of the physiological as well as
pathological properties of gp130 signaling in vivo.
116

Figure 23. gp130-mediated signaling pathways and physiological processes.
Ligand binding to its gp130-associated receptor complex leads to the activation of the
downstream JAK/Stat3, PI3K/Akt, and Ras/Raf/Mek/Erk1/2 signaling cascades. The
differential activation of these signaling pathways by different gp130 cytokines regulates
highly diverse physiological processes, not limited to but including neuron protection,
aging, development, immune response, nerve response, heart protection, angiogenesis,
liver regeneration and metabolism.

117
IV-5: DYSREGULATED
GP130-‐MEDIATED
SIGNALING
IN
CANCER
Due to the fact that gp130-mediated signaling is involved in a variety of
physiological processes, its dysregulation is associated with the development of
various diseases. Here, we are emphasizing in the role of gp130 in cancer
progression. Gene expression microarray studies provide an important tool for
evaluating gp130 expression levels in different cancer types. By analyzing
existing human cancer datasets in Oncomine
TM
, we found that gp130 expression
is significantly up-regulated (P < 10
-4
, fold change > 1.5) in select cancer types,
including brain tumor (Figure 24A) (2008; Shai et al., 2003; Sun et al., 2006),
kidney tumor (Figure 24B) (Beroukhim et al., 2009; Jones et al., 2005), myeloma
(Figure 24C) (Zhan et al., 2002), and teratoma (Figure 24D) (Korkola et al.,
2006). In addition, cytokines of gp130 and other components of the gp130
receptor complexes exhibit correlative upregulation with gp130 in select cancer
types. For example, in brain tumor, the expression levels of cytokines IL-6 and
LIF, and receptor components IL-6R, OSMR, LIFR and IL-27R are also
significantly increased (Figure 24E). Furthermore, gp130 expression is also
associated with cancer patients’ clinical outcomes. For example, we analyzed the
data from a TCGA large sample gene expression microarray study on brain
tumors, and found that patients with lower gp130 expression in brain tumors had
a 2-fold increase in their survival rate compared to those with high gp130
expressions in brain tumors (Figure 24F) (2008). Together, these results imply
that gp130 and its signaling networks may be associated with cancer
progression.
118

Figure 24. Dysregulation of the expression of gp130 in cancers. gp130 expression
is up-regulated in (A) brain tumor (2008; Shai et al., 2003; Sun et al., 2006), (B) kidney
tumor (Beroukhim et al., 2009; Jones et al., 2005), (C) myeloma (Zhan et al., 2002), and
(D) teratoma (Korkola et al., 2006). (E) Expression levels of cytokines and receptor
components of gp130 receptor complexes are up-regulated in correlation with gp130. (F)
Brain tumor gp130 expression level is associated with patients’ survival rate (2008; Shai
et al., 2003; Sun et al., 2006). Data sets were obtained from Oncomine
TM
(Compendia
Bioscience, Ann Arbor, MI), and analyzed using Prism 5 (GraphPad Software, Inc).
Student t-test was chose for statistical analysis. Box: 25% - 75%. Whiskers: Min and
Max.
119
In addition to gene expression microarray data, increasing evidence from
functional studies also support the involvement of each gp130-mediated cytokine
signaling in a vast variety of cancer types. Below we will discuss the relationship
of each gp130 receptor pathway and cancer based on the published literature.
IL-6/gp130 signaling
IL-6 was identified as a cytokine of the gp130/IL-6R receptor complex in 1989
(Taga et al., 1989). Among the different gp130 signaling pathways, the IL-
6/gp130 signaling is the most studied gp130-mediated signaling in cancer, with a
large body of published data indicating that abnormal activation of IL-6/gp130
promotes tumorigenesis and cancer progression in a variety of cancer types.
Myeloma. IL-6 was reported to promote the growth, proliferation and survival in
human myeloma cells via autocrine and/or paracrine mechanisms, with the
upregulation of both membrane and soluble forms of IL-6R and the slight
downregulation of cell surface-bound gp130 (Cheung and Van Ness, 2002;
Jourdan et al., 2005; Kovacs, 2003). IL-6 trans-signaling maintained the
proliferation of IL-6-dependent human multiple myeloma (MM) cell lines (Gaillard
et al., 1997). Compared to human IL-6, vIL-6 was less effective in triggering the
proliferation of myeloma INA-6 cells (Burger et al., 1998). IL-6/gp130/Stat3
signaling could be enhanced by beta1 integrin adhesion (Shain et al., 2009), and
synergized with CD9/HB-EGF/erbB1 pathways to promote growth and survival of
myeloma cells (Wang et al., 2002). Suppressing IL-6/gp130 signaling activation
using AR-42 (a phenylbutyrate-derived histone deacetylase inhibitor) (Zhang et
120
al., 2011), epitope peptides from IL-6R (Halimi et al., 1995), IL-6R
superantagonists (Demartis et al., 1996), AG490 (a JAK2 inhibitor) (De Vos et
al., 2000), or 18AD (an acidic peptide inhibiting Src family kinases) (Hausherr et
al., 2007), has been shown to inhibit IL-6/gp130-mediated cell growth,
proliferation and survival in human MM cells in vitro. In vivo, anti-human IL-6
monoclonal antibody (mAb) (MH166) and anti-human IL-6R mAb (PM1) exhibited
substantial inhibition of S6B45 tumor growth in a SCID mouse model, whereas
anti-human gp130 mAb (AM64) showed weak inhibition (Suzuki et al., 1992). In
addition, cell-cell contact of myeloma-derived ARH77 cells with bone marrow
stromal cells (BMSCs) activated the expression of IL-6 and gp130 genes in
BMSCs (Juneja et al., 2001). Mutations in the gp130 cytoplasmic domain were
reported in a human plasma cell line (HMCL) and in tumor plasma cells of one
patient, whereas DNA and RNA polymorphisms in the gp130 CHR domains were
detected in tumoral samples as well as in blood samples from healthy donors
(Rodriguez et al., 1994). Clinically, significantly high serum levels of sIL-6R were
detected in MM patients compared to normal controls and were correlated with
poor survival (Kyrtsonis et al., 1996).
Prostate cancer. As an autocrine and/or paracrine cytokine, IL-6 stimulated the
growth of human prostatic carcinoma cells in vitro, but did not exhibit significant
stimulatory effects on benign prostatic hyperplasia (BPH)-derived epithelial cells
(Lou et al., 2000; Okamoto et al., 1997b). Clinically, plasma IL-6 and sIL-6R
levels were significantly elevated in association with prostate cancer progression
and metastasis (Shariat et al., 2001). Immunohistocheical (IHC) analysis of
121
human prostatic biopsies showed that IL-6 was scarcely localized in the basal
cells of the epithelium whereas gp130 was detected exclusively in the stroma in
normal prostate. In benign prostatic hyperplasia, IL-6 preferentially localized to
the epithelium where as gp130 was detected both in the epithelium and in the
stroma. In prostatic carcinoma, IL-6 localized in all cell types, whereas gp130
was detected in the epithelium and in the stroma, and increased expression of
both IL-6 and gp130 was correlated with the Gleason grades (Royuela et al.,
2004). SNPs in gp130, IL-6 and Stat3 were found to be nominally associated
with the risk of prostate cancer in Caucasians (Kwon et al., 2011). In addition, IL-
6/gp130 signaling pathway partners with other pathways to foster prostate cancer
progression. It was reported that IL-6 cooperate with Fer to activate downstream
Stat3, resulting in human prostate cancer cell growth (Zoubeidi et al., 2009).
Prostaglandin E(2) (PGE(2)) stimulated prostatic intraepithelial neoplasia cell
growth was mediated by the IL-6/gp130/Stat3 signaling pathway (Liu et al., 2002).
IL-6 also increased androgen biosynthesis by activating the expression of genes
encoding steroidogenic enzymes including HSD3B2 and AKR1C3 in LNCaP cells
in vitro as well as in LNCaP-IL6(+) tumor in the prostates of castrated male nude
mice (Chun et al., 2009). In a transgenic mouse model of prostate
adenocarcinoma, genetic ablation of PKCε decreased IL-6 and gp130 expression
as well as Stat3 activation, accompanied by inhibition of cancer development and
metastasis (Hafeez et al., 2011). Although Stat3 signaling pathway is the major
downstream pathway of IL-6/gp130, IL-6 also documented to stimulate the
proliferation of prostate cancer 22Rv1 cells through the PI3-kinase/Akt pathway
122
without activating Stat3 (Godoy-Tundidor et al., 2005). Consistently, IL-6-
activated PI3-kinase prevented LNCaP cells from apoptosis (Chung et al., 2000).
In addition, IL-6/gp130 signaling also contributed to PC-3 cells’ resistance to
etoposide and cisplatin-mediated cytotoxicity, which most likely involved a
downstream Ras-dependent pathway (Borsellino et al., 1999). Moreover, there is
published data supporting the involvement of sgp130 in prostate cancer. In vitro,
sgp130 promoted invasion of androgen responsive prostate cancer cells and
induced a significant decrease in E-cadherin, whereas in prostate cancer patients
higher level of plasma sgp130, which was suggested to be produced by tumor
cells, was significantly associated with cancer invasion and progression, but
weakly correlated with plasma IL-6 and sIL-6R levels (Shariat et al., 2011). In
addition, it was also suggested that during prostate cancer progression from the
hormone-dependent stage to the hormone refractory stage, IL-6 underwent a
functional transition from paracrine growth inhibitor to autocrine growth stimulator
(Chung et al., 1999).
Colorectal cancer (CRC). IL-6 was shown to stimulate colony formation of both
primary and metastatic human CRC cells (Schneider et al., 2000). Clinically IL-6
and p-Stat3 were both highly expressed in ulcerative colitis (UC)-related CRC (Li
et al., 2010). More studies on the effects of IL-6/gp130 signaling were performed
using mouse CRC models. It was reported that gp130-mediated IL-6/Stat3
signaling activation was required for tumor survival and development in mouse
colitis-associated cancer (CAC) models (Bollrath et al., 2009; Grivennikov et al.,
2009). IL-6 trans-signaling plays an important role in tumor progression, which
123
could be suppressed by soluble gp130Fc in a colitis-associated premalignant
cancer (CApC) model (Matsumoto et al., 2010), or by TGF-beta in a CRC model
(Becker et al., 2004). Besides its role in trans-signaling that promotes CRC
progression in mouse models, sIL-6R was also suggested to mediate the
metastatic initiation of CRC (Dowdall et al., 2002).
Ovarian cancer. IL-6 is constitutively produced by ovarian cancer cell lines and
by primary ovarian tumor cultures that could be induced by reproductive
hormones and stimulated the proliferation of epithelial ovarian cancer cells via
gp130/Stat3 signaling (Syed et al., 2002; Wang et al., 2005; Watson et al., 1990).
Autocrine production of IL-6 in ovarian cancer cells also conferred resistance to
cisplatin and paclitaxel, associated with the increased expression of multidrug
resistance-proteins (MDR1 and GSTpi) and anti-apoptotic proteins (Bcl-2, Bcl-XL
and XIAP) as well as the activation of the Ras/MEK/ERK and PI3K/Akt signaling
pathways (Wang et al., 2010). Clinically, IL-6 levels were increased in the serum
and peritoneal fluid from ovarian cancer patients (Plante et al., 1994; Scambia et
al., 1994; Tempfer et al., 1997). Higher IL-6 levels in body fluids were not only
correlated with poor prognosis (Scambia et al., 1995), but also associated with
chemoresistance to paclitaxel (Penson et al., 2000).
Breast cancer. Addition of human recombinant IL-6 to cultured T47D cells
resulted in increased cell migration (Badache and Hynes, 2001). Anti-IL-6
antibody, IL-6 antisense ODN, and IL-6R antisense ODN have been shown to
significantly inhibit cell proliferation in MCF-7 cells (Jiang et al., 2011). In vivo,
MDA-231 cells expressing a dominant-negative gp130 protein (DN gp130) had
124
markedly decreased engraftment, tumor size, metastasis and angiogenesis in an
orthotopic nude mouse model (Selander et al., 2004). Significantly high levels of
serum IL-6 and sIL-6R are frequently detected in breast cancer patients that
positively correlate with the stage and the axillary lymph node involvement
(Kuang et al., 1998). The expression of IL-6, IL-6R and gp130 proved to be a
positive prognostic factor for patients’ overall survival (OS) and disease free
survival (DFS) (Karczewska et al., 2000).
Lung cancer. Human lung adenocarcinoma cells with somatic activating
mutations in EGFR produce high levels of IL-6 that contribute to tumorigenesis
via the activation of downstream Stat3 signaling (Gao et al., 2007). IL-6 was also
reported to function as an autocrine growth stimulator in vitro and in vivo for PG
cells (a high-metastatic human lung giant cell carcinoma cell line) that produced
both IL-6 and IL-6R (Fu et al., 1998). Additionally, interaction between lung
cancer HARA-8 cells and astrocytes via specific cytokines up-regulate the
expression of IL-6 and gp130 in HARA-8 cells and the production of IL-6, TNF-α
and IL-1β in astrocytes, resulting in cancer cell proliferation and perhaps brain
metastasis (Seike et al., 2011). Additionally, non-small cell lung carcinoma
(NSCLC) specimens from patients show significant tumor expression of IL-6, IL-
6R and gp130 (Haura et al., 2006), suggesting the contribution of IL-6/gp130
signaling to the pathogenesis of NSCLC.
Renal cell carcinoma (RCC). IL-6 plays an important role in RCC progression.
IL-6 and IL-6R expression in RCC were indicated as potential prognostic factors
due to their correlation with tumor size, stage, nuclear grade, proliferation index,
125
and patients’ overall survival (Costes et al., 1997). In addition, serum level of IL-
6, mainly released from RCC cells (Blay et al., 1994), was also indicated as a
prognostic factor in patients with metastatic RCC (Blay et al., 1992; Dosquet et
al., 1997; Yoshida et al., 2002). IL-6 exerts endocrine activities and inhibits
dendritic cell differentiation to mediate tumor cells escaping from immune
recognition (Menetrier-Caux et al., 1998). IL-6 also acts as an autocrine growth
factor in some RCC cell line. It was observed that IL-6 protected RCC cell lines
from cisplatin-induced apoptosis, and anti-IL-6 mAb or anti-IL-6R mAb sensitized
RCC cells to cisplatin (Mizutani et al., 1995). In addition, IL-6 may act through an
intracrine loop in some renal cancers. In eight RCC cell lines, IL-6 exerted a
consistent growth promoting activity in the absence of cell surface IL-6R, and an
IL-6 oligonucleotide but not a blocking IL-6 Ab suppressed RCC cell proliferation
(Alberti et al., 2004). These results indicate that IL-6 plays multiple roles in RCC,
and it might have similar effects in other cancers.
Kaposi’s sarcoma (KS). KS is a type of skin cancer associated with HHV-8
infection and is very common in AIDS patients (Sgadari et al., 2011). vIL-6 is
involved in in KS development. IL-6 and IL-6R produced by cells derived from KS
lesions of patients with AIDS (AIDS-KS cells) promote proliferation of KS cells
(Miles et al., 1990). In addition, vIL-6 could induce angiogenesis and
hematopoiesis by up-regulating VEGF expression (Aoki et al., 1999). Its soluble
receptor sIL-6R is a potent growth factor for AIDS-KS cells and could be
antagonized by sgp130 (Murakami-Mori et al., 1996). A synergistic effect was
reported between gp130 cytokines (IL-6/sIL-6R and OSM) and dexamethasone
126
on KS cell proliferation (Murakami-Mori et al., 1997). Also, vIL-6 but not human
IL-6 elicited proliferation and triggered a marked and sustained expression of an
acute-phase protein PTX3 in human primary KS cells, which were gp130-positive
and IL-6R negative (Klouche et al., 2002). DAB389-IL-6, a chimeric protein
linking IL-6 to a truncated diphtheria toxin, selectively exhibited cytotoxicity in IL-
6R-positive AIDS-KS cells, providing a potential therapy for KS treatment
(Masood et al., 1994).
Other cancers. Besides the cancer types we discussed above, where the role of
IL-6/gp130 has been relatively extensively studied, the IL-6/gp130 signaling was
also documented in the progression of other cancers. In inflammatory
hepatocellular adenomas (IHCAs), which are benign liver tumors, frequent
somatic mutations in gp130 and Stat3 resulted in hyperactivation of Stat3
signaling independent of IL-6 stimulation, and are accompanied by β-catenin-
activating mutations (Pilati et al., 2011; Rebouissou et al., 2009). Coexpression
of IL-6 and sIL-6R was reported to cause nodular regenerative hyperplasia and
adenomas of the liver in mice (Maione et al., 1998). IL-6 functioned as an
autocrine growth factor produced by bladder carcinoma cell lines (253J, RT4 and
T24) but not by normal urothelial cells, and stimulated the growth of the bladder
carcinoma cells substantially more effectively than that of normal urothelial cells
(Okamoto et al., 1997a). Downregulation of IL-6 and IL-6R expression using
short hairpin RNAs (shRNAs) significantly suppressed growth and invasion in
glioma stem cells (GSCs) and increased the survival of mouse xenografts with
intracranial human glioma, whereas upregulation of IL-6R and gp130 was
127
significantly associated with the decrease in survival of glioma patients (Wang et
al., 2009a). In a human choriocarcinoma cell line (JEG-3), monoclonal antibodies
against IL-6, IL-6R or gp130 failed to inhibit cell proliferation, whereas the
antisense oligonucleotides against IL-6 mRNA suppressed cell growth,
suggesting IL-6 may promote cell growth via an intracellular autocrine growth
mechanism (Kong et al., 1996), similar to the observation we discussed above in
RCC (Alberti et al., 2004). Different expression profiles of IL-6, IL-6R and gp130
genes were investigated in three acute leukemia subtypes, including acute
myeloid leukemia (AML), acute lymphoblastic leukemia (ALL), and acute mixed
lineage leukemia (AMLL) (Wang et al., 2009a). In AML signaling through IL-6R
promoted blast cell proliferation (Saily et al., 1998). Lymphoma cells infected with
HHV-8 secreted vIL-6 as an autocrine growth factor to support cell proliferation
and block the antiviral activity of host cell-induced IFN-α (Chatterjee et al., 2002).
Recombinant IL-6 promoted the in vitro growth of human medulloblastoma cells,
which are gp130- and IL-6R-positive but IL-6-deficient (Liu et al., 1995). In
malignant mesotheliomas, IL-6/sIL-6R/gp130/Stat3 signaling induced both cell
growth and VEGF production (Adachi et al., 2006), suggesting its ability to induce
tumor angiogenesis. In pancreatic cancer cells, IL-6 inhibited radiation-induced
apoptosis with the upregulation of Bcl-2 and Bcl-X
L
(Miyamoto et al., 2001). In
vitro functional studies showed that intratumoral IL-6 might contribute to
excessive growth hormone production in the majority of pituitary adenomas
(Thiele et al., 2003). In squamous carcinoma cells, IL-6/gp130/Stat3 signaling
supported cell survival via the upregulation of serpin B3/B4 (Ahmed and Darnell,
128
2009). For the middle-sized T antigen (PmT)-transformed murine endothelial
cells, IL-6 is an in vitro and in vivo autocrine growth factor (Giraudo et al., 1996).
Via an autocrine growth mechanism IL-6 was also shown to promote the
proliferation of human plasmacytoma cells mediated by gp130 (Nishimoto et al.,
1994). The expression of transfected IL-6 substantially accelerated the growth of
weakly tumorigenic rat urothelial cells, but failed to induce a tumorigenic
phenotype in non-tumorigenic cells (Okamoto and Oyasu, 1996).
IL-11/gp130 signaling
IL-11 was first isolated from bone marrow-derived stromal cells in 1990, and
identified as a gp130-related cytokine in 1993 (Yin et al., 1993). With a variety of
biofunctions overlapping with IL-6, IL-11/gp130 signaling is also indicated to
promote progression in multiple cancers, particularly gastric cancer.
Gastric cancer. Many cancer-related studies on IL-11 have been performed in
gastric cancer. IL-11 signaling induced gastric tumorigenesis through
downregulation of growth inhibitory signals, including members of the TGF-β
family (Jackson et al., 2007; Merchant, 2008). It has been suggested that IL-11
linked inflammation to cancer via Stat3 activation (Putoczki and Ernst, 2010). IHC
analysis of surgically resected human gastric adenocarcinomas showed that
increased IL-11 and IL-11R expression also correlated with tumor invasion,
which was confirmed by recombinant human IL-11-promoted migration in SCH
cell line in vitro (Nakayama et al., 2007). In a gp130(757F/F) mouse model, IL-11
level was considerably increased concomitant with gastric tumor progression
129
accompanied by Stat1/3 activation and increased MMP-9 and MMP-13
expression (Howlett et al., 2005), and IL-11 signaling was suggested to mediate
inflammation-associated gastric tumorigenesis and in an indirect manner, antral
tumor progression (Ernst et al., 2008; Judd et al., 2009).
Other cancers. Besides gastric cancer, IL-11 has also been documented to be
involved in several other cancers. In acute myelogenous leukemia cells, IL-11
was shown to enhance the clonal proliferation (Kimura et al., 1999). IL-11 also
enhanced invasiveness of human choriocarcinoma JEG-3 cells by increasing the
activation and expression of Stat3, whereas no significant effect was detected on
cell proliferation (Suman et al., 2009). By using gp130/Stat3 mutant mouse
models, IL-11/gp130 signaling was found to support cell survival and cell cycle
progression during colitis-associated colon cancer development (Bollrath et al.,
2009). IL-11 expression could be induced by IL-1β, phorbol ester, and calcium
ionophore in human glioblastoma U373 and U87 cell lines but not in
neuroblastoma SH-SY5Y cell line (Bollrath et al., 2009). IL-11/gp130 signaling
enhanced cell growth of human myeloma cell lines (Zhang et al., 1994). In
ovarian cancer, IL-11R and gp130 were commonly expressed in both malignant
and nonmalignant ovarian tissues, whereas IL-11 was only expressed in a few
malignant samples (Campbell et al., 2001a). Compared to nonmalignant prostate
samples, prostate carcinoma had elevated expression of IL-11 receptor complex
that resulted in constitutive Stat3 activation (Campbell et al., 2001b).
130
LIF/gp130 signaling
LIF was originally identified for its ability to induce differentiation in a murine
myeloid leukemia cell line and subsequently discovered to be an essential
mediator of embryonic development. Accumulating evidence suggests that LIF
signaling is also associated with select human cancers.
Breast cancer. LIF signaling is associated with breast cancer progression. Both
LIF and LIFR expression were up-regulated and associated with the
development of breast cancer (Garcia-Tunon et al., 2008). LIF triggered
proliferation of human breast cancer cell lines MCF-7 and SKBR-3, but did not
show no significant effects on normal mammary epithelial cell growth (Estrov et
al., 1995; Liu et al., 1998).
Other cancers. LIF's distinct outcomes on cancer cells are related to the
differences of cellular context. In addition, LIF promoted proliferation and survival
of human myeloma cells, accompanied by the activation of the ERK1/2, Stat3,
and PI3K/Akt/FKHR pathways (Lentzsch et al., 2004; Zhang et al., 1994). In
medullary thyroid carcinoma cells, LIF/JAK/STAT pathway induced growth arrest
and differentiation mediated by IL-1β (Park et al., 2005). In human
medulloblastomas, LIF was reported as an autocrine growth factor (Liu et al.,
1996). In human pancreas carcinoma cells and human plasmacytoma cells, LIF
also functioned as a growth factor (Kamohara et al., 2007; Nishimoto et al.,
1994). IHC analysis showed that LIF localized to the epithelium and the stroma in
normal prostate and benign prostatic hyperplasia, and to all cell types in prostate
131
carcinoma with increasing expression associated with the Gleason grades
(Royuela et al., 2004).
OSM/gp130 signaling
Secreted by macrophages, monocytes and activated T lymphocytes, OSM was
confirmed as a gp130-related cytokine in 1992 (Gearing et al., 1992). The OSM-
stimulated cellular signaling can be mediated by LIFR/gp130, which triggers
biological activities that overlap with those induced by LIF. In addition, OSM can
bind to OSMR/gp130 signaling complex, which elicits OSM unique functions that
cannot be stimulated by LIF or other gp130 cytokines and may be distinct from
those triggered by OSM/LIFR/gp130 signaling. One of OSM’s unique functions is
its tumor suppression effect in select cancer types. Below we will focus on
discussing its cytostatic effects in melanoma, brain and CNS, breast, and lung
cancers.
Melanoma. OSM’s growth-inhibitory activity was observed in melanoma A375
cells when it was first isolated from the supernatant of PMA-treated U-937 cells
(Zarling et al., 1986). As a result, in the early studies on OSM, A375 was used a
standard for testing its activity (Brown et al., 1987; Malik et al., 1989). It was
further indicated that OSM-induced growth inhibition of A375 cells was Stat3-
dependent and associated with upregulation of p27/Kip1 (Kortylewski et al.,
1999). In addition to A375, OSM also inhibited the growth of 7 of 11 cell lines
grown from various metastatic sites of melanoma patients, whereas in two of the
132
cell lines, OSM was observed to stimulate cell growth (Gibbs et al., 1998),
making the effects of OSM on melanoma complicated.
Brain and CNS tumors. Compared with IL-6, LIF, and CNTF, OSM is the most
potent activator of Stat molecules in a variety of brain tumor-derived cell lines,
including U373-MG, U118-MG, SNB-19, U87-MG, A172, and H4

(Schaefer et al.,
2000). Another study showed that OSM but not LIF or IL-6, in a side-by-side
comparison, inhibited the proliferation of glioma A172, GOS3, T98G, and U343
cells (Halfter et al., 1998; Halfter et al., 2000; Schaefer et al., 2000), and also
confirmed that activation of the JAK-STAT and MAPK pathways by OSM is not
sufficient to cause growth inhibition of human glioma cells (Halfter et al., 2000). In
addition, OSM/gp130/OSMR signaling inhibited glioblastoma 86HG39 cells from
proliferation with cell cycle arrest in G1 phase, and triggered astrocyte
differentiation by suppressing Skp2, Cks1 and cyclin A expression and up-
regulating p21 expression (Halfter et al., 2006). Similarly, in cerebral
meningiomas, recombinant OSM decreased meningioma cell growth by 66%,
whereas recombinant IL-6 or LIF had no significant growth modulating effects in
side-by-side comparisons (Schrell et al., 1998).
Breast cancer. OSM has also been reported to inhibit the proliferation of human
breast cancer cells. OSM induces growth arrest in vitro in breast cancer H3396,
H3630, H3680B, H3730, H3922, MCF-7 and ZR-75-1 cell lines that is associated
with the downregulation of c-Myc (Brown et al., 1987; Malik et al., 1989). OSM’s
effect on cell growth was also observed in normal human mammary epithelial
(HME) cells (Liu et al., 1998). It has been further confirmed that OSM’s inhibitory
133
activity in both normal and malignant mammary epithelial cell growth is mediated
by the OSM/OSMR/gp130 complex, not by the OSM/LIF/gp130 complex (Liu et
al., 1998). Further, the JNK pathway was identified to be specifically activated by
the OSM/OSMR/gp130 complex, not the OSM/LIFR/gp130 complex, leading to
the activation of downstream transcription factor (ATF)/cyclic AMP-responsive
element binding protein family member, ATF3 (Underhill-Day and Heath, 2006).
Lung cancer. An early study screened the effects of recombinant OSM in a large
panel of human cancer cell lines (Horn et al., 1990). It was found that OSM
inhibited the growth in six out of seven lung cancer cell lines tested. The six
responding cell lines are Calu-1, H125, SK-LU-1, A549, H2981, and SK-MES-1,
and the non-responding one is 9812, whereas OSM bound to all seven cell lines.
Further study using Calu-1 cells indicated that OSM, not LIF, IL-6 or IL-11,
induced the expression of tissue-type plasminogen activator (tPA) and
plasminogen activator inhibitor-1 (PAI-1) largely dependent upon activation of the
MEK1/2 pathway, whereas the Jak3/Stat3 pathway played a secondary role in
these regulatory events (Spence et al., 2002).

On the other hand, OSM shares downstream signaling pathways with other
gp130 ligands. Particularly, making use of the LIFR/gp130 complex for signaling,
OSM and LIF share some functional similarities. Like IL-6, IL-11 and LIF, OSM
signaling has been reported to promote the progression in multiple cancer types,
particularly prostate cancer.
134
Prostate cancer. IHC analysis of human prostatic biopsies showed that both
OSM and OMSR were detected in the stroma and LIFR was scarce in normal
prostates. In benign prostatic hyperplasia, OSM preferentially localized to the
stroma, OSMR was detected both in the epithelium and in the stroma, and LIFR
was only localized to the epithelium. In prostate cancer, OSM localized in all cell
types and OSMR was detected in the epithelium and in the stroma. There is also
a correlation between the increased OSM/OSMR expression and the Gleason
grade. LIFR was only detected in the epithelium of low Gleason grades (Royuela
et al., 2004). The cellular effects of OSM in vitro are prostate cancer cell type-
dependent. In DU145 cells (OSMR- and LIFR-positive), OSM/gp130 signaling
promoted cell growth with the activation of Stat3 (Mori et al., 1999). In 22Rv1
cells (LIFR positive and OSMR negative), OSM triggered cell proliferation in an
OSMR-independent manner through the p38 MAPK and PI3K/Akt pathways
without activating Stat3 (Godoy-Tundidor et al., 2005). In PC-3 (OSMR positive)
cells, OSM/gp130 signaling supported cell growth and rendered chemoresistance
to etoposide and cisplatin-mediated cytotoxicity (Borsellino et al., 1999).
Other cancers. OSM’s role in promoting cancer progression was also reported
in several other cancers. For examples, some myeloma cell lines produced
autocrine OSM, which might be induced by IL-10 and supported growth of
myeloma cells but not normal B lineage cells (Zhang et al., 1994) (Gu et al.,
1996a; Westendorf and Jelinek, 1996). In KS cells, OSM was indicated as a
potent mitogen with the ability to activate ERK1/2 (Amaral et al., 1993; Miles et
al., 1992; Nair et al., 1992).
135
Taken together, the response of cancer cells to OSM depends on the cellular
context (Grant and Begley, 1999). The expression and abundance of LIFR and
OSMR, the ratio between the amount of LIFR and OSMR on the cell surface, the
downstream signaling network structure, and the activation kinetics determine
OSM’s effects on the response and behavior of a cell. Further, the different
activities of OSM render the role of gp130 complicated in cancer progression.
CNTF/gp130 signaling
CNTF was originally identified for its ability to promote survival of neurons of the
ciliary ganglion, and exhibits survival and differentiative actions on cells of the
nervous system. It was identified to bind to gp130 in 1992, and the tyrosine
phosphorylations and gene activations induced by CNTF are very similar to
signaling events that characterize LIF and IL-6 responses in hematopoietic cells
(Ip et al., 1992). Based on our current knowledge, CNTF signaling has a limited
involvement in human cancers compared to the previously introduced gp130-
mediated signaling. CNTF was reported to serve as a growth factor for human
myeloma XG-4 and XG-6 cell lines (Zhang et al., 1994). Addition of soluble
CNTF receptor increased the potency of CNTF, whereas anti-gp130 antibodies
abrogated its growth-promoting effect. A CNTF-sensitive cell line XG4-CNTF
responded to CNTF in correlation with expression of cell surface CNTFR (Gu et
al., 1996b). The coexpression of CNTF and its receptor components (CNTFR,
LIFR and gp130) was detected in approximately half of human glioma samples
tested (Weis et al., 1999). It was recently reported that CNTFR expression is up-
regulated in undifferentiated tumor-initiating cells and gliomas of increasing tumor
136
grade, suggesting that CNTFR may serve as a tumor-initiating cell marker;
inhibition of CNTFR using anti-CNTFR antibody resulted in increased antibody-
dependent cell-mediated cytotoxicity in CNTFR expressing medulloblastoma
DAOY cells (Lu et al., 2012). Although effects of CNTF-related signaling on
cancers remain to be elusive, accumulated evidence suggests CNTF may
stimulate cell growth and proliferation in select cancers in a similar manner with
IL-6, IL-11 and LIF.
CLC/gp130 signaling
CLC was discovered as a second ligand for the CNTFR/LIFR/gp130 tripartite
receptor based on the phenotypic differences between CNTF- and CNTFR-
deficient mice (Elson et al., 2000). So far, however, little is known about its
relationship with cancer progression. Limited evidence showed that CLC
functions as a growth and survival factor for myeloma (Burger et al., 2003).
IL-27/gp130 signaling
As one of the latest discovered gp130 cytokines, only a few data are available for
the role of IL-27 signaling in human cancers. It was reported that IL-27R
mediated transformation of hematopoietic cells accompanied by the activation of
the JAK/STAT pathway (Pradhan et al., 2007). In melanoma, IL-27 exhibited an
antiproliferative activity through WSX-1/Stat1 pathway (Nagai et al., 2010;
Yoshimoto et al., 2008).
137
CT-1/gp130 and NP/gp130 signaling
There are currently no publications on the role of CT-1 and NP signaling in
human cancers.
138
IV-‐6:
GP130
IN
OTHER
DISEASES
AND
ITS
THERAPEUTIC
IMPLICATIONS

Since gp130 is involved in diverse physiological processes, dysregulation of
gp130 signaling is involved in many human diseases in addition to cancer.
Comprehensively reviewed by Jazayeri et al., major gp130-associated diseases
include rheumatoid arthritis (IL-6), castleman’s disease (IL-6), cardiac myxoma
(IL-6, IL-11), crohn’s diseases (IL-6), psoriasis (OSM, IL-11), asthma (IL-6, IL-
11), amyotrophic lateral sclerosis (CNTF), diabetes (IL-6), atherosclerosis (IL-6,
IL-11), systemic lupus (IL-6), obesity (CNTF), and postmenopausal osteoporosis
(IL-6). Therefore, targeting gp130 signaling is also promising for other human
diseases besides cancer.
For the treatment of gp130-dependent diseases, antagonizing gp130 signaling
pathways can be achieved by neutralizing gp130 cytokines, inhibiting gp130-
ligand binding, and suppressing downstream signaling cascades. Multiple
humanized monoclonal antibodies against IL-6 or IL-6R are currently under
clinical trials for various diseases (Table 7, www.clinicaltrials.gov). Although both
anti-IL-6 and anti-IL-6R antibodies successfully block IL-6 signaling, anti-IL-6R
antibodies have advantages in that anti-IL-6 antibodies can cause a massive
systemic increase in IL-6 because antibody-associated IL-6 cannot be cleared
from the circulation (Lu et al., 1992). An anti-IL-6R mAb, tocilizumab (drug name:
ACTEMRA, Genentech), was approved by the FDA in 2010 to treat adults with
moderately to severely active rheumatoid arthritis (RA). Considering the overlap
of signaling pathways stimulated by different gp130-related cytokines and cancer
cells’ ability to bypass a single inhibited pathway via utilizing alternative signaling
139
pathways, it is promising to develop inhibitors targeting gp130, the hub of these
signaling networks. B-R3, a pan-blocking anti-gp130 antibody, is widely used in
preclinical research to inhibit gp130 signaling (Chevalier et al., 1996), whereas
two other anti-gp130 mAbs B-S12 and B-P8 were developed as agonists for
gp130 signaling (Gu et al., 2000). Recently, we have reported the first-in-class
small-molecule gp130 inhibitor SC144 (Figure 25A) that shows gp130 inhibitory
activity and cytotoxicity in vitro and in vivo ovarian cancer models, with oral
bioavailability but no significant off-target toxicity to normal cells or tissues. The
development of small-molecule gp130 inhibitors will not only facilitate gp130-
based research, but also advance gp130-based therapies. Compared to
antibodies, small-molecule compounds possess the advantages in oral
bioavalability, ease of manufacturing, cost, stability and patient preference.
In addition, natural antagonists of gp130 receptor complexes are indeed
present in sera. For example, sgp130 is a natural competitive inhibitor selective
for the IL-6 trans-signaling. Sant7 was reported to exhibit a high affinity binding to
IL-6R subunit and behave as a receptor antagonist (Savino et al., 1997). A
soluble form of OSMR (sOSMR) was also detected with the ability to neutralize
OSM (Diveu et al., 2006). Recently, it has been shown that the IL-27 subunit p28
alone antagonizes cytokine signaling through gp130 independently of EBI3
(Stumhofer et al., 2010). On the other hand, numerous inhibitors targeting
downstream gp130 signaling molecules have been developed. Many of these
inhibitors are small-molecule compounds with anti-cancer activities, such as Jak2
140
inhibitor AG490, Stat3 inhibitor STA-21, PI3-kinase inhibitor, Akt inhibitor MK-
2006, Raf inhibitor SB-590885, and Mek inhibitor AZD6244.
Table 7. Antagonists against cytokine ligands or subunits of gp130 receptor
complexes in clinical trials
Drug Description Sponsor Phase Diseases
CNTO 328 /
Siltuximab
anti-IL-6 mAb Centocor, Inc II castleman’s disease
II myelodysplastic
syndrome
I/II multiple cancers
CNTO 136 anti-IL6 mAb Centocor, Inc II lupus nephritis
Olokizumab anti-IL-6 mAb UCB, Inc II rheumatoid arthritis
ALX-0061 anti-IL6R
nanobody
Ablynx I/II rheumatoid arthritis
Tocilizumab /
INN /
atlizumab
anti-IL-6R mAb Hoffmann-La
Roche (Genentech)
III systemic juvenile
idiopathic arthritis
(sJIA)
III rheumatoid arthritis
II systemic sclerosis
III ankylosing spondylitis
Hospital for Special
Surgery, New York
II polymyalgia rheumatic
(PMR)
National Cancer
Institute (NCI)
II castleman’s disease
Hospital Clinico
Universitario de
Santiago
III graves’
ophthalmopathy
Medical College of
Wisconsin
I/II acute graft-versus-host
disease
University Hospital
Inselspital, Berne
II giant cell arteritis
MAR 003 US anti-IL-6R mAb NIH clinical center I systemic lupus
erythematosus

141
IV-‐7:
CONCLUDING
REMARKS

gp130 is a shared part of receptor complexes for at least nine cytokines, the
signaling pathways of which regulate highly diverse biological processes.
Abnormal expression and/or activation of gp130 have been reported to be
associated with cancer progression as well as the development of a variety of
other human diseases. Based on the accumulating evidence, gp130-implicated
signaling supports cancer cell growth, proliferation, survival, migration, and/or
drug resistance in most cancer types. However, the detailed biology of gp130
signaling is cancer type-dependent mainly because of the differential involvement
of its cytokines as well as the various cellular contexts in different cancers. Due
to the chemotherapeutic resistance, which is one of the most significant issues in
cancer treatment, anticancer agents with new targets and positive therapeutic
impacts are always in medical demand. To develop novel anticancer agents
targeting gp130-mediated signaling, humanized monoclonal antibodies against
gp130 cytokines (particularly IL-6), gp130 and other gp130-related receptor
subunits as well as antagonists of downstream gp130 signaling molecules are
under intensive research. However, there is still a dearth of potent small-
molecule inhibitors of gp130. Due to the complicated role of gp130 in various
human cancers, gp130’s effects should be comprehensively understood on a
specific cancer type when antagonists of the gp130 signaling or agonists of the
OSM/gp130 signaling are applied for clinical use.

142
CHAPTER V: DISCOVERY OF AN ORALLY ACTIVE
INHIBITOR OF GP130

gp130 is part of the receptor-signaling complexes for at least eight cytokines (IL-
6, IL-11, IL-27, LIF, CNTF, OSM, CT-1 and CLC) (Jones et al., 2011). Ligand
binding induces the association of gp130 with a cytokine-specific receptor-α
chain, followed by the activation of downstream signaling cascades, including
JAK/STAT, RAS/RAF/MAPK, and PI3K/AKT pathways (Wang et al., 2009c). It
has been shown that phosphorylation of gp130 at Ser782 down-regulates cell
surface expression of gp130 (Gibson et al., 2005; Gibson et al., 2000). As a
ubiquitously expressed receptor, gp130 is involved in a wide range of important
biological processes including inflammation, autoimmunity, cancer (Silver and
Hunter, 2010), stem cell maintenance, and embryonic development (Rose-John,
2002). In recent years, increasing attention has been paid to the relationship
between gp130 and cancer progression. However, the role of gp130 in ovarian
cancer progression remains to be elusive, and there is a dearth of small-
molecule inhibitors selectively targeting gp130.
Previously, my lab discovered novel quinoxalinhydrazides to have desirable
physicochemical and drug-like properties and a broad-spectrum anticancer
activity (Grande et al., 2007; Oshima et al., 2009; Plasencia et al., 2009). Here, I
show that the lead compound, SC144 is a first-in-class small-molecule gp130
inhibitor with oral activity in ovarian cancer.
143
V-‐1:
RESULTS

V-1-1: SC144 Exhibits Anticancer Activity in a Panel of Human
Ovarian Cancer Cell Lines
SC144 exhibited cell growth inhibition in a panel of human ovarian cancer cell
lines with IC
50
values in a submicromolar range (Table 8, Figure 25A). SC144’s
potency towards NCI/ADR-RES (paclitaxel- and doxorubicin-resistant (Alvarez et
al., 1995; Lee et al., 1994; Roschke et al., 2003), IC
50
= 0.43 µM) and HEY
(cisplatin-resistant, IC
50
= 0.88 µM) indicates an ability to overcome drug
resistance in ovarian cancer. SC144 also inhibited colony formation in Caov-3
and OVCAR-8 cells (Figure 25B). We compared SC144’s abilities to induce
apoptosis and cell death in human ovarian cancer cells (OVCAR-8 and Caov-3)
with human normal epithelial cells (kidney epithelial cells and endometrial
epithelial cells). The kidney epithelial cells and the endometrial epithelial cells do
not express cancer specific marker survivin (Figure 25C) (Altieri, 2003). SC144
induced substantial apoptosis in OVCAR-8 and Caov-3 cells, but no considerable
apoptotic effect was induced in the kidney epithelial cells or the endometrial
epithelial cells (Figure 25D). Consistently, trypan blue staining showed that
SC144 significantly caused cell death in OVCAR-8 and Caov-3 cells, whereas no
significant cell death was induced by SC144 in the kidney epithelial cells or the
endometrial epithelial cells (Figure 25E). These results indicate that SC144
exhibits cytotoxicity in human ovarian cancer cells but not in human normal
epithelial cells.
144
Table 8. Cytotoxicity of SC144 in a panel of human ovarian cancer cell lines

IC 50 (µM)
a

OVCAR-8 OVCAR-5 OVCAR-3 Caov-3 SKOV-3 HEY NCI/ADR-RES
SC144 0.72 ± 0.32
b
0.49 ± 0.44 0.95 ± 0.35 0.44 ± 0.14 0.53 ± 0.04 0.88 ± 0.46 0.43 ± 0.12
a
IC 50 is defined as the drug concentration causing a 50% decrease in the cell population.
b
Values (mean ± SD) are calculated from three independent experiments.
145

Figure 25. SC144 exhibits cytotoxicity in human ovarian cancer cell lines. (A) chemical
structure of SC144. (B) SC144 inhibited colony formation of OVCAR-8 and Caov-3 cells. After
indicated SC144 treatments for 48 h, cells were cultured in fresh medium and allowed to form
colonies (10 days for OVCAR-8 and 15 days for Caov-3), followed by crystal violet staining. (C)
Survivin protein levels in a panel of human cancer cells and normal epithelial cells detected by
immunoblotting. (D) SC144’s apoptotic effect on OVCAR-8, Caov-3, normal kidney epithelial and
normal endometrial cells. After treatment with SC144 (2 µM), paclitaxel (2 µM) or equal amount of
DMSO for 72 h, cells were stained with Annexin V-FITC and propidium iodide (PI), and then
analyzed by flow cytometry. Upper panel: cells in the bottom left quadrant of each panel (Annexin
V-negative, PI-negative) are viable, whereas cells in the top left quadrant (Annexin V-positive, PI-
negative) are in the early apoptotic stage, and cells in the top right quandrant (Annexin V-positive,
PI-positive) are in the late apoptotic/necrotic stage. Lower panel: the percentage of apoptotic cells
is shown in a histogram. (E) Analysis of SC144 induced cell death in OVCAR-8, Caov-3, normal
kidney epithelial and normal endometrial cells. After treatment with SC144 (2 µM), paclitaxel (2
µM) or equal amount of DMSO for 72 h, cells were stained with trypan blue. Histogram shows the
percentage of trypan blue-stained cells after indicated treatment (***, P < 0.001. Bar, SEM).
146
V-1-2: SC144 Induces gp130 (Ser782) Phosphorylation and Down-
Regulates gp130 Glycosylation
SC144 treatment substantially increased the phosphorylation of gp130 (S782) in
both OVCAR-8 and Caov-3 cells in a time- (Figure 26A) and dose-dependent
manner (Figure 26B). In both cell lines, total gp130 protein was detected as two
individual bands in immunoblotting (Figure 26A). Silencing of gp130 using
siRNAs decreased the abundance of both bands (Figure 26C), confirming that
both bands are gp130. Treatment of OVCAR-8 and Caov-3 cells with brefeldin A
(BFA), an inhibitor of protein glycosylation produced a time-dependent shift of
gp130 into the lower band (Figure 26D), indicating that the upper band
represents glycosylated gp130 and the lower band represents unglycosylated or
minimal core glycosylated gp130. After 6 h continuous SC144 treatment in
OVCAR-8 and Caov-3 cells, the upper bands of total gp130 shifted to the lower
band (Figure 26A), indicating that SC144 treatment induced gp130
deglycosylation following the stimulation of gp130 phosphorylation. Also, SC144
is mechanistically different from BFA, which did not induce gp130
phosphorylation (Figure 26E). In addition, we found that gp130 ligands IL-6 and
LIF substantially enhanced Stat3 (Y705) phosphorylation, whereas the
phosphorylation of gp130 (Ser782) was only increased by LIF treatment (Figure
26F). This result suggests that phosphorylation at Ser782 is cytokine specific and
is not required for gp130 activation.
147

Figure 26. SC144 induces gp130 phosphorylation and deglycosylation. Immunoblot
detection of phosphorylation of gp130 (Ser782) in OVCAR-8 and Caov-3 cells treated by
SC144 in a (A) time- and (B) dose- dependent manner as indicated. C, Immunoblot
detection of gp130 after gp130 knockdown using gp130 siRNAs, as described in
Materials and Methods. D, Inhibition of gp130 glycosylation by brefeldin A (BFA).
OVCAR-8 and Caov-3 cells were treated with BFA (1 µg/mL) for the indicated periods of
time, followed by immunoblotting with a specific anti-gp130 antibody. E, BFA had no
significant effects on gp130 (Ser782) phosphorylation. OVCAR-8 cells were treated with
SC144 or BFA as indicated. Phospho-gp130 (Ser782) and total gp130 were determined
by immunoblotting. F, LIF but not IL-6 stimulated phosphorylation of gp130 (Ser782).
OVCAR-8 cells were treated with 50 ng/mL of IL-6 or LIF for 15 min or left untreated.
The levels of phospho-gp130 (Ser782) and phospho-Stat3 (Tyr705) were determined by
immunoblotting.
148
V-1-3: SC144 Decreases gp130 Protein Levels on Cell Surface
Previous studies have shown that phosphorylation of Ser782 leads to down-
regulation of gp130 cell-surface expression (Gibson et al., 2005; Gibson et al.,
2000; Radtke et al., 2010). Therefore, we assessed the cell-surface levels of
gp130 before and after SC144 treatment. After 24 h SC144 treatment, no
significantly change in cell surface-localized gp130 was detected with 2 µM
SC144, whereas SC144 significantly down-regulated surface-localized gp130 at
10 µM (Figure 27A), which is much higher than the concentration that induced
gp130 phosphorylation and deglycosylation. In comparison, IL-6 induced
immediate gp130 internalization as early as 10 min after stimulation (Figure 27B).
These results indicate that the downregulation of cell surface-localized gp130 by
SC144 does not directly caused by the induction of gp130 phosphorylation or
deglycosylation, and SC144 down-regulates cell surface-localized gp130 by a
different mechanism than cytokine-induced gp130 internalization.
149

Figure 27. SC144 decreases gp130 cell surface expression. (A) Cell surface-bound
gp130 was down-regulated in OVCAR-8 by SC144. After treatment, cells were collected
and stained with anti-gp130 (B-R3) antibody, and analyzed by FACS. The upper panel
shows data representing three independent experiments. The lower panels are statistical
analyses of data from the three independent experiments. Error bars = SEM. *** P <
0.001, paired t test. (B) IL-6 stimulated gp130 internalization. OVCAR-8 cells were
treated with IL-6 (50 ng/mL) for 15 min or left untreated. After treatment, cells were
collected and stained with anti-gp130 (B-R3) antibody, followed by FACS analysis.
150
V-1-4: SC144 Directly Binds gp130
Induction of gp130 phosphorylation and deglycosylation suggests that SC144
targets gp130. Binding of a small-molecule compound to its target protein can
result in conformational changes and proteolytic stabilization of the protein by
decreasing its sensitivity to proteases (Lomenick et al., 2009; Park and
Marqusee, 2005). Similar in concept to DNA footprint assay, Drug Affinity
Responsive Target Stability (DARTS) assay (Lomenick et al., 2009; Lomenick et
al., 2011) was used to test the binding of SC144 to gp130. Incubation of SC144
with OVCAR-8 cell lysate protected gp130 from digestion in a dose-dependent
manner as evident by the increased abundance of the gp130 band (Figure 28).
Particularly, 100 µM and 1000 µM of SC144 significantly increased gp130
abundance by 63% (P = 0.035) and 102% (P = 0.020) compared to vehicle
control. These results demonstrate the direct binding of SC144 to gp130, and
suggest SC144 may induce conformational changes in gp130 and affect its
activity.
151

Figure 28. SC144 directly binds gp130. DARTS assay was used to assess the binding
of SC144 to gp130. OVCAR-8 cell lysate was incubated with indicated concentrations of
SC144 for 1 h at room temp, followed by pronase digestion for 30 min at room temp and
analysis by Western blotting for protection of gp130. Upper panel, one of three
representative experiments is shown. Lower pane, statistical analyses of data from the
three independent experiments. Error bars = SEM. *, P < 0.05, paired t test.
152
V-1-5: SC144 Suppresses the Activation of Stat3
Stat3 is a primary downstream signaling molecule of gp130, and it transduces
gp130 signaling from cell surface to nucleus, where it activates target gene
expression (Debnath et al., 2012). Treatment of OVCAR-8 and Caov-3 cells with
an anti-gp130 (B-R3) antibody (Figure 29A) or silencing of gp130 using siRNAs
in OVCAR-8 cells (Figure 29B) substantially decreased constitutive Stat3 (Y705)
phosphorylation, indicating that the constitutive Stat3 activation in ovarian cancer
cells is mainly maintained by extracellular gp130 ligands constitutively produced
by ovarian cancer cells via an autocrine mechanism (Watson et al., 1990).
Therefore Stat3 activation can be used as a predictive marker for gp130 activity.
SC144 treatment suppressed the constitutive phosphorylation of Stat3 (Y705) in
a time- (Figure 29C and D) and dose-dependent manner (Figure 29E) in
OVCAR-8 and Caov-3 cells. The total Stat3 level was decreased after 24 h of
SC144 treatment (Figure 29D), perhaps due to an autoregulatory role of Stat3
(Ichiba et al., 1998). In comparison, stattic, a previously reported Stat3 inhibitor
(Schust et al., 2006), inhibited Stat3 (Y705) phosphorylation in OVCAR-8 cells in
a time-dependent manner (Figure 29F), whereas stattic exerted no substantial
effect on phosphorylated and total gp130, indicating that SC144-induced gp130
phosphorylation and deglycosylation do not result from the direct inhibition of
Stat3 (Y705) phosphorylation.
153

Figure 29. SC144 inhibits constitutive Stat3 phosphorylation in ovarian cancer
cells. (A) Inhibition of Stat3 phosphorylation by anti-gp130 antibody. Protein extracts
from OVCAR-8 and Caov-3 cells treated with a monoclonal antibody against an
extracellular region (B-R3) of gp130 at 10 µg/mL for the indicated periods of time were
analyzed for phospho-Stat3 (Ser782) and total Stat3. (B) Stat3 phosphorylation was
decreased after gp130 knockdown using siRNA. SC144 inhibited Stat3 (Y705)
phosphorylation in OVCAR-8 and Caov-3 cells in (C and D) a time-dependent manner at
2 µM and (E) a dose-dependent manner after 1 h. (F) Effects of stattic on phospho-
gp130 (Ser782), total gp130, phospho-Stat3 (Tyr705) and total Stat3 were examined in
OVCAR-8 cells after treatment with Stattic (5 µM) for the indicated periods of time.
154
V-1-6: SC144 Decreases the Expression of gp130/Stat3 Target Genes
The activation of gp130/Stat3 promotes the expression of multiple genes,
including Bcl-2 (Selvendiran et al., 2008), Bcl-X
L
(Duan et al., 2006), Cyclin D1
(Cai et al., 2010), survivin (Cai et al., 2010), Ape1/Ref-1 (Haga et al., 2003), and
MMP-7 (Fukuda et al., 2011), which are involved in cell growth, survival, cell
cycle progression, proliferation and metastasis. The role of gp130/Stat3 in
promoting the expression of these genes in OVCAR-8 cells was confirmed by
silencing of gp130 (Figure 30A) or Stat3 (Figure 30B) using respective siRNAs.
Importantly, Stat3 knockdown also reduced gp130 expression, which is
consistent with a previous study unveiling a STAT-binding element within the
gp130 promoter region (O'Brien and Manolagas, 1997). During SC144 treatment
in OVCAR-8 cells, survivin, MMP-7 and gp130 were the most sensitive to SC144
treatment; their abundance was significantly decreased during the first 24 h
(Figure 30C), whereas Cyclin D1, Bcl-X
L
, Bcl-2 and Ape1/Ref-1 expression
started to decrease after 24 h. Stat1 level was not affected by SC144. These
results also provide additional evidence that SC144 is mechanistically different
from BFA, which increased gp130 level (Figure 26D). These results further
confirm that SC144 inhibits the activation of the Stat3 signaling pathway
downstream of gp130.

155

Figure 30. SC144 suppresses the expression of gp130/Stat3 target genes. Silencing
of (A) gp130 or (B) Stat3 down-regulated the protein levels of gp130, Stat3, Bcl-2, Bcl-
X
L
, survivin, MMP-7 and Ape1/Ref-1, measured by immunoblotting. (C) gp130/Stat3-
mediated gene expression was inhibited in OVCAR-8 cells treated with SC144 (2 µM) for
the indicated periods of time.
156
V-1-7: SC144 Potently Blocks Cytokine-Triggered gp130 Signaling
We compared the effects of SC144 on the activation of downstream signaling
stimulated by gp130 cytokines, including IL-6 and LIF, and non-gp130 cytokines,
including an interferon family member IFN-γ, a GPCR (CXCR4) ligand SDF-1α,
and a growth factor PDGF. IL-6 stimulates Stat3 phosphorylation in a dose-
dependent manner (Figure 31A). When 3 ng/mL IL-6, which is close to the IL-6
concentration in the human body (Watson et al., 1990), was used to induce Stat3
phosphorylation, SC144 exhibited the inhibitory effect at 2 µM and showed
complete inhibition at 5 µM. When 50 ng/mL IL-6 was used to stimulate the
downstream pathways of gp130, SC144 exhibited the inhibitory effect at over 10
µM. These results indicate that SC144 competitively inhibits the downstream
signaling activation elicited by IL-6. In addition, SC144 inhibited LIF-induced
phosphorylation of Stat3 (Y705) and Stat1 (Y701) in a dose- and time-dependent
manner (Figure 31B). SC144 also inhibited LIF-induced Akt activation. In
contrast, SC144 did not affect the activation of phosphorylation of Stat3, Stat1
and Akt stimulated by IFN-γ (Figure 31C). Akt phosphorylation induced by SDF-
1α (Figure 31D) or PDGF (Figure 31E) was not affected by SC144. Neither SDF-
1α nor PDGF exhibited significant effects on Stat3 phosphorylation in OVCAR-8.
These results demonstrate that the inhibitory effect of SC144 on the downstream
signaling is gp130-specific and gp130-dependent.
157

Figure 31. SC144 inhibits downstream signaling induced by gp130 cytokines.
SC144 inhibited the downstream signaling elicited by (A) LIF or (B) IL-6 in a dose-
dependent manner without affecting the downstream signaling induced by (C) IFN-γ, (D)
SDF-1α or (E) PDGF. OVCAR-8 cells were serum-starved overnight prior to 4 h
pretreatment (or 1 h pretreatment for LIF) with SC144 at the indicated concentrations,
followed by stimulation with IL-6 (3 ng/mL or 50 ng/mL as indicated) for 10 min, LIF (50
ng/mL) for 15 min, IFN- (50 ng/mL) for 20 min, SDF-1α (80 ng/mL) for 10 min, or
PDGF (20 ng/mL) for 15 min. Whole-cell lysates were subjected to immunoblotting using
specific antibodies for the indicated proteins.
158
The selectivity of SC144 in inhibiting the activation of downstream signaling
stimulated by gp130 cytokines was also confirmed by examining the nuclear
translocation of Stat3 (Figure 32) and Stat1 (Figure 33) induced by LIF, IL-6 or
IFN-γ, which occurs following tyrosine phosphorylation. Immunofluorescence
microscopy showed that SC144 treatment effectively inhibited LIF- and IL-6-
induced nuclear translocation of Stat3 and Stat1 but was not able to affect
STATs nuclear translocation induced by IFN-γ in OVCAR-8 cells.
159

Figure 32. SC144 inhibits nuclear translocation of Stat3 induced by gp130
cytokines. OVCAR-8 cells were serum-starved overnight prior to 4 h pretreatment with
or without SC144 (20 µM). Cells were treated with IL-6 (50 ng/mL) for 10 min, LIF (50
ng/mL) for 15 min, or IFN-γ (50 ng/mL) for 20 min, or left untreated. Cells were then
fixed, permeablized, and stained with anti-Stat3 or anti-Stat1 antibody (Red), and
SYTOX
®
GREEN nuclear staining (Green). Images were captured using Leica TCS SP
confocal system with 40x or 100x magnification and are representative of three
independent experiments. Scale bar: 50 µm.
160

Figure 33. SC144 inhibits nuclear translocation of Stat1 induced by gp130
cytokines. OVCAR-8 cells were serum-starved overnight prior to 4 h pretreatment with
or without SC144 (20 µM). Cells were treated with IL-6 (50 ng/mL) for 20 min, LIF (50
ng/mL) for 15 min, or IFN-γ (50 ng/mL) for 20 min, or left untreated. Cells were then
fixed, permeablized, and stained with anti-Stat3 or anti-Stat1 antibody (Red), and
SYTOX
®
GREEN nuclear staining (Green). Images were captured using Leica TCS SP
confocal system with 40x or 100x magnification and are representative of three
independent experiments. Scale bar: 50 µm.
161
V-1-8: Inhibition of gp130/Stat3 Leads to Cytotoxicity
We further assessed whether the inhibition of gp130/Stat3 signaling contributes
to the cytotoxicity of SC144 in ovarian cancer cells. By using gp130 siRNAs or
Stat3 siRNAs, we found that silencing of gp130 (Figure 34A) or Stat3 (Figure
34B) significantly inhibited OVCAR-8 cell growth. In addition, anti-gp130 (B-R3)
antibody reduced OVCAR-8 cells’ ability to form colonies in a dose-dependent
manner (Figure 34C). Pharmacological suppression of Stat3 activity by stattic
inhibited cell growth in OVCAR-8, Caov-3 and NCI/ADR-RES cells (Table 9).
Taken together, these results demonstrate that the gp130/Stat3 inhibition results
in cytotoxicity in human ovarian cancer cells.

162

Figure 34. gp130/Stat3 inhibition causes cytotoxicity in human ovarian cancer
cells. (A) gp130 or (B) Stat3 knockdown inhibited cell growth in ovarian cancer cells.
OVCAR-8 cells were transfected with gp130 siGP130s, siSTAT3s or siCtrl in 96-well
plates. After 96 h, cells were stained with MTT for 3 h at 37°C and then dissolved in
DMSO. Absorbance was measured in a 96-well plate reader at 570 nm. **, P < 0.01, ***,
P < 0.001. B, Colony formation in OVCAR-8 cells was inhibited by 48 h treatment with
anti-gp130 (B-R3) antibody at indicated concentrations.

Table 9. Cytotoxicity of stattic in a panel of human ovarian cancer cell lines

IC
50
(µM)
a

OVCAR-8 Caov-3 NCI/ADR-RES
stattic 6.1 ± 0.1
b
13.7 ± 1.3 3.4 ± 0.2
a
IC
50
is defined as the drug concentration causing a 50% decrease in the cell population.
b
Values (mean ± SD) are calculated from three independent experiments.

163
V-1-9: SC144 and Its Active Analogs Inhibit gp130/Stat3 Signaling and
Show Cytotoxicity in Other Cancer Types
We further tested SC144 and its analogs SC186, SC204 and SC205 (Figure
35A) in a panel of cancer cells. SC144 exhibited similar cytotoxicity in human
ovarian, colon, pancreatic, and prostate cancer cells as well as mouse PTEN−/−
prostate cancer cells, evaluated by both MTT assay (Table 10) and colony
formation assay (Figure 35B). Particularly, prostate cancer cell lines LNCaP and
CE1 are androgen-sensitive, whereas DU145 and E2 are androgen-refractory.
The cytotoxicities of SC186 and SC204 were ~ 10-fold less than that of SC144
(Table 10, Figure 35B). Compared with the structure of SC144, SC186 and
SC204 lack the fluorine, which may lead to efficient metabolism and decrease in
gp130 binding affinity of the metabolites, resulting in lower inhibitory activities
and cytotoxicity. In addition, SC205 exhibited a complete loss of activity, probably
because of the hydroxybenzoate that makes it an immediate substrate for
metabolism, and the lack of a pyrazine nitrogen, a potential hydrogen bond
acceptor. Consistently, SC144 inhibited gp130/Stat3 signaling in all the cell lines
tested (Figure 36). SC186 and SC204 showed similar activities at higher
concentrations, whereas SC205 had no effects at concentrations up to 10 µM.
These results also suggest the importance of gp130/Stat3 signaling and the
potential use of SC144 in these cancer types.
164
Table 10. Cytotoxicity of SC144 and its analogs in a panel of cancer cell lines
IC50
a
(µM)
SC144 SC186 SC204 SC205
Human
ovarian
cancer
NCI/ADR-RES 0.16 ± 0.05
b
2.7 ± 0.3 4.6 ± 0.4 > 10
OVCAR-8 0.36 ± 0.04 2.5 ± 0.3 2.0 ± 0.5 > 10
colon
cancer
HCT116 p53+/+ 0.14 ± 0.01 1.6 ± 0.1 1.1 ± 0.1 > 10
pancreatic
cancer
Mia-Paca2 0.20 ± 0.02 2.4 ± 0.3 2.2 ± 0.1 > 10
Panc-1 0.19 ± 0.01 4.3 ± 0.4 2.2 ± 0.1 > 10
prostate
cancer
LNCaP 0.12 ± 0.02 2.7 ± 0.3 2.0 ± 0.1 > 10
DU145 0.26 ± 0.05 3.3 ± 0.9 3.2 ± 0.1 > 10
Mouse
PTEN−/−
prostate
cancer
CE1 0.54 ± 0.03 3.4 ± 0.3 3.0 ± 0.1 > 10
E8 0.60 ± 0.01 2.2 ± 0.1 2.0 ± 0.1 > 10
a
IC
50
is defined as the drug concentration causing a 50% decrease in the cell population.
b
Values (mean ± SEM) are calculated from three independent experiments.

165

Figure 35. Effects of SC144 and its analogs on the abilities of a panel of cancer
cells to form colonies. (A) Chemical structures of SC186, SC204 and SC205. (B)
SC144 and its active analogs SC186 and SC204 impairs colon formation in indicated
cell lines. Cells were treated with indicated compounds for 24 h, and then allowed to
form colonies in fresh medium without drugs. Colonies were stained with crystal violet.
166

Figure 36. Effects of SC144 and its analogs on the activity of the gp130/Stat3
signaling. Cells were treated with SC144, SC186, SC204 and SC205 at indicated
concentrations. Levels of gp130, phospho-Stat3 (Y705), Stat3, and surviving were
analyzed using immunoblotting.

167
V-1-10: SC144 Suppresses Tumor Growth in Human Ovarian Cancer
Xenografts
To determine in vivo efficacy of SC144, we tested its effect on established
tumors after subcutaneous inoculation of human ovarian cancer OVCAR-8 cells
in the flank of nude mice. Compared with vehicle control treatment, daily i.p.
administration of SC144 (10 mg/kg) for 58 days significantly inhibited tumor
growth by ~73% (from 835.2 ± 228.1 mm
3
to 228.1 ± 92.5 mm
3
; P = 0.032)
(Figure 37A). To assess SC144’s oral bioavailability, we tested SC144 through
daily p.o. administration. After we sacrificed the mice in the control group on day
69 due to their large tumors, we continued SC144 treatment for another 35 days.
The average tumor volume in mice receiving daily p.o. administration of SC144
(100 mg/kg) was ~82% smaller than that in the control group (on Day 69, 975.8 ±
247.9 mm
3
vs. 175.4 ± 80.8 mm
3
, P = 0.014) (Figure 37B). No significant body
weight loss was detected during both treatment periods (Figure 37C and D),
indicating that SC144 did not produce significant adverse effects in mice at its
effective anticancer dosage.
168

Figure 37. SC144 suppresses human ovarian cancer xenograft in nude mice. (A)
Growth curves of s.c. tumors in nude mice treated with vehicle or SC144. OVCAR-8
tumor-bearing mice were treated daily with SC144 at 10 mg/kg (n = 4) or vehicle (n = 4)
through i.p. injection for 5 days, followed by 2 days of rest, over a treatment period of 58
days. (B) Growth curves of s.c. tumors in the SC144 (p.o.) treatment group (n = 4) or
control group (n = 5). OVCAR-8 tumor-bearing mice were treated daily with or without
SC144 (100 mg/kg) through p.o. administration for 5 days, followed by 2 days of rest.
Mice in the control group (blue line) were sacrificed on Day 69. Mice in the treatment
group (red line) were sacrificed after 104 days of SC144 treatment. Results are
presented as mean ± SEM (*, P < 0.05). (C) Comparison of body weight of control mice
(blue line) and SC144 (i.p.)-treated mice (red line). (D) Comparison of body weight of
control mice (blue line) and SC144 (p.o.)-treated mice (red line).
169
While the tumors excised from control mice appeared red, the ones from
SC144-treated mice were pale (Figure 38A), suggesting that SC144 treatment
effectively inhibited tumor angiogenesis. SC144-induced reduction in tumor
microvessel density was further confirmed by the significant decrease in both
CD31 (Figure 38B) and von Willebrand factor (vWF) staining (Figure 38C). While
SC144 treatment resulted in extensive areas of necrosis in the tumor (Figure 39A
and B), no substantial toxicity was observed in liver, kidney, spleen, lung, heart,
pancreas and brain (Figure 39C and D), further demonstrating the safety of
SC144.
170

Figure 38. SC144 treatment reduced tumor blood vessels in OVCAR-8 xenograft
mice. (A) tumors excised from i.p. vehicle-treated and SC144-treated mice are shown.
Microvascular density was confirmed by IHC staining of (B) CD31 and (C) von
Willebrand factor (vWF). Percentage of positively stained areas was determined by
areas of stained microvessels divided by number of nuclei. Bar, SEM. *, P < 0.05; **, P <
0.01.
171

Figure 39. SC144 treatment induces extensive areas of necrosis in tumor but no
damage in normal tissue. Comparisons of control tumor sections with SC144 (A) i.p.
treated or (B) p.o. treated tumor sections. Proportion of necrosis was statistically
analyzed with five different scopes. ***, P < 0.0001. Tissue sections from indicated
organs from SC144 (C) i.p. treated or (D) p.o. treated mice are shown.
172
Consistent with our in vitro data, SC144 treatment triggered apoptosis and
suppressed cell proliferation in tumor, as is confirmed by the significant increase
in TUNEL and cleaved caspase 3 staining and significant decrease in Ki67
staining, respectively (Figure 40A). The levels of gp130, MMP-7, Bcl-X
L
, Bcl-2
and Ape1/Ref-1 in the tumor were substantially decreased in the SC144
treatment group (Figure 40B), which is consistent with our in vitro data (Figure
40C). Phospho-gp130 (S782) was decreased after SC144 treatment, which may
result from the downregulation of total gp130 protein. The decrease in Bcl-X
L
,
Bcl-2 and Ape1/Ref-1 explains the extensive necrosis in the tumor tissue from
the SC144 treatment mice due to the roles of these gp130 effectors in protecting
cancer cells from apoptosis (Fishel and Kelley, 2007; Patel et al., 2009), whereas
the reduced level of MMP-7 contributes to both apoptosis and anti-angiogenesis
(Ii et al., 2006; Sang, 1998).
173

Figure 40. Effects of SC144 on in vivo ovarian tumor. (A), IHC staining of TUNEL,
cleaved caspase 3 and Ki67. Percentage was determined by the number of double-
stained cells divided by the number of nuclei. Bar, SEM. **, P < 0.01; ***, P < 0.001. (B)
SC144 (i.p.) treatment down-regulated gp130/Stat3-mediated gene expression in
OVCAR-8 tumors from xenograft mice. Tumor homogenates were analyzed using
immunoblotting with indicated antibodies.

174
V-‐2:
DISCUSSION

IL-6 and Stat3 are involved in cancer progression and drug resistance in a variety
of human cancers including ovarian cancer. Inhibition of IL-6 secretion by CDDO-
Me induced apoptosis and sensitized drug-resistant ovarian cancer cells to
paclitaxel and cisplatin (Duan et al., 2009). CNTO 328, a monoclonal antibody to
IL-6, is currently in a Phase I clinical trial for ovarian cancer. On the other hand,
Stat3 is persistently active in ovarian cancer cell lines but not in normal ovarian
epithelial cells (Huang et al., 2000). In vitro and in vivo studies confirmed that
inhibition of constitutive Stat3 activation by using a Jak-selective inhibitor, a
dominant negative Stat3, or shRNA suppressed the growth and development of
ovarian cancer (Burke et al., 2001; Catlett-Falcone et al., 1999; Huang et al.,
2008; Niu et al., 1999). Considering that gp130 is positioned at the junction of
this oncogenic signaling network and is decisive for the network activation,
blocking gp130 activity using nonpeptidic small molecules is an attractive
therapeutic approach to gp130-dependent cancers. However, so far no selective
small-molecule gp130 inhibitors have been developed. A small molecule HDAC
inhibitor AR-42 (formerly known as (S)-HDAC-42) decreases gp130 expression
and inhibits Stat3 phosphorylation (Zhang et al., 2010). Nevertheless, its effect
on down-regulating gp130 expression may result from the Stat3 inhibition. In our
study, we developed a quinoxalinhydrazide compound SC144, which inhibits
gp130-cytokine signaling, as well as the downstream gene expression, and
exhibits in vitro and in vivo potency in ovarian cancer. Due to the fact that the
gp130 cytokine IL-6 is constitutively produced and the key effectors (Stat3, Akt)
175
in the gp130 downstream signaling pathways are persistently active in ovarian
cancer (Cheng et al., 2002; Liu et al., 2001; Nicosia et al., 2003; Watson et al.,
1990), ovarian cancer cells might be more sensitive to gp130 inhibitors compared
to normal cells. This hypothesis was confirmed by our results that SC144 exhibits
cytotoxicity in human ovarian cancer cells without inducing apoptosis or cell
death in human normal epithelial cells, and inhibits human ovarian tumor growth
in a mouse xenograft model without causing significant damage to normal
tissues. Unlike AR-42, SC144 dose not exhibit an inhibitory effect on HDAC
(Figure 41). To our knowledge, SC144 is the first-in-class small-molecule gp130
inhibitor with oral activity in ovarian cancer. In addition, our results showing that
gp130 persistently activates Stat3 in ovarian cancer cells, promoting cell growth
and proliferation, are consistent with a similar discovery in colitis-associated
tumorigenesis (Bollrath et al., 2009). Although the identification of residues on
gp130 that SC144 interacts and the elucidation of off-targets are beyond the
scope of this present study, the following evidence together demonstrates gp130
is directly inhibited by SC144: (a) SC144 binds to gp130 and induces gp130
(S782) phosphorylation, deglycosylation and internalization; (b) anti-gp130
antibody is able to completely suppress constitutive Stat3 activation, indicating
Stat3 activation as an indicator for SC144’s effect on gp130; (c) SC144 inhibits
Stat3 activation and the expression of downstream genes; (d) SC144 potently
inhibits the activation of the downstream signaling pathways stimulated by gp130
ligands but exhibits no effects on the activation of the downstream signaling
pathways stimulated by non-gp130 ligands.
176

Figure 41. SC144 has no significant effects on HDAC activity. Nuclear protein
extracts from HCT116 cells were prepared and used for an HDAC activity assay. (A)
Deacetylated standard curve. (B) HDAC activity was tested in vitro as described in
Supplementary Materials and Methods. Nuclear protein extracts (2 mg/mL) in a 96-well
plate were incubated with SC144 at the indicated concentrations, 1 µM Trichostatin A
(TSA) was a positive control, and an equal amount of DMSO was a negative control, as
were untreated extracts, followed by the examination of HDAC activity using a
fluorescence plate reader (EX: 355 nm; EM: 460 nm).
177
The role of phospho-S782 in gp130 was first documented in 2000 to down-
regulate gp130 cell surface expression (Gibson et al., 2000). Recently, it has
been reported that phospho-S782-associated downregulation of gp130 cell
surface expression is mediated by immediate internalization (Radtke et al.,
2010). Indeed, phosphorylation of many transmembrane receptors at Ser/Thr
residues is commonly associated with the negative modulation of receptor
activity (Sibley et al., 1987). The induction of gp130 glycosylation and the
decrease in total gp130 protein level may also contribute to Stat3 inhibition at a
later time. Moreover, inhibition of gp130/Stat3 amplifies SC144’s cytotoxicity
through down-regulating the expression of a large number of downstream genes,
such as Bcl-2, Bcl-X
L
, Cyclin D1, MMP-7, Ape1/Ref-1 and survivin in ovarian
cancer cells, which results in the submicromolar IC
50
values after 72 h SC144
treatment (Table 8 and 10).
On the basis of our and previous studies, we established a plausible model for
the anticancer mechanism of SC144 in ovarian cancer (Figure 42). SC144
binding induces conformational changes and activity loss in the cell surface-
bound gp130 receptor, exposing the Ser782 residue to protein kinases, such as
CaMKII, CaMKIV, and MK2 (Gibson et al., 2005; Radtke et al., 2010). The
decrease in gp130 activity leads to the inactivation of downstream signaling
pathways involving Stat3 and Akt. The suppression of gp130/Stat3-mediated
gene expression amplifies SC144’s potency due to the inhibition of cell cycle
progression and angiogenesis, and the promotion of apoptosis and cell death. In
addition, the downregulation of the PI3K/Akt pathway also contributes to
178
apoptosis and cell death. Our working model may serve to identify a unifying
anticancer mechanism likely to be shared by future gp130 inhibitors.

Figure 42. A working model for the anticancer mechanism of SC144 in ovarian
cancer. We surmise that SC144 is cytotoxic to ovarian cancer cells via a mechanism
involving the inhibition of gp130 activity, which leads to the inactivation of Akt and Stat3
as well as the suppression of Stat3-regulated gene expression. As a result, SC144
treatment eventually causes cell cycle arrest, anti-angiogenesis and apoptosis.

179
V-‐3:
MATERIALS
AND
METHODS

Reagents
SC144, a quinoxalinhydrazide derivative, was synthesized as described (Grande
et al., 2007). Stattic, a Stat3 inhibitor (Schust et al., 2006), was purchased from
Sigma-Aldrich. Stock solutions of 10 mM SC144 and Stattic were prepared in
DMSO and stored at -20°C. Further dilutions were made fresh in cell-culture
medium. IL-6, IFN-γ and PDGF were purchased from PeproTech. LIF was
purchased from Santa Cruz Biotechnology. SDF-1α plasmid was a generous gift
from Dr. Ghalib A. Alkhatib (Indiana University, Indianapolis, Indiana) and the
protein was expressed and purified as described (Altenburg et al., 2007). Anti-
phospho-Stat3 (Y705, 9145), anti-Stat3 (9132), anti-gp130 (3732), anti-phospho-
Stat1 (Y701, 9171), anti-Stat1 (9172), anti-cyclin D1 (2978), anti-Bcl-X
L
(2764),
anti-Bcl2 (2870), anti-phospho-Akt (T308, 2965), anti-phospho-Akt (S473, 4060),
anti-Akt (9272) and anti-Ape1 (4128) were purchased from Cell Signaling
Technology. Anti-phospho-gp130 (Ser782, 12978-R), anti-Actin (1616-R), normal
rabbit IgG (3888), and goat anti-rabbit IgG-HRP (2030) were purchased from
Santa Cruz Biotechnology. Anti-gp130 (B-R3, 852-060-000) was purchased from
Cell Sciences. Mouse IgG2a (010-0141) and goat anti-mouse IgG-HRP (610-
1302) were purchased from Rockland. Goat anti-mouse Cy5-linked IgG and goat
anti-rabbit Cy5-linked were purchased from GE Life Sciences. gp130 siRNA,
Stat3 siRNA, control siRNA, siRNA transfection reagent, and siRNA transfection
medium were purchased from Santa Cruz Biotechnology. Trypan Blue 0.4%
solution was purchased from Lonza Inc.
180
Cell Culture
OVCAR-8, OVCAR-5 and OVCAR-3 cells (National Cancer Institute,
Developmental Therapeutics Program) were maintained in RPMI-1640
supplemented with 10% heat-inactivated FBS (Gemini-Bioproducts). NCI/ADR-
RES cells (National Cancer Institute) were maintained in RPMI-1640
supplemented with 10% heat-inacivated FBS and 5 mM L-glutamine. HEY and
CAOV-3 cell lines were kindly provided as gifts by Dr. Louis Dubeau (University
of Southern California, Keck School of Medicine, Los Angeles, CA). HEY cells
were maintained in DMEM supplemented with 10% heat-inacivated FBS and 5
mM L-glutamine. CAOV-3 were maintained in MEM supplemented with 10%
heat-inactivated FBS. Human normal kidney epithelial and human normal
endometrial epithelial cells were kindly provided by Dr. Alan Epstein (University
of Southern California, Keck School of Medicine, Los Angeles, CA) and
maintained in RPMI-1640 supplemented with 10% heat-inactivated FBS. Mouse
PTEN-/- prostate cancer cell lines CE1 and E8 were kindly provided by Dr. Padip
Roy-Burman (University of Southern California, Keck School of Medicine, Los
Angeles, CA). Cells were grown as monolayers at 37°C in a humidified
atmosphere of 5% CO
2
. To remove adherent cells from the flask for subculture
and counting, cells were washed with PBS without calcium or magnesium,
incubated with a small volume of 0.25% trypsin-EDTA solution (Mediatech, Inc.)
for 5-10 min, resuspended with culture medium and centrifuged. All experiments
were performed using cells in the exponential growth phase. Cells were routinely
checked for mycoplasma contamination by using PlasmoTest (InvivoGen).
181
Colony Formation Assay
Colony formation assays were performed as described (Munshi et al., 2005).
Briefly, OVCAR-8 cells (600 cells/well) and Caov-3 cells (800 cells/well) were
seeded in 24-well plates and allowed to attach. After 48 h, serial dilutions of the
corresponding compounds or antibodies were added to the culture medium and
incubated for 48 h. Cells were cultured until colonies were formed (10 days for
OVCAR-8 and 15 days for Caov-3), then subsequently washed, stained with
crystal violet solution (2%) for 1 h, and thoroughly washed with water.
Western Blotting
Cells (only attached cells were collected) or xenograft tumor tissues were
washed with ice-cold PBS, and lysed in cold lysis buffer (20 mM Tris-HCl, 150
mM NaCl, 1 mM EDTA, 1% Triton X-100, pH 7.5) with 1x protease and
phosphatase inhibitors. Protein concentration was determined by BCA protein
assay (Thermo Scientific). Proteins were resolved on 8% or 10% SDS-PAGE
and electrotransferred to Immun-Blot PVDF membrane (Bio-Rad). After blocking
with 5% milk in TBST, membranes were probed with the indicated primary
antibodies, subsequently with horseradish peroxidase-conjugated secondary
antibody, and developed using Dura Extended Duration Substrate (Thermo
Scientific). Immunoreactive proteins were visualized with the Chemi-Doc System
(Bio-Rad).
182
Growth Inhibition Assay
Growth inhibition was assessed using a 3-(4,5-dimethylthiazol-2-yl)-2,5-
diphenyltetrazolium bromide (MTT) assay as described earlier (Yamada et al.,
2010). Cells were seeded in 96-well microtiter plates, allowed to attach 24 h prior
to siRNA transfections or the addition of corresponding compounds to the culture
medium. After 72 h, cells were incubated with 0.3 mg/ml MTT (Amresco) for an
additional 3 h at 37°C. After removal of the supernatant, dimethyl sulfoxide
(DMSO) was added to the wells and the absorbance was read at 570 nm. All
assays were performed in triplicate. Percentage of cell growth inhibition was
expressed as: (1-A/C) x 100% (A and C were the absorbance values from
experimental and control cells, respectively). Inhibitory concentration 50% (IC
50
)
values were determined for each drug from a plot of log (drug concentration)
versus percentage of cell growth inhibition. Standard deviations were calculated
based on the IC
50
values obtained from at least three independent experiments.
Flow Cytometry Analysis of Cell Surface gp130 Expression
OVCAR-8 cells were grown in full media and subjected to SC144. Only attached
cells were collected by detaching with a non-enzymatic cell dissociation solution
(Sigma-Aldrich) and centrifugation, and washed with cold PBS. After blocking
with 10% goat serum for 15 min, cells were stained with anti-gp130 (B-R3)
antibody (Cell Sciences, 852.060.000) or isotype IgG on ice for 1 h, subsequently
exposed to Cy5-linked goat anti-rabbit secondary antibody on ice for 1 h, and
analyzed using a LSR II flow cytometer (Becton Dickinson).
183
DARTS Assay
The Drug Affinity Responsive Target Stability (DARTS) assay was optimized and
used to assess the binding of SC144 on gp130 via the protease protection from
pronase (Roche Applied Science, Inc) as previously described (Lomenick et al.,
2009; Lomenick et al., 2011). OVCAR-8 cells were lysed using M-PER (Thermo
Scientific, Inc) supplemented with protease and phosphatase inhibitors. The
supernatant of cell lysate containing 4 – 6 µg/µL total proteins was incubated with
SC144 at indicated concentrations at room temperature for 1 h, followed by
proteolysis with 1 µg pronase to every 9,600 µg of lysate for 30 min at room
temperature. Final concentration of DMSO was 1% in all samples. To stop
proteolysis, 5× SDS sample loading buffer (Tris-HCl 0.25 M, pH 6.8, SDS 10%,
glycerol 50%, bromophenol blue 0.5%, DTT 100 mM) was added to each sample
at a 1:4 ratio, mixed well, and boiled at 100°C for 5 min. Samples were analyzed
by Western blotting.
Annexin V-FITC Apoptosis Assay
OVCAR-8 and Caov-3 cells (2 x 10
5
) were seeded in 35 mm dishes, allowed to
attach overnight, and then received indicated treatments. Cells were washed with
cold PBS. Both floating and attached cells were collected, stained with the
Annexin V-FITC apoptosis detection kit (BioVision) according to the
manufacturer’s protocol. The resulting fluorescence was measured by a LSR II
flow cytometer (Becton Dickinson).
184
Fluorescence Confocal Microscopy
OVCAR-8 cells were grown on poly-L-lysine (Sigma-Aldrich) coated glass
coverslips and subjected to serum-free RPM1-1640 overnight. With or without
compound pretreatments, cells were stimulated with 50 ng/mL IL-6 for 20 min (for
Stat1) or 10 min (for Stat3), 50 ng/mL LIF for 15 min, or 50 ng/mL IFN-γ for 20
min. Cells were then fixed with 3.7% formaldehyde in PBS for 15 min at room
temperature and permeabilized for 10 min with ice-cold 100% methanol at -20°C.
After blocking with 5% goat serum, slides were incubated with anti-Stat1 (1:200)
or anti-Stat3 (1:1000) antibodies overnight at 4°C, followed by simultaneous
incubation with Cy5-linked goat anti-mouse or goat anti-rabbit secondary
antibody (1:1000) and 100 nM SYTOX
®
Green nucleic acid stain (Invitrogen) at
room temperature for 1 h. Images were collected on a Leica TCS SP confocal
system with 40x or 100x magnification.
siRNA Transfections
siRNAs for gp130 (siGP130.1: Santa Cruz, #sc-29333; siGP130.2, siGP130.3,
siGP130.4: OriGene, #SR302381) and Stat3 (siSTAT3.1: Santa Cruz, #sc-
29493; siSTAT3.2: Santa Cruz, #sc44275) were used in this study. Subconfluent
OVCAR-8 cells were transfected in 6-well or 96-well plates with 80 nM gp130,
Stat3, or control siRNA according to the manufacturer’s protocol. Protein
expression was tested by immunoblotting after 72 h of siRNA exposure. Cell
viability was evaluated by MTT staining after 48, 72 and 96 h of siRNA exposure.
185
Mice
Athymic mice (Charles River Laboratories, Wilmington, MA) were used for in vivo
efficacy studies. Mice were fed ad libitum and kept in air-conditioned rooms at 20
± 2°C with a 12 h light-dark period. Animal care and manipulation were in
agreement with the USC institutional guidelines, which were in accordance with
the Guidelines for the Care and Use of Laboratory Animals.
In Vivo Tumor Xenograft Studies
OVCAR-8 cells in the logarithmic growth phase from in vitro cultures were
implanted in athymic mice (5 x 10
5
cells in 100 µL of PBS/mouse) under aseptic
conditions as previously described (Xu et al., 2012). Tumor growth was assessed
by biweekly measurement of tumor diameters with a Vernier caliper. Tumor
volume was calculated according to the formula: Tumor volume (mm
3
) = D ×
d
2
/2, where D and d are the longest and shortest diameters, respectively. For i.p.
administration, tumors were allowed to grow to an average volume of 70 mm
3
.
Mice were then randomly assigned into 2 groups, 4 mice per group, for daily
vehicle control or SC144 (10 mg/kg) treatment. To prepare SC144 solution for
i.p. administration, 200 mg/ml DMSO stock solution of SC144 was diluted to 20
mg/mL in propylene glycol, and further added to 0.9% NaCl with 40% propylene
glycol. For p.o. administration, tumors were allowed to grow to an average
volume of 40 mm
3
. Mice were then randomly assigned into control (5 mice) and
treatment (4 mice) groups. SC144 (100 mg/kg) was orally delivered every day to
the treatment group. To prepare SC144 solution for p.o. administration, 300
186
mg/mL DMSO stock solution of SC144 was mixed with sesame oil. Mouse body
weight and tumor volume were measured twice a week.
Histochemical Analysis
Upon autopsy, tumors, kidneys, livers, pancreas, spleens, hearts, lungs and
brains from mice were collected, fixed in formalin, embedded in paraffin, and
sectioned. Sections (4 µm) were stained with H&E to facilitate histologic
examination.
Immunohistochemistry
Immunohistochemistry was performed on formalin-fixed, paraffin-embedded
sections, following antigen retrieval. Internal hydrogen peroxidase activity was
blocked with 3% hydrogen peroxide for 15 min. Tissue sections were
subsequently blocked by Power block for 10 min and 10% goat serum for 30 min,
followed by incubation with indicated antibodies at 4 ºC overnight. Anti-von
Willebrand factor antibody (Dako, A0082) was kindly provided by Dr Florence M
Hofman (University of Southern California, Keck School of Medicine). Anti-Ki67
mAb (Thermo Scientific, RM-9106-S) was kindly provided by Dr Bangyan Stiles
(University of Southern California, School of Pharmacy). Anti-CD31 antibody
(Santa Cruz biotechnology, inc, sc-8306) was kindly provided by Dr Jason Wu.
Anti-cleaved caspase 3 antibody was purchased from Cell Signaling, Inc (9661).
Tissue sections were then incubated with biotin-conjugated secondary antibody
(BioGenex, HK337-5G) for 20 min, and peroxidase-conjugated streptavidin
(BioGenex, HK330-5K) for 20 min. TSA Cyanine 3 plus evaluation kit
187
(PerkinElmer, Inc, NEL744E001KT) was used to generate fluorescent signals,
while nucleus was stained by DAPI. TUNEL staining was performed according to
manufacturer’s instruction (Roche, 11 684 817 910).
Statistical Analysis
The Student’s test was used for statistical analysis and p-value determination via
SPSS 16.0 (SPSS Inc.) and Prism 5 (GraphPad Software, Inc.). Differences were
considered statistically significant at P < 0.05.

188
CHAPTER VI: FUTURE WORK
During my Ph.D. study, I stepped into the following fields in the context of cancer:
PDI, PACMA, gp130, Stat3, survivin, and SC144. Scientists have dedicated huge
amount of effort to exploring the stories behind them, but a lot of questions have
not yet got satisfying answers. Please allow me to use the saying from Dr. Walter
Biship (TV series “Fringe”), “so much happened here, and so much is about to”.
Below, I list some of my thoughts for future work in these areas.
VI-‐1:
Is
Cancer
a
Curable
Disease?

Although we already have quite a few chemotherapeutic drugs as powerful
weapons against cancer, they only delay our loss in the battle when we face late-
stage cancers. In these cancer cases, chemotherapeutic drugs do no more than
slightly extending patients’ lives. Optimistically, I think some cancer subtypes are
eventually curable by chemotherapy, but not all. Therefore, it is necessary and
important to choose the right cancer subtype to begin the research with. Cancers
are like out-of-control cars, and there are a vast variety of causes, such as flat
tires, and malfunctions in the break or the wheel. However, broken windows or
damaged bumpers should not be counted, although they are significantly
associated with all out-of-control cars. Same for cancer. Identifying the critical
cause for a cancer subtype is necessary (but not sufficient) for the development
of effective, specific and safe chemotherapeutic approaches. We cannot deny
some out-of-control cars cause unfixable damages. In addition, it is important for
scientists who are dedicated to developing targeted approaches for cancer
189
therapy to comprehensively understand the mechanisms of their cancer subtypes
and the roles of the targets.
VI-‐2:
Destroy,
Control
or
Reprogram

All anticancer agents fall into two catalogs. One is to destroy cancer cells, such
as induction of apoptosis and activation of immune response. The other is to
control cancer cell progression, such as inhibition of metastasis. However,
according to the knowledge accumulated from stem cell research, there may be
an alternative option: cancer reprogramming. Cancer cells continuously grow,
proliferate, and migrate via regaining the stem-like properties and abilities, or in
other words, via de-diferentiation. It may be possible to develop approaches to
re-differentiate cancer cells, turning them back to normal status. Such
approaches should have less toxicity to normal cells and tissues. To achieve this
goal, it is necessary to understand the roles of development-related molecules in
cancer. However, although the possibility of reprograming cancer cells to normal
cells could be limited by genetic mutations above certain levels ( the “point of no
return” for cancer cells), transformed cells before reaching the “point of no return”
should be reprogrammable.
VI-‐3:
Dark
Matters
in
Cancer

Traditional cancer research focuses on the genetic and protein levels. Many
genes, promoters, enhancers and DNA modifications (epigenetics) have been
reported to contribute to cancer progression. Recently, microRNAs have got
scientists’ attention due to their regulatory functions on protein abundance and
190
even cell signaling pathways. However, there may be intracellular or extracellular
"dark matters" associated with cancer. These dark matters may be beyond our
current knowledge of biomaterials, and their detection may be limited by our
current technology. They could even be energy fields, such as electric and
magnetic fields, because a large amount of biological molecules are charged and
movements of charged molecules generate magnetic fields. As life is
complicated, oftentimes we need to think and look out of the box.
VI-‐4:
Effects
of
PACMAs
on
PDI

In my results, I showed that PACMA 31 inhibited PDI’s enzymatic activity in the
insulin turbidity assay. This assay only measures the reductase activity of PDI.
However, PDI is a multi-function protein with at lease oxidase, reductase,
isomerase and chaperone activities. PACMA 31’s effects on other PDI activities
remain unclear. Because PACMA 31 binds to PDI’s active-site cysteines, which
contribute to the reductase, oxidase and isomerase activities of PDI, I predict that
PACMA 31 also suppresses PDI’s oxidase and isomerase activities. However,
the chaperone activity of PDI primarily depends on the substrate binding domain
b’, which does not have active-site cysteines. Therefore, PACMAs may not affect
PDI’s chaperone activity.
VI-‐5:
Other
Targets
of
PACMAs

PACMAs covalently bind to target proteins via their terminal propynoic moieties.
PDI was first selected from our fluorescent 2D gel because it has the strongest
fluorescence intensity amount all fluorescent spots. However, there may be other
191
target proteins contributing to the toxicity of PACMAs. Because many other PDI
family proteins share the same or similar active sites with PDI, it is possible that
active PACMAs may bind and inhibit some of them. It would also be interesting to
compare the binding affinity of active PACMAs to PDI and other PDI family
proteins. These results would be useful to expand our knowledge about PACMAs’
cytotoxic mechanism.
VI-‐6:
Toxicity
Associated
with
PDI
Inhibition

Accumulating evidence from silencing of PDI using siRNA or shRNA has told us
that complete depletion of PDI is cytotoxic to normal cells. This is not surprising
because PDI is such an important protein in regulating a board range of cellular
events. Therefore, when we use PDI inhibitors for the treatment of cancer as well
as other diseases (HIV-1 infection, Huntington disease, et al.), it is critical to
figure out their therapeutic windows.
VI-‐7:
Other
Potential
Targets
of
SC144

I have shown that SC144 targets gp130/Stat3 signaling pathway. However,
SC144 may have other targets responsible for its cytotoxic mechanisms. Based
on my observation, SC144 has no significant toxicity after 24 h treatment in
cultured cancer cells at concentrations up to 10 µM. After 48 h treatment at 0.5 –
10 µM, there are vesicles accumulating inside the cells, which can be clearly
observed under microscope. These vesicles share similar morphology with
autophagosomes. In fact, I also detected using Western blotting an increase in
the LC3 protein levels after 24 – 48 h SC144 (2 µM) treatment (data not shown).
192
These observations suggest that SC144 may induce autophagy or inhibit the
fusion of autophagosome and lysosome. After 72 h treatment, even at 2 µM
SC144 shows killing effects in over 90% of total cells. Therefore, SC144 does not
exert an immediate cytotoxic effect. Instead, it regulates multiple signaling
pathways and affects the abundance of a board range of cellular proteins. In
addition, although surivin is an important target gene of Stat3, SC144 can also
suppress survivin expression in cells with inactive Stat3, such as MCF-7, SKBR-
3 and PC3 (data not shown). These observations indicate there are Stat3-
independent mechanisms by which SC144 decreases survivin expression. In the
DARTS assay, SC144 shows protection of multiple proteins against pronase,
suggesting SC144 binds to several cellular proteins, and some of these binding
events may be associated with the decrease of protein activity. If I had another 5
years to further explore the molecular mechanisms of SC144, I would primarily
focus on its effects on the reduction of survivin and the induction of
autophagosome-like vesicles. They could be two independent events, or they are
subsequently triggered by the same pathway. Moreover, although SC144
induces apoptosis, but the percentage of apoptosis is far lower than the
percentage of SC144-induced cell death measured by the MTT assay or the
colony formation assay. Therefore, SC144 mainly causes apoptosis-independent
cell death. After SC144 treatment, trypan blue only stains a small portion of the
cells, excluding necrosis as an important cause. There are also interesting
observations from SC144-treated mouse xenografts. First, SC144 treatment in
mouse xenografts substantially decreased the number of inflammatory cells in
193
almost all organs, including tumors, suggesting that SC144 may also have a role
in regulating the activity of immune system. Secondly, SC144 substantially
reduces tumor angiogenesis. Although MMP-7 is associated with angiogenesis,
the decrease in MMP-7 may not be the only cause, or even a main cause. The in
vivo anti-angiogenic mechanism of SC144 remains unclear. These questions
about SC144 need future work to provide answers.

194
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Abstract (if available)

Abstract Ovarian cancer is one of the leading cancers causing women’s death in the United States mainly due to its late diagnosis and resistance to current clinically used drugs. Therefore, the development of novel potent drugs for the treatment of ovarian cancer is in urgent medical need. ❧ Protein disulfide isomerase (PDI), an endoplasmic reticulum (ER) chaperone protein, catalyzes disulfide bond breakage, formation and rearrangement. The effect of PDI inhibition on ovarian cancer progression is not yet clear, and there is a dearth of potent, selective and safe small-molecule inhibitors of PDI. In chapter III, I report a novel class of propynoic acid carbamoyl methyl amides (PACMAs) that are active against a panel of human ovarian cancer cell lines. Using fluorescent derivatives, 2D gel electrophoresis, and mass spectrometry, we established that PACMA 31, one of the most active analogs, act as irreversible small-molecule inhibitors of PDI. We also demonstrated that PDI is essential for the survival and proliferation of human ovarian cancer cells. In vivo PACMA 31 showed tumor targeting ability and significantly suppressed ovarian tumor growth without causing damage in normal tissues. These irreversible small-molecule PDI inhibitors represent a new approach for the development of targeted anticancer agents for ovarian cancer therapy, and can also serve as useful probes for investigating the biology of PDI-implicated pathways. ❧ On the other hand, despite abundant evidence that ovarian cancer progression is dependent upon IL-6/Stat3 signaling, the role of glycoprotein 130 (gp130), the signal transducer of this signaling axis, is unclear in ovarian cancer, and there is a dearth of small-molecule inhibitors of gp130. In chapter V, I report that gp130 is an attractive drug target in ovarian cancer due to its role in promoting ovarian cancer progression via the activation of its downstream Stat3 signaling, and identify a small-molecule gp130 inhibitor SC144 with anticancer activity. In vitro, SC144 exhibits potency in human ovarian cancer cells without cytotoxicity in normal epithelial cells. SC144 binds gp130, induces gp130 phosphorylation (S782) and deglycosylation, abrogates Stat3 phosphorylation and nuclear translocation, and thus inhibits the expression of downstream target genes. In addition, SC144 shows potent inhibition of gp130 ligand-triggered signaling. In vivo, SC144 suppresses tumor growth with oral bioavailability in a mouse xenograft model of human ovarian cancer without causing damage to normal tissues. ❧ My discovery of PDI and gp130 as potential drug targets for ovarian cancer treatment, and the identification of PACMA 31 and SC144 as inhibitors of PDI and gp130, respectively, not only promote the development of targeted therapy, but also bring new hope for cancer patients.

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Creator Xu, Shili (author)

Core Title Discovery of novel small molecules for ovarian cancer treatment

School School of Pharmacy

Degree Doctor of Philosophy

Degree Program Pharmaceutical Sciences

Publication Date 04/23/2013

Defense Date 03/12/2013

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

Tag chemotherapy,drug resistance,ER stress,glycoprotein 130 (gp130),inhibitor,OAI-PMH Harvest,oral bioavailability,ovarian cancer,PACMA 31,propynoic acid carbamoyl methyl amide (PACMA),protein disulfide isomerase (PDI),quinoxalinhydrazide,SC144,small molecule,Stat3,unfolded protein response (UPR),xenograft

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Advisor Neamati, Nouri (committee chair), Duncan, Roger (committee member), Stiles, Bangyan L. (committee member), Zandi, Ebrahim (committee member)

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