Combination of Th1 Cytokines Plus Small Molecule Kinase

Home , Palbociclib

COMBINATION OF TH1 CYTOKINES PLUS SMALL MOLECULE KINASE

INHIBITORS PALBOCICLIB OR SUNITINIB POTENTIATE APOPTOSIS IN BREAST

CANCER CELL LINES

A thesis submitted

To Kent State University

Fulfillment of the requirements for the

Degree of Master of Science

Nirmala Ghimirey

August, 2018

Except for previously published materials Thesis Written by

Nirmala Ghimirey

B.S., Kent State University, Kent, Ohio, 2016

MS., Kent State University, Kent, Ohio 2018

Approved by

Gary Koski, Ph.D. , Advisor

Ernest J. Freeman, Ph.D. , Chair, School of Biomedical Sciences

James L. Blank, Ph.D. , Dean, College of Arts and Sciences

TABLE OF CONTENTS

Table of contents………………………………………………………………………………...iii

List of Figures…………………………………………………………………………………...iv

List of Table……………………………………………………………………………………vii

Acknowledgements………………………………………………………….…………………viii

Chapters

1) Introduction……………………………………………………………………………1

2) Methods and materials……………………………………………………………….29

3) Results…………………………………………………………………………...…...34

4) Discussion……………………………………………………………………………62

5) Future Direction…………………………………………….. ……………………....71

6) References…………………………………………………………………………....74

iii

LIST OF FIGURES

Figure 1. Growth factor activated receptor tyrosine kinases pathway…………………………17

Figure 2. Schematic of intrinsic and extrinsic apoptosis pathways ……………………………18

Figure 3. Schematic of caspase 3 cleavage of PARP…………………………………………...19

Figure 4. Alamar Blue staining to determine cellular metabolic activity……………………….20

Figure 5. Trypan Blue staining to distinguish living from dead cells…………………………...21

Figure 6. TMRE staining to determine mitochondrial membrane potential…………………….22

Figure 7. Annexin V/ Propidium iodine (PI) staining examining apoptosis…………………….23

Figure 8. Ibrance chemical structure…………………………………………………………….24

Figure 9. Cell cycle……………………………………………………………………………...25

Figure 10. Sunitinib chemical structure…………………………………………………………26

Figure 11. Sunitinib action of mechanism………………………………………………………27

Figure 12. Combination of Th1 cytokines and ibrance enhance metabolic suppression of breast cancer cells.………………………………………………………………………………………38

Figure 13. Metabolic activity of breast cancer cell lines is maximally suppressed by dual Th1 cytokine and ibrance treatment.……………………………………………………………….....39

Figure 14. Microscopic examination of treated cells reveals evidence of cell death..…………..40

Figure 15. Histogram analysis of Trypan Blue staining reveals enhanced cell death with combined Th1 cytokine and ibrance treatment ………………………………………………….41

Figure 16. Combination of Th1 cytokines and ibrance significantly increase breast cancer cells death ……………………………………………………………………………………………..42

Figure 17. Ibrance plus Th1 cytokines causes maximized decrease mitochondria membrane potential, an indicator of apoptosis ……………………………………………..……………….43

Figure 18. Combination of Th1 cytokines plus ibrance significantly decrease mitochondria membrane potential over single treatments for breast cancer cells……………………………...44

Figure 19. Combination of ibrance and Th1 cytokines maximizes markers of apoptosis in breast cancer cell lines ………………………………………………………………………………….45

Figure 20. Combined Th1 cytokine and ibrance treatment minimizes levels of total Rb protein in

SKBR-3 cells…………………………………………………….………………………………46

Figure 21. Detection of phosphorylated Rb protein in SKBR-3………………………………...47

Figure 22. Combination of Th1 cytokines and sunitinib enhance metabolic suppression of breast cancer cells over single treatments………………………………………………….…………...53

Figure 23. Metabolic activity of breast cancer cell lines is maximally suppressed by dual Th1 cytokines and sunitinib treatment…………………………………………………………...... 54

Figure 24. Microscopic examination treated cells with sunitinib plus Th1 cytokines reveals evidence of cell death…………………………………………………………………………….55

Figure 25. Histograms analysis of Trypan Blue staining reveals enhanced cell death with combined Th1 cytokines and sunitinib treatment………………………..………………………56

Figure 26. Combination of Th1 cytokines and sunitinib significantly increased breast cancer cell death over single treatments ……………………………………………..………………………57

Figure 27. Combination of Th1 cytokines and sunitinib maximized markers of apoptosis in breast cancer cell lines……………………………….…….…………………………………….58

Figure 28. Dual treatment significantly suppresses HER-2 expression over single treatments...59

Figure 29. Cytochrome c release in SKBR3 cell lines demonstrate apoptosis….………………60

Figure 30. Dual Th1 cytokine treatment plus sunitinib results in a trend toward lowered PARP levels in SKBR-3 cells……………………………………………………….…………………..61

LIST OF TABLE

Table 1. Breast cancer classification…………………………………………………………….28

vii

ACKNOWLEDGEMENTS

I am very grateful for all the help I received from several individuals that led to the completion of this thesis. Without their input, it would not have been possible. Foremost, I am very thankful for endless support, guidelines, and motivation Dr. Koski provided throughout my research experience. Dr. Koski’s constant suggestions and improvements were very meaningful and gave hope during the writing process of this thesis, which I greatly appreciated.

I would like to thank my committee members, Dr. Colleen Novak and Dr. Gail Fraizer for their valuable time and effort to improve this thesis and for serving on my committee.

I am very thankful to Lori Showalter, for teaching me how to do several experiments and bearing with me along every step. I really appreciate her patience with me for answering my questions and providing support for completion of this thesis. Thank you, Crystal Oechsle, for suggestions and technical assistance. I would also like to thank my other lab members, Aaron

Title, Sierra Glass, and Chase Steel.

Special thanks to Adam Kulp for his feedback on drafts. I really appreciated the morale support I got from Adam, Devanshi Metha, and Tim Niepokney.

Lastly, I want to thank my grandparents/mentors (Terry and Liz Kuhn) and family for all the encouragement and support I received throughout my graduate school experience and during the writing process.

viii

Chapter 1

Introduction

General Background

Breast cancer is a major health problem with over 230,000 new cases of invasive breast cancer expected to be diagnosed each year (Desantis et al., 2016). Breast cancer is classified based on the expression of several biological markers, including presence or absence of hormone receptor (estrogen or progesterone) or overexpression of human epidermal growth factor receptor

2 (HER-2) protein (Anderson et al., 2014). These markers allow the disease to be placed in various phenotypic groups (Table 1).

In breast cancer, HER-2 overexpression has been associated with metastasis, resistance to chemotherapy, and poor prognosis (Harari and Yarden, 2000). In mouse models, tissue-specific forced overexpression of HER-2 is sufficient to induce high rates of spontaneous breast cancer development, indicating that HER-2 may be involved in the earliest stages of oncogenesis

(Siegel and Muller, 1996; Finkle et al., 2004). Because of these findings, HER-2 has become the target of a number of efforts to develop targeted drugs and immune-based therapies with the hope of crippling this important oncodriver, or marking cells that express it for immune- mediated attack (lapatinib (Burris, 2004), trastuzumab (Vogel et al., 2002), including anti-HER-2 vaccine efforts (Peoples et al., 2005; Sears et al., 2011)). For example, it has been shown that

HER-2-targeted, dendritic cell (DC)-based neoadjuvant vaccine therapy for early breast cancer

(ductal carcinoma in situ; DCIS) resulted in pathological complete response (pCR; i.e. no disease 1

at the time of surgery) in about a third of subjects that do not also co-express estrogen receptor

(ER). This was accompanied by strongly suppressed expression of HER-2 in about half of the patients with remaining disease. In contrast, only 5% of vaccine recipients with ERpos disease experienced pCR (Sharma et al., 2012). It is not immediately apparent how ER expression could make breast cancer cells resistant to vaccine therapy. However, it was considered that both HER-

2 and ER both act as growth factor receptors and initiate biochemical signaling events that drive proliferation of breast cancer cells (Osborne et al., 2005). This means that for breast cancer cells expressing HER-2 but not ER, an anti-HER-2 vaccine may interfere with the only growth factor receptor capable of supporting proliferation and continued viability, leading inevitably to death of the cancerous cells. On the other hand, a tumor cell that expresses both HER-2 and ER can afford to have HER-2 signaling disrupted, because in can fall back on ER for growth factor signaling. If this is the actual explanation, then it follows that disruption of ER signaling in conjunction with vaccination should normalize response rates in HER-2pos ERpos vaccine recipients. Fortunately for the testability of this hypothesis, a number of FDA-approved anti- estrogen drugs are available. These work by preventing estrogen binding to its receptor (e.g.

Tamoxifen (Mandlekar and Kong, 2001)) or by inhibiting aromatase, thus preventing estrogen biosynthesis (e.g. Anastrozole (Miller, 2003)). In a follow-up study to the original HER-2 vaccine trial, a short course of these anti-estrogen drugs were added concurrent with vaccine therapy. The results were dramatic: ERpos vaccine recipient pCR rates jumped from 5% to about

30%, a response rate not statistically differing from their ERneg counterparts (Lowenfeld, Mick, et al., 2017).

The anti-estrogen experience showed several things. First, it suggested that at least one reason a subset of patients were non-responsive to vaccine therapy was that their tumors

possessed additional growth factor signaling pathways that allowed them to survive HER-2 targeting. Second, it demonstrated that small-molecule drugs targeting these additional pathways could be used to enhance vaccine efficacy. Finally, it highlights the possibility that the now 70% of patients that still do not experience pCR from vaccine therapy, or vaccine therapy plus anti- estrogen do so because their tumors possess yet additional escape pathways that may be potentially exploited by other targeted drugs to further enhance efficacy of therapy.

It remains not entirely certain which effectors of vaccine-induced immunity are primarily responsible for anti-tumor activity. However, considerable indirect evidence points to soluble factors produced by CD4pos T cells of the “Th1” phenotype. The traditional effectors of antitumor immunity are CD8pos “cytotoxic” T cells (Mahmoud et al., 2011). However, immunohistochemical staining of excised DCIS samples before and after vaccination show relatively few such cells infiltrating areas of disease after vaccination (Czerniecki et al., 2007).

In contrast, the numbers of CD4pos T cells increase dramatically for many patients (Czerniecki et al., 2007; Sharma et al., 2012), and stimulation of extracorporealized T cells show enhancement of IFN-γ responses (characteristic of Th1 T cells) after vaccination (Czerniecki et al., 2007;

Koski et al., 2012). Finally, in vitro studies using a variety of human and murine breast cancer lines showed that exposure to a combination of two cytokines (IFN-γ and TNF-α) characteristic of Th1 cells result in high levels of cellular apoptosis as well as suppression of expression of surface HER-2. Therefore, Th1 cytokines can mimic in vitro many of the in vivo effects of vaccination (Namjoshi et al., 2016).

Interestingly, Th1 cytokines were used as an in vitro surrogate for vaccine-induced immunity to test the efficacy of anti-estrogen drugs prior to the clinical trial combining anti- estrogen therapy with vaccination for ERpos DCIS patients (Lowenfeld, Zaheer, et al., 2017). The

ERneg lines were highly sensitive to Th1 cytokines, while ERpos lines were relatively resistant.

However, the presence of anti-estrogen drugs rendered the ERpos lines sensitive to cytokines, while as expected, they had no additive effect on ERneg lines. This not only served as pre-clinical data to support the anti-estrogen trial, but it also demonstrated the general principle of screening targeted drugs in vitro for their additive or synergistic activity with Th1 cytokines against breast cancer lines as a way to predict which agents would likely work cooperatively with vaccination to improve overall clinical responses. It is the goal of this thesis to expand upon this idea and examine additional targeted drugs that can be quickly translated into clinical trials to improve pCR rates in response to vaccination. The drugs selected for these studies are palbociclib, an inhibitor of cyclin-dependent kinase 4/6, and sunitinib, a relatively broad-spectrum receptor tyrosine kinase antagonist. In order to understand why these drugs should be considered, a brief discussion of biochemical pathways regulating growth and proliferation in cancer cells is in order.

Cell signaling and proliferation

Receptor tyrosine kinases (RTKs) are a class of transmembrane proteins that play an important role in cell signaling pathways for cell proliferation, migration, differentiation, and survival. Some of the subfamilies of RTKs include vascular endothelial growth factor receptors

(VEGFRs), epidermal growth factor receptors (EGFRs), insulin-like growth factor receptors

(IGFRs), platelet-derived growth factor receptors (PDGFRα/β), stem cell factor receptors (Kit), and fetal liver tyrosine kinase receptor 3 (FLT3) (Paul and Mukhopadhyay, 2004). Usually, when a ligand (EGF, VEGF etc) binds to these RTKs in the extracellular domain, monomeric RTKs dimerize. This close association results in trans-phosphorylation of the receptor’s intracellular domains. An adaptor protein, Grb2, then binds to the phospho-tyrosine and also associates with

guanine-nucleotide exchange factor (GEF protein) (Weinberg, 2013). GEF exchanges GTP for

GDP and leads to activation of Ras (small monomeric G-binding protein). Ras activation further phosphorylates and activates multiple proteins downstream such as Raf, PI3K, and Ral-GEFs.

Activation of these proteins lead to protein synthesis, cell cycle progression, transcription factor activation, and cell migration (Figure 1) (Weinberg, 2013).

For example, activation of Ras mitogenic signaling leads to activation of cyclin D1 and E

(Weinberg, 2013). Cyclin-dependent kinases (CDK) play a key role in control of the cell cycle progressing from G1 (growth), S (replication), G2 (growth), and M (mitosis) phase (Weinberg,

2013). Regulation of progression from G1 to S phase is a critical restriction point for cell cycle progression. Interaction of CDK with cyclin protein is involved in regulating the restriction points. Cyclin D forms an association with cyclin dependent kinases 4 and 6 (CDK4/6) and the activated complex then phosphorylates retinoblastoma protein (Rb, a tumor suppressor) and other members of Rb protein family including p107 and p130 (Caldon et al., 2006; Giacinti and

Giordano, 2006; Peyressatre et al., 2015). In quiescent cells, Rb is hypophosphorylated and binds to E2F (transcription factor) and suppresses E2F function (Knudsen and Knudsen, 2008). When

Rb is hyperphosphorylated E2F is released, allowing transition from G1 to S-phase leading to cell cycle progression (Figure 1) (Nevins, 2001). Therefore, a defined and continuous chain of known biochemical events link RTK activation with cell cycle regulation in part through the regulated activity of cyclin-dependent protein kinases.

The other side of the coin: Biochemical events associated with apoptotic cell death p53 is a master regulator of cell life and death

The human body tightly regulates cell growth and apoptosis. Cells have to go through several checkpoints to move onto the next phase of mitosis for replication and proliferation as

described previously. However, many environmental and intrinsic factors can affect whether a cell continues to survive and proliferate or whether it dies via apoptosis. Cell apoptosis is characterized by several morphological changes such as cell shrinkage, chromatin condensation, formation of membrane-bound apoptotic bodies, and nuclear breakdown (Saraste and Pulkki,

2000). Ultra-violet radiation, reaction with oxidative free radicals, and genotoxic agents lead to

DNA damage (Gudkov and Komarova, 2003). When DNA is damaged, several proteins and cell signaling cascades activate to initiate repair of the damaged DNA or to trigger apoptosis. For example, if there is damage to double-stranded DNA, there is rapid induction of p53 protein. The p53 protein is a tumor suppressor that halts progression into the cell cycle (Vogelstein et al.,

2000). P53 activation leads to ATM kinases initiation, which in turn activate chk1 kinases

(Appella and Anderson, 2001). Activation of chk1 kinase phosphorylates p53 and protects it from Mdm2, a protein that ubiquinates unphosphorylated p53 leading to its degradation (Michael and Oren, 2003). Activated p53 can in turn activate DNA repair enzymes and return the cell to active proliferation. If the DNA damage is too severe to repair, it can lead to a block to angiogenesis and trigger initiation of apoptosis (Harris and Levine, 2005).

Two general pathways for induction of apoptosis: Intrinsic and Extrinsic

Intrinsic pathway is triggered when a signal for apoptosis is initiated from within the cells

(Christensen et al., 2013). Activated p53 can inhibit pro-survival signals (Bcl-2/Bcl-x) (Kluck,

Bossy-Wetzel, et al., 1997) and activate pro-apoptosis signals (Bax/Bak/Bid) (Zimmermann and

Green, 2001) through the intrinsic pathway. Pro-apoptotic signals lead to opening of mitochondrial voltage-dependent anion channels to allow release of cytochrome c into the cytoplasm (Kluck, Martin, et al., 1997). Cytochrome c is normally located in the inner membrane of mitochondria transporting electrons from complex 3 to complex 4 during routine

oxidative metabolism. Bax or Bak also causes loss of mitochondria membrane potential and cell swelling (Narita et al., 1998) due to the loss of cytochrome c. Release of cytochrome c prevents electrons from transferring between complexes (Christensen et al., 2013), hence mitochondria membrane potential is no longer maintained. The released cytochrome c binds to Apaf-1 protein and forms a seven-spoked wheel-like complex called the apoptosome (Zou et al., 1997).

Procaspase 9, an initiator procaspase, then binds at the middle of the wheel and gets cleaved by the apoptosome to generate its active form (Stennicke et al., 1999). Apoptosome formation leads to recruitment of executioner procaspase 3, 6, and 7, which are cleaved by caspase 9 and thus activated (Figure 2) (Zimmermann and Green, 2001). Activated caspase 3 further cleave PARP

(poly [ADP-ribose] polymerase), thus converting it from active to an inactive form leading to apoptosis (Figure 3) (Boulares et al., 1999; Soldani and Scovassi, 2002).

Cells can also go through apoptosis via extrinsic pathways. In extrinsic pathways, an extracellular signaling agent (such as cytokines or other factors) binds the extracellular domain of a transmembrane receptor, causing its activation. When initiating apoptosis such a receptor is refer to as a death receptor (Ashkenazi, 2002). The ligands of death receptors include the Th1- associated cytokine TNF-α as well as other tumor necrosis factor (TNF) members such as Fas ligand, and TRAIL (Zimmermann and Green, 2001). Ligand binding activates the receptor, and the death domain of the receptor binds and activates Fas-associated death domain proteins. This results in formation of a complex called death-inducing signaling complex (DISC) (Wajant,

2002). DISC triggers cleavage of inactive pro-enzyme caspase 8 and caspase 10 (initiator caspases) to the active forms. These in turn cleave and activate executioner caspases 3, 6, and 7

(Boatright and Salvesen, 2003). However, caspase 3 is considered to be primary an executioner caspase due to its importance in apoptosis-mediated DNA fragmentation, chromatin margination

and nuclear collapse (Slee et al., 2001). It is at this point (caspase 3 activation) where intrinsic and extrinsic pathways converge. However, in Fas-mediated apoptosis, Bid (pro-apoptosis) is activated by caspase 8, which leads to release of cytochrome c from the mitochondria (Figure 2)

(Zimmermann and Green, 2001). Therefore, cytochrome c release can occur through both extrinsic and intrinsic pathways.

Alteration of membrane structure in apoptotic cells

Activated caspase 3 has been linked to inhibition of flippase activity (Mandal et al.,

2002). Flippase is an ATP-dependent enzyme that transport aminophospholipids from the extracellular leaflet of the plasma membrane to the intracellular leaflet causing specific cytosol- directed transbilayer movement (Leventis and Grinstein, 2010). In an asymmetrical lipid bilayer, phosphatidylcholine and sphingomyelin are mainly located in the outer leaflet, whereas phosphatidylserine and phosphatidylethanolamine are restricted to intracellular leaflet (Leventis and Grinstein, 2010). In an apoptotic cell, phosphatidylserine is exposed to extracellular leaflet as a signal for engulfment by phagocytes (Nagata et al., 2010). Although there remain some controversies to be resolved, (Darland-Ransom et al., 2008), the exposure of phosphatidylserine residues on the extracellular leaflet is considered by some to be due to both inactivation of flippase activity (Segawa et al., 2014) and scramblase activation (Balasubramanian et al., 2007).

Scramblase is a ATP-independent enzyme that randomized phospholipid distribution across lipid membrane (Leventis and Grinstein, 2010).

Interferon gamma

We have discussed above the role of the Th1 cytokine TNF-α in initiating apoptosis, and will now comment on the other major Th1 cytokine, IFN-γ and its known role in affecting regulation of cell growth. Interferons (IFNs) are part of the human body’s defense against

microbes, viruses, and tumor cells (De Andrea et al., 2002). Interferons are a type of cytokine that was initially discovered to interfere with viral replication (De Andrea et al., 2002; Schroder et al., 2004). They are classified into Type I, Type II, and Type III interferons, based on receptor binding and sequence homology (Pestka et al., 2004). The Type I IFN includes IFN-α, IFN-β,

IFN-κ, IFN-ε, and IFN-ω; Type II includes IFN-γ only, and Type III includes three IFN-λ molecules (IFN-λ1, IFN-λ2, and IFN-λ3) (Vilcek, 2003; Pestka et al., 2004). Type I IFNs, mainly α and β are expressed ubiquitously and they bind to heterodimer receptor IFNAR, which consists of IFNAR1 and IFNAR2 chains (Schroder et al., 2004). IFN-γ was originally thought to be produced mainly by CD4pos T helper cell type 1 lymphocytes, CD8pos cytotoxic lymphocytes and Natural Killer cells (Young, 1996; Bach et al., 1997); however, now there is evidence that B cells and professional antigen presenting cells also secrete IFN-γ (Gessani and Belardelli, 1998;

Flaishon et al., 2000; Frucht et al., 2001). IFN-γ binds to IFN-γR, which consists of two subunits: IFN-γRI and IFN-γRII, (also known as IFNγ-Rα and IFNγ-Rβα) (Soh et al., 1994;

Pestka et al., 2004) and activates JAK/STAT intracellular signaling pathway (Darnell et al.,

1994). IFN-γ has been shown to up-regulate transcription expression of p21 and p27 (cyclin- dependent kinases inhibitors) and arrest cell cycle (Chin et al., 1996; Gooch et al., 2000; Takami et al., 2002). Upregulation of p21 and p27 leads to inhibition of cyclin E/CDK2 and cyclin

D/CDK4 complexes, which results in cell cycle arrest at G1/S phase boundary (Ekholm and

Reed, 2000). Inhibition of cyclin/CDK complexes results in hypophosphorylation of Rb, which sequesters E2F with the result that cell cycle cannot progress from G1 to S phase (Nevins, 2001).

How cell death and evidence of apoptosis are quantified in the laboratory

Because of the many defined biochemical processes that occur in cells as they undergo apoptosis, there are correspondingly numerous assays that can be performed to detect and

characterize these events. However, there is no single assay sufficient to positively identify when apoptosis is occurring. Instead, a battery of tests needs be performed, first to detect cell death, and then to demonstrate that the observed death is associated with multiple hallmarks of apoptosis.

Alamar Blue dye assay

Because our laboratory is testing large numbers of drugs in the search for candidates that will work cooperatively with the effectors of vaccination, we require a simple, fast and inexpensive assay compatible with high throughput efficiency in order to screen out unsuitable molecules and identify promising ones for further testing. Although assessing neither apoptosis nor cell death directly, the Alamar Blue assay serves as an ideal screening tool. Alamar Blue, is a water-soluble non-toxic dye that has been used in vitro to measures the viability of cells (Fields and Lancaster, 1993). Living cells are constantly metabolizing, through a series of oxidation- reduction reactions, and via these processes are capable of reducing the resazurin salt present in the Alamar Blue dye preparation to resofurin. This redox reaction changes the dye color from blue to pink (Figure 4). This transformation changes the absorbance properties of the dye which can be followed spectrophotometrically, and can thus be used as an indicator for cell metabolic activity and viability (Al-Nasiry et al., 2007). It should be noted, however, that lower metabolic activity in a test group may not always be synonymous with cell death. For example, a particular treatment over the course of several days could conceivably slow tumor cell proliferation without killing them. When the dye is added, the fewer cells in the test group would metabolize the dye more slowly than the control. To make certain that actual cell death is happening, another assay is employed, Trypan Blue staining.

Vital staining with Trypan Blue

Trypan Blue is a dye that stains dead cells since the cell membrane can no longer act as a barrier to prevent dye from entering the cells (Figure 5) (Louis and Siegel, 2011; Avelar-Freitas et al., 2014). Trypan Blue staining has traditionally been assessed by manually observing and enumerating stained cells microscopically, with the aid of a hemocytometer. This, however, is a slow, labor-intensive process incompatible with high-throughput screening, and is also subject to bias. Interestingly, Trypan Blue also has fluorescence properties, and this molecule can be excited by 642 nm laser via flow cytometry. The population of cells with higher fluorescent intensity represents those cells capable of taking up more Trypan Blue dye (i.e. dead cells).

Assessment of mitochondrial membrane potential using TMRE staining

Because mitochondrial breakdown is a feature occurring in both the intrinsic and extrinsic apoptotic pathways, tetramethylrhodamine ethyl esther (TMRE) staining can be regarded as one indicator of apoptosis. TMRE is a cationic fluorescent dye which accumulates in functional mitochondria due to their negative charge (Figure 6). The mitochondrial membrane potential must be maintained for TMRE to accumulated inside the cells (Christensen et al.,

2013). When cells undergo apoptosis, cytochrome C is released from the mitochondria, disrupting the electron transport chain and preventing maintenance of the mitochondrial membrane electrochemical potential. When this membrane potential is lost, the dye no longer sequesters there (Christensen et al., 2013). Thus, strong TMRE staining, as assessed by flow cytometry, is associated with viable cells, while low TMRE staining is a marker of apoptosis.

Analysis of plasma membrane structure and integrity using Annexin V and PI staining

Annexin V is a calcium-dependent phospholipid binding protein that binds to phosphatidylserine with high affinity (Van Engeland et al., 1998). Annexin V and propidium

iodide (PI) stains are often examined in combination to inspect if the death is due to apoptosis.

Annexin V can only bind extracellularly if the phosphatidylserines are exposed to the plasma membrane surface indicating early apoptosis (Figure 7) (Koopman et al., 1994). PI is an intercalating agent, and is impermeable to membranes of intact healthy cells; however, as cells die, PI is able to penetrate cells and bind double stranded DNA (Riccardi and Nicoletti, 2006).

When cells are stained positive for both Annexin V and PI, this indicates the cells are in late apoptosis (Figure 7).

Indirect assessment of Caspase 3 activity via PARP degradation

To further examine if the cell death was via apoptosis, PARP activity can be measured using western blot. PARPs are a family of enzymes that trigger the transfer of ADP-ribose to their target protein (Morales et al., 2014). PARPs are activated by single strand DNA breaks and are involved in base excision repair, nucleic acid metabolism, DNA synthesis, and ADP ribosylation of various substrate (D'amours et al., 1999). PARP-1 uses NAD+ as a substrate to catalyze poly ADP-ribosylation (Boulares et al., 1999). During apoptosis, cysteinyl-aspartate proteases (caspases) are activated to promote cell death (Li and Yuan, 2008). One of the hallmarks for apoptosis is the cleavage of PARP-1 by caspases (Kaufmann et al., 1993). Caspase

3 is primarily responsible for the cleavage of PARP during apoptosis (Tewari et al., 1995) .

Therefore, PARP cleavage stands as a surrogate marker for caspase 3 activity. Caspase 3 cleaves

PARP-1 into 89 kDa and 24 kDa fragments (Soldani and Scovassi, 2002). The 89 kDa fragment contains the catalytic domain of the enzyme and reduces DNA binding ability (Soldani et al.,

2001). The 24 kDa fragment binds irreversibly to damaged DNA and inhibits DNA repair enzyme binding (D’amours et al., 2001). Cleavage of PARP-1 leads to inactivation of poly(ADP-ribosylation) of cellular processes (Scovassi and Poirier, 1999), loss of activation

towards DNA damage (Soldani and Scovassi, 2002), and prevents NAD and ATP depletion

(Boulares et al., 1999). PARP levels can be detected using anti-PARP antibody by western blot.

Regulation of cell cycle in cancer using pharmacological agents

Normally, RTKs signals are tightly regulated in healthy cells allowing cell proliferation only when it is necessary. Modes of regulation include degradation of signaling ligands, as well as dephosphorylation, internalization via endocytosis and ubiquitination and turnover of the

RTKs themselves (Wiley and Burke, 2001). However, in cancer cells RTK regulation is often circumvented through mutation or overexpression, permitting constant proliferation (Sangwan and Park, 2006). Under these circumstances, RTKs act as oncogenes. For example, HER-2 is overexpressed in 10-30% of breast cancer patients (Iqbal and Iqbal, 2014) and cyclin D1 protein is overexpressed in 50% of human breast cancer (Mohammadizadeh et al., 2013). Targeting

RTKs and different mitogenic proteins involved in the cell cycle progression have become attractive strategies for developing anti-cancer drugs. Two drugs targeting mitogenic pathways that have recently been developed are palbociclib (ibrance) and sunitinib (sutent).

Palbociclib

Palbociclib (ibrance) is a small molecule selective inhibitor for CDKs 4/6 (Cadoo et al.,

2014), which is taken orally. Ibrance consists of group of pyridopyrimidine compounds developed due to its promising pharmaceuticals and physical properties (Fry et al., 2004) (Figure

8). For CDK4/6 to be activated, cyclin D1 needs to bind and form an activate complex that can phosphorylate Rb. Ibrance is very potent and selectively inhibits both CDKs 4/6 kinase activity, with little effect on other protein kinases (Fry et al., 2004). Suppression of the cyclin

D1/CDK4/6 axis leads to reduction of phosphorylation of Rb protein and arresting cells in G1

(Dean et al., 2010), as shown in Figure 9. Ibrance has been shown to decrease DNA replication

and suppresses cellular proliferation at low concentrations in Rb-positive cancer cell lines

(Cadoo et al., 2014). Ibrance is FDA-approved and has been in clinical trials for treatment of estrogen-positive breast cancer (Finn et al., 2009; Schwartz et al., 2011; Finn et al., 2015).

Ibrance has also shown antitumor effect in other human tumor cell lines including bladder cancer

(Pan et al., 2017), renal cell carcinoma (Logan et al., 2013), and mantle cell and B cell lymphoma (Leonard et al., 2012). For breast cancer, ibrance has been demonstrated to work best in conjunction with other agents such as anti-estrogen drugs and the HER-2-targeting monoclonal antibody drug trastuzumab. Furthermore, ibrance seems to sensitize cells to chemotherapy and radiation therapy (Finn et al., 2009).

Sunitinib

Sunitinib malate (Figure 10) is an oral small molecule inhibitor that interferes with the activity of multiple RTKs (Figure 11). Sunitinib has shown to potently inhibit VEGFR, KIT,

FLT3, and PDGFR (Mendel et al., 2003) making it less selective than palbociclib. In vitro studies have demonstrated that sunitinib inhibits the growth of human umbilical vein endothelial cells that are dependent on VEGF, PDGF and stem-cell factor (SCF) Sunitinib demonstrated both time- and dose-depended antitumor activities in human tumor xenograft models (Mendel et al., 2003) for colon (Morimoto et al., 2004), glioma, melanoma, and breast (Mendel et al., 2003) cancer cell lines. It has also been in clinical trial for breast cancer patients and has been examined in combinations with other drugs such as trastuzmab (Bachelot et al., 2014), docetaxel

(Bergh et al., 2012), and capecitabine (Crown et al., 2013). These combinations of sunitinib with another drug resulted in better outcomes than sunitinib treatment alone (Bachelot et al., 2014).

Hypotheses and Aims:

The dendritic cell-based vaccine technology developed by our laboratory and its collaborators targets the HER-2 oncodriver through the generation of powerful Th1-polarized immune responses. Clinically, the vaccine effect includes complete pathological responses in some patients (where tumor cells are clearly destroyed since the tumor entirely disappears) and strongly suppressed HER-2 expression in about 50% of patients with remaining disease. In vitro studies show that Th1 cytokines mimic vaccine effects in vitro (a moderate level of induced apoptosis coincident with partial inhibition of HER-2 expression as well as other HER-family members). In this regard, our vaccine can be considered a therapy that works in part by interfering with HER-2 signaling. In addition, small molecule inhibitors of estrogen signaling enhances vaccine effects for hormone receptor-expressing individuals, suggesting that judicious combinations of vaccine and drug can leverage the therapeutic effects of vaccination. Small molecule inhibitors of other mitogenic pathways including various RTKs (sunitinib) and the cyclin-dependent kinases (palbociclib) have already been shown most effective when combined with HER-2 targeted drugs such as trastuzumab. We therefore hypothesize that the drugs sunitinib and palbociclib will work in concert with Th1 cytokines to potentiate apoptotic cell death in a variety of human breast cancer cell lines. This thesis has two Specific Aims:

Aim 1: To determine the effects of combined Th1 cytokine plus palbociclib, an inhibitor of cyclin-dependent kinases 4 and 6, on human breast cancer cell lines of diverse phenotypic characteristics. We hypothesize that the combination of Th1 cytokines plus the cyclin-dependent kinase inhibitor palbociclib (ibrance) will show either additive or synergistic effects to enhance the expression of characteristic features of apoptotic cell death.

Aim 2: To determine the effects of combined Th1 cytokines and the broader-spectrum receptor tyrosine kinase (RTK) inhibitor, sunitinib on human breast cancer cell lines of diverse phenotypic characteristics. We hypothesize that the combination of Th1 cytokines plus sunitinib will show either additive or synergistic effects to enhance expression of characteristic features of apoptotic cell death.

Figure 1: Growth factor activated receptor tyrosine kinases pathway.

Binding of growth factor (GF) to receptor (RTKs) leads to dimerization and transphosphorylation by the tyrosine kinase (TK) domain. Grb2 bind at the site of phosphorylate

(P) TK domain and binds guanine exchange factor (GEF) causing GDP to GTP exchange by Ras protein. Activated Ras further induces downstream proteins and eventually activating cyclin D1, which bind and form complex with CDK4/6. This cyclin D1 and CDK4/6 complex leads to phosphorylation of retinoblastoma protein (Rb) and frees E2F.

Figure 2: Schematic of intrinsic and extrinsic apoptosis pathways.

DNA damage activates p53 that could lead to intrinsic pathway activation of cytochrome c release. Cytochrome c binds seven Apaf-1 proteins to form an apoptosome. The apoptosome cleaves and activates procaspase 9 leading to activation of caspase 3, 6, and 7. Extrinsic pathways is activated by extracellular signaling agents that bind death receptors (DR) and form a

DISC complex. DISC cleaves caspase 8 function that leads to activation of Bid and cleavage of procaspase 3, 6, and 7 to active form.

Figure 3: Schematic of caspase 3 cleavage of PARP.

Pro-caspase 3 which is cleaved and activated by caspase 9 leads to the proteolytic inactivation of

PARP and induction of apoptosis. However, when levels of DNA damage are low, PARP remains intact and promotes DNA repair.

Figure 4: Alamar Blue staining to determine cellular metabolic activity.

In live cells, resazurin sodium salt (blue color) is reduced to resorufin sodium salt (pink color).

This conversion can be monitored spectrophotometrically by determining optical density at 630 nm.

Figure 5: Trypan Blue staining to distinguish living from dead cells.

Trypan Blue dye is a non-permeable to live cells, but it can enter and accumulate in dead cells which can be observed microscopically as a blue color, or can be detected via flow cytometry using a 642nm excitation laser.

Figure 6: TMRE staining to determine mitochondrial membrane potential.

Cells with functional mitochondria have a strong negatively charged mitochondrial membrane and are stained with TMRE (positively charged) dye, imparting to the cells a strong fluorescent signal. Cells undergoing apoptosis lose this charge, so little dye accumulates, producing a weak fluorescent signal.

Figure 7: Annexin V/ Propidium iodine (PI) staining examining apoptosis.

In healthy cells, the membrane component phosphatidylserine is located primarily on the inner leaf of the plasma membrane. When cells undergo apoptosis, these residues become exposed on the outer leaflet of the plasma membrane. Annexin V (AV), which is substituted with a fluorochrome, binds to phosphatidylserine, imparting to the cell a strong fluorescent signal by itself indicative of early apoptosis. PI can only enter cells with degrading membrane and bind strongly to DNA, imparting fluorescence at a different wavelength than AV. Simultaneous staining with both AV and PI indicates late apoptosis.

Figure 8: Ibrance chemical structure. Image adapted from (Dhillon, 2015).

Figure 9: Cell cycle.

Several kinases and cyclins control transition of the cell cycle from one phase to the next. Cell replication progresses from G1 (Growth phase) to S (DNA replication) to G2 (growth phase 2) and then to M (mitosis phase). For transition from G1 to S phase, Rb (retinoblastoma) protein needs to be phosphorylated by CDK4/6 complex with cyclin D, which is inhibited by palbociclib, and results in cell cycle arrest (figure modified from (Lutful Kabir et al., 2016)]).

Figure 10: Sunitinib chemical structure.

Figure 11: Sunitinib mechanism of action.

Sunitinib blocks tyrosine receptor kinases such as vascular endothelial growth factor receptor

(VEGFR), stem cell factor receptor (KIT), platelet-derived growth factor receptor (PDGFR), and fetal liver tyrosine kinase receptor 3 (FLT3).

Group 1: Group 2: Group 3: Group 4: Luminal A Luminal B HER-2pos Basal-like

ERpos ERpos ERneg ERneg

PRpos PRneg PRneg PRneg

HER-2neg HER-2pos HER-2pos HER-2neg

Table 1: Breast cancer classification. Estrogen receptor (ER), progesterone receptor (PR), human epidermal growth receptor 2 (HER-2)

Chapter 2

Methods

Reagents and Cell Culture:

Palbociclib (PD0332991, Pfizer) was resuspended in water and diluted in cell media or

PBS for in vitro use. Sunitinib (SU11248, Pfizer) was resuspended in dimethyl sulfoxide

(DMSO) and diluted in cell media or PBS for in vitro use. Human breast cancer cell lines MDA-

MB-231, MDA-MB-468, SKBR-3, and MCF-7 were obtained from American Type Culture

Collection (ATCC) (Manassas, VA), JIMT-1 obtained from DSMZ (Braunschweig, Germany), and HCC-1419 obtained from Dr. Czerniecki lab. The MDA-MB-231, HCC-1419, and MDA-

MB-468 cell lines were cultured in RPMI media (Sigma-Aldrich, St. Louis, MO). MCF-7 cell lines were cultured in EMEM media (ATCC; Manassas, VA). JIMT-1 cell lines were cultured in

DMEM media (VWR; Sanborn, NY). SKBR-3 cells were cultured in McCoy’s 5A media (Gibo;

Rockville, MD). In all media, 10% fetal bovine serum (FBS) (Atlantic biologics), 1% Pen/strep

(Gibco), 1% sodium pyruvate, 1% NEAA and 1% glutamine were added and cultured at 37˚C and 5% CO2 humidified atmosphere.

Alamar Blue Assay:

Breast cancer cell lines were plated in 96-well cluster plates at densities ranging from 5-

7*103 cells per well depending on the growth characteristics of individual lines. After 12 hours of initial culture, cells were treated with Th1 cytokines (TNF-ɑ and IFN-ɣ; 5 ng/mL each) and either ibrance (0.325-80 µM) or sunitinib (0.325-40 µM), then cultured for 72 additional hours, at which time 20 µL of 0.15 mg/mL of resazurin in PBS (Alamar Blue dye) was added to each

well. After 6-7 hours of additional culture, the optical density of the culture supernatants was measured at 630 nm using a BioRad (model number) microplate reader using Gen5 software.

Viability staining with Trypan Blue:

Breast cancer cells were seeded at densities 5*104 in 12-well cluster plates and incubated overnight. The next day the cells were treated with cytokines (5ng/mL each TNF-α and IFN-γ), drugs (either 10 μM ibrance or 10 μM sunitinib), combinations of single drugs plus cytokines, or left untreated. After 72 hours further incubation, cells were harvested by scraping, and 5 μL

Trypan Blue dye added in about 20 to 30 μL volume of cell suspension. Dye uptake assessed via flow cytometric analysis using 642 nm laser with and analysis performed using IDEAS version

6.2 software.

Mitochondrial membrane potential determination via TMRE staining

Cultured breast cancer lines were treated as describe for Trypan Blue studies. Thirty minutes prior to harvest, 10μL/mL of tetramethylrhodamine, ethyl ester (TMRE) was added to each well of cultured cells, and cells collected by scraping. Dye uptake in mitochondria was determined via flow cytometry using 488 nm excitation laser and analysis performed using

IDEAS version 6.2 software. Because Trypan Blue and TMRE have discrete fluorescent properties, for some experiments, cells were double-stained, and analyses performed simultaneously.

Annexin V/propidium iodide staining:

Breast cancer cell lines were treated and cultured as described for Trypan Blue studies.

At 72 hours post-treatment, cells were harvested by scraping and washed with Annexin V staining buffer. Cells were then stained with fluorochrome-labeled Annexin V (3 μL was added in 30-40 μL) at 4˚C for 15 minutes, followed by the addition of propidium iodide (PI) (0.5 μL)

immediately prior to analysis using a Flowsight imaging flow cytometer running IDEAS version

6.2 software.

Mitochondria Cytochrome C assessment of apoptosis

SKBR-3 cell lines were treated as described for Trypan Blue studies. At 72 hours post- treatment cells were harvested with the aid of trypsin and resuspended in 100 μL ice-cold permeabilization buffer (100 mM KCL, 100 μg/mL digitonin in PBS). These were incubated for

5 minutes and 100 μL paraformaldehyde (4% in PBS) was added to the permeabilized cells.

Samples were centrifuged at 500 RPM at 4˚C for 5 minutes followed by removal of the supernatant. Cells were then incubated with 4% paraformaldehyde for 20 minutes at room temperature. Cells were then washed three times in 200 μL PBS (700 RPM for 5 min at 4 ˚C).

Cells were resuspended in 200 μL blocking buffer (0.05% Saponin and 3% BSA in PBS) and incubated for 1 hour at RT. APC-conjugated anti-cytochrome c (1:200) was add and incubated overnight at 4˚C. Next day, cells were washed three times in 200 μL PBS (700 RPM for 5 min at

4 ˚C). Anti-mouse secondary (1:200) antibody was added, diluted in 100 μL blocking buffer and incubated for an hour at RT. Cells were washed three more times in 200 μL PBS (700 RPM for 5 min at 4 ˚C). Expression of cytochrome c was analyzed using a Flowcyto imaging cytometer employing a 642 nm and Ideas software suite version 6.2.

Assessment of HER-2 surface expression

Breast cancer cell lines were treated as described for Trypan Blue studies. At 72 hours post-treatment, cells were harvested by scraping, washed in PBS and resuspended in 1 mL FACS buffer (PBS+1%FBS+0.01% sodium azide). APC-conjugated anti-HER-2 antibody was then added for 15 minutes at 4˚C and cells analyzed the HER-2 expression by flow cytometry using

642 nm excitation laser on a Flowsight imaging cytometer running Ideas 6.2 analysis software.

Western Blot:

SKBR-3 cells were seeded at 3*106 cells/mL in 10mL total volume in a petri dish. The next day, cells were treated with 5ng/mL of Th1 cytokines, 10uM Ibrance, or both combined.

After incubation for 72 hours, samples were collected. Harvested cells were centrifuged at 800

RPM for 5 minutes and supernatants removed and discarded. The cell pellets were lysed using

100 µL of RIPA lysis buffer (50 mM Tris-base, 150mM NaCl, 1% Trition-X 100, 0.5% Sodium deoxycholate and 0.1% sodium dodecyl sulfate) with added protease inhibitor cocktail (Pierce) and PhosStop phosphatase inhibitors cocktail (Roche). These extracts were centrifuged at 20,000

RPM for 20 minutes at 4˚C. Supernatants were collected and stored at -80˚C prior to analysis.

The BCA protein assay was used to generate a standard curve to estimate the amount of proteins in each extract. Extracts were diluted in 5X loading buffer that contained β- mercaptoethanol and glycerol. Samples were vortexed briefly and boiled for 5 minutes to denature the proteins. Thirty micrograms total protein from each sample were loaded into 4-15%

Mini Protean TGX gels (Bio-Rad) and separated for 40 minutes at 160 volts. Proteins from the gel were electrotransferred onto PVDF membrane (Bio-Rad) at 100 volts for an hour.

Membranes were washed 3 times with TBS-T (20mM Tris, 150mM NaCl, and 0.1%Tween 20 with pH of 7.6) at 5-minute intervals. The membranes were blocked with 5% milk for 30 minutes and incubated overnight with primary antibodies: 1:1000 anti-phospho-Rb anti-rabbit (Cell

Signaling, Denver MA), 1:1000 total-Rb anti-mouse (Cell Signaling, Denver MA), 1:1000 PARP anti-rabbit (Cell Signaling, Denver MA), and 1:1000 anti-β-actin anti-mouse (GenScript,

Piscataway, NJ). The membrane was washed with TBS-T buffer and appropriate secondary HRP tagged antibody was added and incubated at room temperature for an hour. It was followed by washing the membrane 3 times with TBS-T at 5-minute intervals. The membrane was developed

by using SuperSignal West Pico Chemiluminescent Substrate (Thermo Scientific).

Chemiluminescent detection was performed using Fuji LAS 3000 detection system (R&D

Systems, Minneapolis, MN). Band intensity and fold change were quantified using ImageJ analysis.

Statistical Method:

One-way Analysis of Variance (ANOVA) was performed for statistical analyses; p-value of less than 0.05 was considered significant. The statistical significance between treatment groups was determined using the Tukey’s Honest Significant Difference test. One-way ANOVA was analysis using SigmaPlot 12.5 software (Systat Software Inc., San Jose, CA).

Chapter 3

Results

Aim 1:

Th1 cytokines and ibrance work together to maximize suppression of breast cancer metabolism.

To screen breast cancer lines for drug sensitivity, and to determine optimal doses of ibrance and Th1 cytokines to use in the ongoing studies, the Alamar Blue assay was performed on three different breast cancer cell lines: MDA-MB-468, SKBR-3, and JIMT-1. These cell lines were treated with 5ng/mL Th1 cytokines and different concentrations of ibrance ranging from

0.3125 to 80uM. Cells were cultured for 72 hours followed by addition of Alamar Blue dye, which helps estimate relative levels of cellular metabolism by spectrophotometrically following the reduction resazurin salt to resofurin. Initial dose response curves indicated that a concentration of 10μM was the optimal dose of ibrance for all three cell lines (lower optical densities correspond to higher metabolic activity). This dose of drug was then used to confirm

Alamar Blue results (Figure 12) and throughout the studies. For all examined cell lines, the combination of ibrance and Th1 cytokines led to a level of metabolic suppression significantly greater (p<.01) than that achieved by either treatment alone (Figure 13).

Th1 cytokines plus ibrance potentiate cell death.

The Alamar Blue assay detects suppression of metabolic activity in cells. However, it does not prove that any observed suppression in metabolism is a consequence of cell death. To confirm cell death, Trypan Blue staining was performed. Cells were exposed to either Th1

cytokines, ibrance, or both for 72 hours, or were left untreated. After 72 hours incubation, microscopic examination of cells suggested massive cell death; all treatment groups displayed fewer viable cells and high levels of subcellular debris, with remaining viable cells exhibiting altered morphological features (Figure 14). Cells that were exposed to both treatments seemed to be most affected. These visual observations were confirmed when the remaining cells were harvested and stained with Trypan Blue, which is excluded from living cells but retained in dead ones, imparting a strong fluorescent signal that can be detected via flow cytometry.

For the three tested cell lines, flow cytometric data of cellular fluorescence of each treatment group is displayed in histogram form (Figure 15). During analysis, a gate was established to distinguish low-staining (living) from high-staining (dead) cells, allowing a percentage cell death assignment for all treatment groups. For example, in one representative experiment, MDA-MB-468 cell death was shown to be highest with combined treatment:

Untreated (13%), Th1 cytokines (36%), ibrance (24%), and Th1 cytokines+ibrance (61%).

SKBR-3 showed similar responses: untreated (25%), Th1 cytokines (48%), ibrance (58%), and

Th1 cytokines+ibrance (80%). JIMT-1 followed the same pattern: 12% Untreated, 34% Th1 cytokines, 26% ibrance, and 56% for Th1 cytokines plus ibrance (Figure 15).

We also analyzed flow cytometry data by overall fluorescent intensity (mean channel fluorescence). Composite data from at least three separate representative experiments are shown for Figure 16. For all breast cancer cell lines tested, the combination of Th1 cytokines plus ibrance led to significantly greater uptake of fluorescent dye (p<0.05) than either treatment alone

(Figure 16). Taken together, these data show that the combination of Th1 cytokines and ibrance significantly increase cell death over either treatment alone.

Th1 cytokines and ibrance combination induce cell death via an apparent apoptotic mechanism.

To determine if the observed cell death occurred through an apoptotic mechanism, breast cancer cell lines were analyzed via two fluorescence-based assays that detect apoptotic features:

TMRE and Annexin V/propidium iodide (PI) staining. TMRE dye is sequester by mitochondria in healthy, metabolically active cells leading to a strong fluorescent signal. In contrast, apoptotic cells do not take up TMRE dye and will be of low fluorescent intensity. With Annexin V/PI assay, fluorochrome-labeled Annexin V binds to phosphoserine residues that gets flipped to the outer leaflet of the plasma membrane during early apoptosis, while PI can enter the nuclei and bind efficiently to fragmenting DNA in late apoptotic cells.

For analysis of TMRE results, data for each treatment group were initially arranged in histogram format (Figure 17). A gate was defined to enumerate high-staining (viable, non- apoptotic cells). In the representative experiment, for MDA-MB-468 cells, the proportion of high fluorescent staining (viable) cells were 52%, 12%, 47% and 2.3% for untreated, cytokine-treated, ibrance-treated and combined treatment, respectively. For SKBR-3, the values were 67%, 41%,

38% and 14%. For JIMT-1 cells, the values were 66.1%, 48.8%, 37.4% and 20.8%, respectively.

We also analyzed flow cytometry data by mean channel fluorescence to look for overall loss of fluorescent staining, indicative of apoptotic changes in the treated cells. Composite data from at least three different experiments demonstrated that for each cell line, combined treatment with Th1 cytokines and ibrance lead to significantly greater (p<0.05) losses in fluorescent staining compared with cells exposed to cytokines or drug alone (Figure 18).

For the Annexin V/PI assay with MDA-MB-468 cells, the proportion of cells that stained brightly for both Annexin V and PI (late apoptotic) were 31.2% for untreated, 42.8% (Th1

cytokines), 46.6% (ibrance), and 55.6% for dual treatment (Figure 19 left panel). For JIMT-1 cells, these values were 7.3%, 37.4%, 16.9%, and 48.7%, respectively (Figure 19 middle panel).

Similarly, the values for SKBR-3 cell line (Figure 19 middle panel) were 17.8%, 34.7%, 26%, and 51.8%, respectively. These results demonstrate that dual treatment of breast cancer cell lines with Th1 cytokines and ibrance resulted in significantly more apoptotic cell death than either treatment alone.

Th1 cytokines and ibrance combination shows decrease total Rb protein.

To determine whether the combination of Th1 cytokines and ibrance led to decreased total Rb production, western blot analysis was performed. SKBR-3 cells were treated as described for the Trypan Blue assay, and cells were harvested and subsequently lysed and equal amounts of protein from each treatment group separated by SDS-PAGE, transferred to PVDF membranes and blotted with anti- Rb and actin (loading control) antibodies (Figure 20A). Rb protein has an expected mobility of 110 kDa. It is clear from the representative blot that dual treatment leads to strongly suppressed levels of Rb protein, an impression verified by composite densitometry analysis of three experiments (Figure 20B) demonstrating that suppressed Rb levels resulting from dual treatment were significant (p<0.01) over either Th1 cytokines or drug alone.

We were surprised, however, when we probed similar blots with a phospho-specific anti-Rb antibody to determine the activation state of the remaining Rb protein. Here, we consistently observed a major band with an apparent molecular mass of 160 kDa rather than the expected 110 kDa (Figure 21A). We attempted to resolve this question by purchasing a different phospho- specific Rb antibody, but unfortunately, identical results with an aberrantly high molecular weight major band were obtained (Figure 21B). Because of this consistently observed mass discrepancy, we regrettably cannot conclude anything about phosphorylated Rb protein.

Figure 12: Combination of Th1 cytokines and ibrance enhance metabolic suppression of breast cancer cells.

SKBR-3, JIMT-1, and MDA-MB-468 cells were seeded at 5x103 per well and were left untreated

(No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and Th1 cytokines for 72 hours. After 72 hours, Alamar Blue dye was added to each well and optical density of the supernatant read spectrophometrically at 630nm.

Figure 13: Metabolic activity of breast cancer cell lines is maximally suppressed by dual

Th1 cytokine and ibrance treatment.

SKBR-3, JIMT-1, and MDA-MB-468 cells were seeded at 5x103 per well and were left untreated

(No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and Th1 cytokines. After 72 hours of incubation, Alamar Blue dye was added to each well and after a period of incubation, after color change became apparent, optical density of the supernatant read spectrophometrically at 630nm. Data represent summary analysis of at least three separate experiments per cell line. Statistical significance determined by one way ANOVA: SKBR-3

[F(3,20)=9.2, p<0.001], JIMT-1 [F(3,56)=62.8, p<0.001], and MDA-MB-468 [F(3,51)=51.9, p<0.001] followed by Tukey test. Significance was set at **p<0.01 and *** p<0.001. Error bars show the standard error of the mean.

Figure 14: Microscopic examination of treated cells reveals evidence of cell death.

SKBR-3, MDA-MB-468, and JIMT-1 cells were seeded at 5*104 per mL and were left untreated

(No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and Th1 cytokines for 72 hours. Photomicrographs were taken using an Olympus CK2 microscope with

20X objective lens.

Figure 15: Histogram analysis of Trypan Blue staining reveals enhanced cell death with combined Th1 cytokine and ibrance treatment.

MDA-MB-468, SKBR-3, and JIMT-1 cells were seeded at 5*104 per mL and were left untreated

(No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and Th1 cytokines for 72 hours. Cells were then harvested and stained with Trypan Blue dye, after which the imparted fluorescent signal was assessed by flow cytometry using 642nm excitation laser.

Figure 16: Combination of Th1 cytokines and ibrance significantly increase breast cancer cells death.

MDA-MB-468, JIMT-1, and SKBR-3 cells were seeded at 5*104 per mL density and were left untreated (No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and

Th1 cytokines for 72 hours. Afterwards, cells were harvested and stained with Trypan Blue dye, which was assessed by flow cytometry using 642nm laser. Data represent summary analysis of mean channel fluorescence of treated, stained cells for at least three separate experiments per cell line. Statistical significance determined by one way ANOVA: MDA-MB-468 [F(3,96)=52.4, p<0.001], JIMT-1 [F(3,20)=19.7, p<0.001], and SKBR-3 [F(3,8)=32.1, p<0.001] followed by post hoc Tukey test. Significance was set at *** p<0.001, **p<0.01 and *p<0.05. Error bars represent the standard error of the mean.

Figure 17: Ibrance plus Th1 cytokines causes maximized decrease mitochondria membrane potential, an indicator of apoptosis.

MDA-MB-468, SKBR-3, and JIMT-1 cells were seeded at 5*104 per mL and were left untreated

(No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and Th1 cytokines for 72 hours. Afterwards, cells were harvested and stained with teramethylrhodamine ethyl ester (TMRE 100nM) dye, with intensity of imparted fluorescence assessed by flow cytometry using a 488nm excitation laser. Histogram analysis displays results from a signal representative experiment. Percentages represent metabolically active (i.e., strongly staining) cells for different treatment groups.

Figure 18: Combination of Th1 cytokines plus ibrance significantly decrease mitochondria membrane potential over single treatments for breast cancer cells.

MDA-MB-468, JIMT-1, and SKBR-3 cells were seeded at 5*104 per mL and were left untreated

(No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and Th1 cytokines for 72 hours. Next, cells were harvested and stained with teramethylrhodamine ethyl ester (TMRE 100nM) dye, which was assessed by flow cytometry using 488nm laser. Data represent composite data of mean channel fluorescence for each cell line and treatment group derived from at least three separate experiments. Statistical significance determined by one way

ANOVA: MDA-MB-468 [F(3,48)=51.1, p<0.001], JIMT-1 [F(3,47)=22.8, p<0.001] and SKBR-

3 [F(3,32)=45.9, p<0.001] followed by post-hoc Tukey test. Significance was set at *** p<0.001,

**p<0.01 and *p<0.05. Error bars represent the standard error of the mean.

Figure 19: Combination of ibrance and Th1 cytokines maximizes markers of apoptosis in breast cancer cell lines.

MDA-MB-468, JIMT-1, and SKBR-3 cells were seeded at 5*104 per mL and were left untreated

(No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and Th1 cytokines for 72 hours. Next, cells were harvested and stained with FITC-Annexin V and propidium iodide (PI), which was assessed by flow cytometry. Two-color dot plot analysis represents percentage of gated cells in each quadrant. Double staining with both Annexin V/PI represents late apoptotic events.

A. kines

NoRx

cyto Th1

Ibrance Both 100 Total RB 110kDa 75 50 Actin 42 kDa 37

Figure 20: Combined Th1 cytokine and ibrance treatment minimizes levels of total Rb protein in SKBR-3 cells.

SKBR-3 cells were seeded at 3*106/mL in 10mL total volume and were left untreated (No Rx) or treated with Th1 cytokines (5ng/mL), or ibrance (10uM) or both ibrance and Th1 cytokines.

After 72 hours of incubation, cells were harvested and extracted in RIPA buffer in the presence of protease and phosphatase inhibitors. Protein lysate (30ug/well) was separated on a 4-15%

SDS-PAGE gel before transferring to nitrocellulose membrane. A. Western blot probed with anti- total Rb protein (110 kDa), anti-Actin (42 kDa, loading control) and bands visualized using chemiluminescence with Fuji LAS 3000 detection system B. ImageJ analysis of western blot image that was normalized to loading control. Data represent at least three separate experiments.

Statistical significance determined by one way ANOVA SKBR-3 [F(3,8)=19.5, p<0.001] followed by Tukey test. Significance was set at *** p<0.001, **p<0.01 and *p<0.05. Error bars represent the standard error of the mean.

A. B.

Figure 21: Detection of phosphorylated Rb protein in SKBR-3 cells.

SKBR-3 cells were seeded at 3*106/mL in 10mL total volume and were left untreated (No Rx) or treated with Th1 cytokines (5ng/mL), ibrance (10uM) or both ibrance and Th1 cytokines for 72 hours. After 72 hours incubation, cells were harvested and extracted in the RIPA buffer in the presence of protease and phosphatase inhibitors. Protein lysate (30ug/well) was separated on a 4-

15% SDS-PAGE gel before transferring to nitrocellulose membrane. A. Western blot probed with anti-phosphorylated Rb protein (160 kDa 9307-S antibody) and Actin (42 kDa, loading control). B. Western blot probed with a different anti-phosphorylated Rb protein (160kDa, 8180-

T antibody) and Actin 42 kDa (loading control). Bands were detected using chemiluminescence with Fuji LAS 3000 detection system.

Aim 2:

Th1 cytokines and sunitinib work together to suppress breast cancer cell metabolism.

To screen breast cancer lines for drug sensitivity, and to determine optimal doses of sunitinib and Th1 cytokines to use in the rest of the studies, Alamar blue assay was performed on four different breast cancer cell lines: MDA-MB-231, MDA-MB-468, MCF-7, and SKBR-3.

These cell lines were treated with 5ng/ml Th1 cytokines and graded concentration of sunitinib ranging from 0.3125 to 40uM (Figure 22). Initial dose response curves indicated that a concentration of 10μM was optimal dose of sunitinib for all cell lines examined (lower optical density correspond to higher metabolic activity). This dose of drug was then used to confirm

Alamar Blue results (Figure 22) and throughout the remaining studies. We added fifth line HCC-

1419, all five cell lines treated with the combination of Th1 cytokines (5ng/mL) and sunitinib

(10uM) had significantly greater suppression of the cell metabolic activity compared to Th1 cytokines or sunitinib alone (Figure 23; p<0.05).

Th1 cytokines plus sunitinib maximize cell death.

Alamar Blue assay detects suppression of metabolic activity in cells. However, it does not prove that any observed suppression in metabolism is a consequence of cell death. To confirm cell death, Trypan Blue staining was performed. Cell lines MDA-MB-231, MDA-MB-

468, MCF-7, HCC-1419, and SKBR-3 were exposed to either Th1 cytokines, sunitinib, or both for 72 hours or left untreated (No Rx). After 72 hours of incubation, microscopic examination of cells suggested massive cell death; all treatment groups displayed fewer viable cells, high levels of subcellular debris, with remaining viable cells exhibiting altered morphological features

(Figure 24). Cells that were exposed to both treatments seemed to be most affected.

These visual observations were confirmed when the remaining cells were harvested and stained with Trypan Blue, which is excluded from living cells but retained in dead ones, imparting a strong fluorescent signal which can be assessed via flow cytometry. For the five tested cell lines, flow cytometric data of cellular fluorescence for each treatment group is displayed in histogram from (Figure 25). During analysis, a gate was established to distinguish low-staining (living) from high staining (dead) cells, allowing a percentage cell death assignment for all treatment groups. For example, in one representative experiment MDA-MB-468 cell death was shown to be highest with combined treatment: untreated (17.7%), Th1 cytokines (37.4%), sunitinib (52.3%), and both treatment (91.1%). MDA-MB-231 showed similar results: untreated

(11.9%), Th1 cytokines (27%), sunitinib (29.6%), and both (95.1%). For SKBR-3 cells, 8.4% dead for untreated, 43.1% for Th1 cytokines, 51.8% for sunitinib and 78.3% for both treatment.

HCC-1419 cell death went from 5% untreated, 20.8% Th1 cytokines, 56.8% sunitinib, and

87.9% for both treatment combinations. Like other cell lines similar results were found for estrogen receptor- positive cell lines, MCF-7 with 31.2% dead in untreated, 56.7% Th1 cytokines, 59.4% sunitinib, and 86.4% in both treatment combinations (Figure 25).

We also analyzed the flow cytometry data by overall fluorescent intensity (mean channel fluorescence). Composite data from at least three separate experiments are illustrated in Figure

(26). For all breast cancer cell lines tested, the combination of both Th1 cytokines plus sunitinib led to significantly greater uptake of fluorescent dye (p<0.05) than either treatment alone (Figure

26). Taken together, these data demonstrated that the combination of Th1 cytokines plus sunitinib significantly increase cell death over either treatment alone.

Th1 cytokines combined with sunitinib potentiate cell death via apoptosis.

To determine if the observed cell death occurred through an apoptotic mechanism, breast cancer cell lines were examined with Annexin V/propidium iodide (PI) staining, a flow cytometry-based assay. For MDA-MB-468 cells the proportion of cells that stained brightly for both Annexin V and PI (late apoptotic) were: untreated 12.6%, Th1 cytokines 28%, sunitinib

46.6%, and both treatment 81.3% (Figure 27 left panel). For MDA-MB-231 cell lines these values were untreated 9.45%, Th1 cytokines 25.4%, sunitinib 36.4%, and both treatment combinations were 51.4% (Figure 27 left middle panel). SKBR-3 cell lines these values were

17.8% (untreated), 34.7% (Th1 cytokines), 47.3% (sunitinib), and 58.4% (both treatments)

(Figure 27 middle panel). HCC-1419 cell lines these values were 17% (untreated), 22% (Th1 cytokines), 22.2% (sunitinib), and 34.8% (both treatments). Similarly, estrogen positive cell lines

MCF-7 represented 33% (untreated), 51.4% (Th1 cytokines), 39.4% (sunitinib), and 77.1% (both treatments) (Figure 27 right panel). These results demonstrate that dual treatment of breast cancer cell lines with Th1 cytokines and sunitinib resulted in enhanced apoptotic cell death than either treatment alone.

Th1 cytokines combined with sunitinib minimizes expression of surface HER-2 over single treatments.

Previous studies had shown that Th1 cytokine treatment led to lower levels of HER-2 expression. To determine whether the addition of sunitinib enhanced this suppression, surface

HER-2 levels were examined in the SKBR-3 cell line. At 72 hours post-treatment cells were harvested and stained with HER-2 antibody and analyzed by flow cytometry with results from a single representative experiment displayed in histogram form (Figure 28A), and composite data from 3 separate experiments analyzed by mean channel fluorescence (Figure 28B). Histogram

data reveal an increasing leftward shift of the curve, indicating suppressed HER-2 expression, which is maximally achieved when SKBR-3 cells were exposed to both Th1 cytokines and sunitinib. Analysis of mean channel fluorescence from three separate experiments verified that the differences seen with dual treatments were significant over single treatments (Figure 28B; p<0.05).

Combined treatment using Th1 cytokines and sunitinib lead to lowered cytochrome C levels.

To examine apoptosis further, mitochondria cytochrome c levels were measured in

SKBR-3 cell lines. Cells were harvested, permeabilized (in a solution containing KCl and digitonin), and stained with anti-cytochrome c antibody. The permeabilizing solution disrupts plasma membrane integrity, allowing cytochrome c released from the mitochondria to leach out of the cell. However, the digitonin in the buffer does not compromise mitochondrial membranes, allowing for retention and staining of cytochrome c in these organelles. This difference allows only cytochrome c associated with the mitochondria to be measured (Christensen et al., 2013).

For the SKBR-3 cell lines, flow cytometric data of cellular fluorescence of each treatment group is displayed in histogram from (Figure 29A). During analysis, a gate was established to distinguish low-staining (loss of cytochrome c) from high staining (cytochrome c retained in mitochondria) cells, allowing a determination of the percentage of cells retaining mitochondria- associated cytochrome c. For example, in one representative experiment of SKBR-3, cytochrome c stain was lowest with both treatment: no treatment (83.9%), Th1 cytokines (83.8%), sunitinib

(23.8%), and both treatment combination (4%) (Figure 29A).

We also analyzed the flow cytometry data by overall fluorescent intensity (mean channel fluorescence). Composite data from at least three separate experiments represented in Figure

(28B). There was no evidence of suppression of mean channel fluorescence values between untreated cells and Th1 cytokine-treated cells. However, sunitinib treatment alone lead to strong suppression of mitochondrial-associated cytochrome c, and a trend toward even lower levels was observed when Th1 cytokines were added, but combined treatment did not achieve statistical significance over sunitinib alone. These studies did indicate, however, that additional apoptotic changes were occurring in the cells as a consequence of treatment.

PARP protein levels were lower in both treatment group.

To indirectly assess caspase 3 activity, we monitored via western blot analysis the disappearance of one of its proteolytic targets, PARP. Although single agents seemed to have little effect on PARP, the combination of sunitinib and Th1 cytokines appeared to display a trend toward loss of PARP protein (Figure 30A). However, when the bands from three separate blots were subjected to densitometry analysis and normalized to an actin control, the differences were not significant (Figure 30B).

Figure 22: Combination of Th1 cytokines and sunitinib enhance metabolic suppression of breast cancer cells over single treatments.

MDA-MB-231, MDA-MB-468, MCF-7, HCC-1419 and SKBR-3 cells were seeded at 5x103 cells per well and were left untreated (No Rx), treated with Th1 cytokines (5ng/mL), sunitinib

(10uM) or both sunitinib and Th1 cytokines for 72 hours. Alamar Blue dye was added to each well and optical density of the supernatant read spectrophometrically at 630nm.

Figure 23: Metabolic activity of breast cancer cell lines is maximally suppressed by dual

Th1 cytokines and sunitinib treatment.

MDA-MB-231, MDA-MB-468, MCF-7, HCC-1419 and SKBR-3 cells were seeded at 5x103 cells per well and were left untreated (No Rx), treated with Th1 cytokines (5ng/mL), sunitinib

(10uM) or both sunitinib and Th1 cytokines. After 72 hours of incubation, Alamar Blue dye was added to each well and incubated further. When a color change became apparent, optical density of the supernatants were read spectrophometrically at 630nm. Data represent summary analysis of at least three separate experiments per cell line. Statistical significance determined by one way

ANOVA: MDA-MB-231 [F(3,56)=242.8, p<0.001], MDA-MB-468 [F(3,24),=5.84, p<0.004],

MCF-7 [F(3,56)=146.3, p<0.001], HCC-1419 [F(3, 56)=95.2, p<0.001], and SKBR-3

[F(3,16)=5.94, p<0.006], followed by post-hoc Tukey. Significance was set at *** p<0.001.

Error bars show the standard error of the mean.

Figure 24: Microscopic examination treated cells with sunitinib plus Th1 cytokines reveals evidence of cell death.

MDA-MB-468, MDA-MB-231, MCF-7, SKBR-3, and HCC-1419 cells were seeded at 5x104 per mL and were left untreated (No Rx), treated with Th1 cytokines (5ng/mL), sunitinib (10uM) or both sunitinib and Th1 cytokines for 72 hours. Photomicrographs were taken using Olympus

CK2 microscope with 20X objective lens.

Figure 25: Histograms analysis of Trypan Blue staining reveals enhanced cell death with combined Th1 cytokines and sunitinib treatment.

MDA-MB-468, MDA-MB-231, SKBR-3, HCC-1419 and MCF-7 cells were seeded at 5x104 per mL and were left untreated (No Rx), treated with Th1 cytokines (5ng/mL), sunitinib (10uM) or both sunitinib and Th1 cytokines for 72 hours. Cells were harvested and stained with Trypan

Blue dye, after which the imparted fluorescent signal was assessed by flow cytometry using a

642nm excitation laser.

Figure 26: Combination of Th1 cytokines and sunitinib significantly increased breast cancer cell death over single treatments.

MDA-MB-231, MDA-MB-468, MCF-7, HCC-1419 and SKBR-3 cells were seeded at 5x104 per mL and were left untreated (No Rx), treated with Th1 cytokines (5ng/mL), sunitinib (10uM) or both sunitinib and Th1 cytokines for 72 hours. Cells were harvested and stained with Trypan

Blue dye, which was assessed by flow cytometry. Data represent summary analysis of mean channel fluorescence of treated, stained cells from at least three separate experiments per cell line. Statistical significance determine by one way ANOVA: MDA-MB-231 [F(3,56)=46.1, p<0.001], MDA-MB-468 [F(3,52),=33.1, p<0.001], MCF-7 [F(3,28)=91.5, p<0.001], HCC-1419

[F(3, 28)=19.1, p<0.001] and SKBR-3 [F(3,24)=22.4, p<0.001], followed by post-hoc Tukey test. Significance was set at *** p<0.001, **p<0.01 and *p<0.05. Error bars represent the standard error of the mean.

Figure 27: Combination of Th1 cytokines and sunitinib maximized markers of apoptosis in breast cancer cell lines.

Annexin V and propidium iodide (PI), and stain uptake was assessed by flow cytometry. Two- color dot plot analysis represents percentage of gated cells in each quadrant. Double staining with both Annexin V/PI represents late apoptotic events.

A. B.

HER-2 fluorescence intensity

Figure 28: Dual treatment significantly suppresses HER-2 expression over single treatments.

SKBR-3 cells were seeded at 5x104 per mL and were left untreated (No Rx), treated with Th1 cytokines (5ng/mL), sunitinib (10uM), or both sunitinib and Th1 cytokines for 72 hours. Cells were harvested and stained with APC-tag HER-2, which was assessed by flow cytometry using

642nm excitation laser. A. Histogram analysis form a single representative experiment for HER-

2 expression in SKBR-3 cell lines B. Composite data analysis of mean channel fluorescence for each cell line and treatment group derived from at least three separate experiments. Statistical significance determine by one way ANOVA: [F(3,20)=65.7, p<0.001] followed by Tukey post- hoc analysis. Significance was set at *** p<0.001, **p<0.01 and *p<0.05. Error bars represent the standard error of the mean.

A. B.

Fluorescence intensity Figure 29: Cytochrome c release in SKBR-3 cell lines demonstrated apoptosis.

SKBR-3 cells were seeded at 5x104 per mL and were left untreated (No Rx), treated with Th1 cytokines (5ng/mL), sunitinib (10uM) or both sunitinib and Th1 cytokines for 72 hours. Cells were harvested, permeabilized, stained with APC-tag cytochrome c, and probed with secondary antibody, which was assessed by flow cytometry using 642nm excitation laser. A. Histogram representations a single experiment of cytochrome c expression in SKBR-3 cell lines B. Analysis represent composite data of mean channel fluorescence for each cell line and treatment group derived from at least three separate experiments. Statistical significance determine by one way

ANOVA: [F(3,8)=158.9, p<0.001] followed by Tukey post-hoc analysis. Significance was set at

*** p<0.001 and **p<0.01. Error bars represent the standard error of the mean.

A. B.

Figure 30: Dual Th1 cytokine treatment plus sunitinib results in a trend toward lowered

PARP levels in SKBR-3 cells.

SKBR-3 cells were seeded at 3X106 per mL and were left untreated (No Rx), treated with Th1 cytokines (5ng/mL), sunitinib (10uM) or both sunitinib and Th1 cytokines for 72 hours.

Following 72 hours incubation, cells were harvested and extracted in RIPA buffer in the presence of protease and phosphatase inhibitors. Protein lysate (30ug/well) was separated on a 4-

15% SDS gel before transferring onto nitrocellulose membrane. A. Western blot was probed with anti-PARP (116 kDa) and anti-Actin (42 kDa) loading control and band visualized using chemiluminescence with Fuji LAS 3000 detection system. B. Quantification of PARP protein was performed using Image J analysis and normalized to loading control. Data represent at least three separate experiments. Error bars represent the standard error of the mean.

Chapter 4

Discussion

It has been shown that a Th1-polarizing, anti-HER-2 DC vaccine can be effective in 30% of time in earliest pre-invasive form of breast cancer (Czerniecki et al., 2007; Sharma et al.,

2012; Lowenfeld, Mick, et al., 2017). However, as the tumors grow and become invasive the tumor burden becomes too great for the immune system alone (Lowenfeld, Zaheer, et al., 2017).

Adding low toxicity targeted drugs may be one way to assist the immune system in its anti-tumor effects without the patient experiencing debilitating side effects often seen with traditional cancer treatments.

Combination of Th1 cytokines (IFN-γ and TNF-α) on cultured breast cancer cell lines had shown a similar effect as observed in vaccinated patients with cell death and suppression of expression of HER-family protooncogenes (Namjoshi et al., 2016). Furthermore, when anti- estrogen drugs were added in combination with Th1 cytokines to ERpos breast cancer cell lines, cell death increased (Lowenfeld, Zaheer, et al., 2017). When a short course of anti-estrogen drugs were subsequently tested in a clinical trial in combination with DC vaccination, it was found that the response rate of ERpos patients increased from 5% to 30% (Lowenfeld, Mick, et al., 2017). Based on the success of this approach, we tested in these studies additional small molecule inhibitors of targets associated with growth factor signaling or regulation of cell proliferation in combination with Th1 cytokines with a view toward enhancing breast cancer cell

death in vitro, thus providing pre-clinical data for future clinical trials aimed at further increasing vaccine response rates.

We therefore examined ibrance, which inhibits CDK4/6. During cell proliferation, transition of G1 to S phase requires phosphorylation of Rb protein, which is bound to E2F.

Binding of cyclin D1 to CDK4/6 activates the complex leading to Rb phosphorylation and freeing E2F leading to DNA replication (S-phase) (Dean et al., 2010). Since ibrance has been shown to inhibit the cell cycle pathway, and because Th1 cytokines are known to reduce surface expression of growth factor receptors such as HER-2 and HER-3 (thereby limiting growth factor signaling), we anticipated that the combinations of ibrance and Th1 cytokines would simultaneously attack these two proliferation-critical pathways and thereby work together to minimize cell growth and maximize cell death. Ibrance has been used in clinical trials for patients with estrogen receptor-positive breast cancer; this led us to examine whether Th1 cytokines plus ibrance would enhance breast cancer death in other, ERneg cell lines, and offering a possible rationale for expanding the indication for this drug when used in conjunction with vaccines.

Therefore, the main focus of our studies with ibrance was on estrogen receptor-negative cell lines. These included the HER-2pos Rbpos lines SKBR-3 and JIMT-1. We also included the

HER-2neg HER-3pos line MDA-MB-468, which is known to be Rbneg. We anticipated that SKBR-

3 and JIMT-1 would demonstrate additive effects when Th1 cytokines and ibrance were combined, but expected MDA-MB-468 to be unresponsive to this pairing, owing to its lack of

Rb expression (Robinson et al., 2013). We found that for the former two cell lines, the combination of Th1 cytokines plus ibrance maximized suppression of metabolic activity (Alamar

Blue assay; Figures 12 and 13), cell death (Trypan Blue assay; Figures 15 and 16), loss of

mitochondrial membrane potential (TMRE assay; Figure 17 and 18) and membrane changes associated with apoptosis (Annexin V/PI staining; Figure 19). Collectively, each of these assays confirmed our original hypothesis that Th1 cytokines plus a drug that inhibited a discrete aspect of cell cycle control would work cooperatively to enhance breast cancer cell death above that achievable by either agent alone.

Our results with MDA-MB-468 were at first surprising, however. We initially included this line because we expected it to react in contradistinction to the other two lines, owing to its lack of Rb protein (Robinson et al., 2013), which is one of the chief substrates of the CDK4/6- cyclin D complex, the activity of which is blocked by ibrance. Consistent with our original hypothesis, other studies have reported that MDA-MB-468 cell lines did not respond to ibrance, and were thus considered resistant (Fry et al., 2004; Finn et al., 2009; Dean et al., 2010). Our studies also confirmed that treatment with ibrance alone did not kill MDA-MB-468 cells; however, when ibrance was combined with Th1 cytokines, cell death was significantly enhanced

(p<0.05, Figure 16). This finding suggests that when Th1 cytokines are added, ibrance inhibition of CDK4/6 might be playing another role in the cell cycle. Some studies have reported that following CDK4/6 inhibition in Rb-deficient cells, there was observed a significant suppression of E2F (target gene) expression, which suggests potential involvement of the p107/p130 pocket protein to compensate Rb loss (Sage et al., 2000; Cobrinik, 2005; Rivadeneira et al., 2010).

Perhaps p107/p130 might be involved in MDA-MB-468 cell lines leading to enhanced cell death, however; more additional experiments need to be performed to test this hypothesis. It should be noted that these findings with ibrance are not the first time that exposure to Th1 cytokines was observed by us to sensitize previously drug-resistant cell lines to drug-induced cell death.

Lapatinib is a small molecule inhibitor of both HER-1 (EGFR) and HER-2, and exposure of

lapatinib-resistant cell lines to Th1 cytokines likewise reverses drug resistance (Showalter, manuscript in preparation). Acquired resistance to small molecule inhibitor drugs is a major problem in cancer therapy (Van Der Kuip et al., 2005; D’amato et al., 2015), and it is fascinating to speculate that the exposure to Th1 cytokines may exploit either a common mechanism, or multiple discrete mechanisms to circumvent resistance to multiple drugs. This would make DC vaccination a preferred adjunct to drug treatment to avoid the problem of resistance.

Since ibrance treatment inhibits cyclin D and CDK4/6 complex, which leads to hypophosphorylation of Rb, we sought to determine levels of both total Rb protein as well as its phosphorylated form by western blot analysis. Since ibrance and Th1 cytokines worked together to maximize cell death, we considered it possible that the cytokines might actually enhance the suppression of Rb phosphorylation by ibrance. We instead discovered that total Rb was significantly reduced for cells treated with both Th1 cytokines and ibrance, compared with single treatment (p<0.05, Figure 20B). However, when we examined phosphorylated Rb in SKBR-3 cell lines using phospho-specific antibodies, we observed the major band at 160 kDa rather than the expected 110 kDa (Figure 21). After making this unexpected observation, we tried a second anti-phospho-Rb antibody and saw reactivity with the same high molecular weight band. We are currently at a loss to explain this discrepancy. If the observed high molecular weight band is due to an adventitious cross-reaction, it is indeed unusual that two different antibodies would have the same cross-reactivity. In addition, it is difficult to explain why we do not also see a strong band at the expected molecular mass, when the total Rb antibody generates a strong signal at 110 kDa. It is also remotely possible that the phosphorylated form of Rb is binding to another protein, giving the complex a higher apparent molecular mass. But this too seems unlikely because the protein extracts are boiled in a loading buffer containing SDS as well as beta-

mercaptoethanol, the combination of which should disrupt virtually all protein-protein interactions. The identity of the high molecular weight band will likely not be resolved without protein sequencing data. However, if this higher molecular protein is in fact phospho-Rb, then its levels are clearly dropping with combined treatment, just as the 110 kDa band identified with anti-total Rb protein antibody.

Th1 cytokines are known to suppress the expression of a number of growth factor receptors such as HER-2 and HER-3 (Namjoshi et al., 2016). Similarly, sunitinib inhibits the activity of multiple RTKs. Therefore we anticipated that the combination of Th1 cytokines with sunitinib would work cooperatively to rob cancer cells of important growth and survival signals, thereby enhancing breast cancer cells death. For these studies, we examined hormone receptor positive (MCF-7), HER-2 positive (SKBR-3 and HCC-1419) and triple negative (MDA-MB-468 and MDA-MB-231) cell lines. In all cell lines the combination of sunitinib and Th1 cytokines effectively suppressed metabolic activity (Alamar Blue assay, Figure 22 and 23), induced cell death (Trypan Blue assay, Figure 25 and 26), and led to apoptosis (Annexin V/PI assay, Figure

27). We further examined the expression level of growth factor receptor HER-2, in SKBR-3 cell lines and discovered that the combination of both sunitinib and Th1 cytokines led to significant suppression (p<0.05, Figure 28) of the growth factor receptor.

Many of the standard assays we employ in our laboratory to assess cell viability, apoptotic cell death and expression of surface markers are fluorescence-based. We encountered an unexpected difficulty during these studies arising from the fact that the drug sunitinib possesses fluorescent properties strong enough to interfere with many of our assays that rely upon excitation with the 488nm laser and detection in certain channels such as those used for the fluorochrome FITC and TMRE dye. In some cases, we were able to work around this limitation

by substituting fluorochromes. For example, for the sunitinib studies, we substituted FITC- labeled Annexin V with an APC conjugate that was excited by the 642nm laser and detected in a channel far removed from the emission spectrum of sunitinib. For TMRE, however, there was no substitute, and we thus could not perform mitochondrial membrane polarization studies when sunitinib was used.

We did, however, examine a different aspect of mitochondrial function associated with apoptosis. One of the hallmarks for apoptosis is the loss of cytochrome c from the mitochondria

(Christensen et al., 2013). We measured cytochrome c levels in SKBR-3 cell lines by flow cytometry using the appropriate fluorochrome-labeled antibodies so as to avoid the problem of interference by the drug. As expected, cytochrome c was lowered in cells treated with both Th1 cytokines and sunitinib compared to treatment alone (Figure 29). We were surprised to observe that Th1 cytokine treatment alone did not result in a decrease in cytochrome c levels, given the fact that this treatment showed evidence of moderate cell death by all other criteria examined in these studies. In addition, TNF-α, one of the Th1 cytokines used, binds directly to extrinsic pathway receptors and should induce apoptosis (Balachandran et al., 1998). TNF-α binds to TNF superfamily receptor and adaptor proteins such as TRADD (tumor necrosis factor receptor type

1-associated death domain protein) and FADD (Fas-associated protein with death domain) binds to the receptor (Yuan et al., 2012). This interaction leads to formation of death inducing signaling complex that cleaves pro-caspase 8 leading to activation of Bid (pro-apoptotic signal), which is known to causes the release of cytochrome c. However, despite the weakness of Th1 cytokines alone for inducing cytochrome c loss, when combined with sunitinib, cytochrome c loss as well as other indicators of apoptosis were increased.

In these studies with sunitinib, we also introduced an additional assay to evaluate cellular apoptosis, which was observing levels of poly (ADP-ribose) polymerase-1 (PARP-1) by western blot. PARP is a substrate of executioner caspase 3, so loss of intact PARP is an indicator of caspase 3 activity, and thus apoptosis. We observed a downward trend in PARP levels with dual

Th1 cytokine/sunitinib treatment, but the differences observed in three separate experiments did not reach the level of significance (Figure 30A). Given the relatively strong trend, additional repetitions of this experiment are likely to achieve statistical significance.

Because each malignant tumor is unique, with its own set of mutations and dysregulated genes, most cancer cell lines behave differently with treatment; however, we were surprised that sunitinib worked very well in an unusually broad set of phenotypically diverse cell lines including hormone receptor positive, HER-2 positive, and triple negative. These finding suggests that sunitinib could be potent drug to be combined in clinical trials with Th1 cytokines vaccination to enhance cancer cell death.

Although these in vitro studies are highly encouraging, using such drugs with Th1- polarizing vaccination has potential limitations that must be considered, since there are a number of points of overlap between the immune system and the drug targets. Studies have demonstrated a potential link between immune system and oncogenic signaling to upregulate the MAPK pathway and impair IL-12 production by dendritic cells in melanoma (Sumimoto et al., 2006).

Small molecule inhibitors have targeted oncogenic pathway and downstream signaling. For example, sunitinib treatment for patients with renal cell cancer demonstrated an increase in frequency of T cells producing IFN-γ (i.e. Th1 cells) and a decrease T cells producing IL-4 (i.e.

Th2 cells), which reverses the Th1/Th2 ratio. This is potent for generating an effective antitumor immune response (Finke et al., 2008). In these renal cancer cells, T regulatory cells

(which favor tumor growth) are elevated, which was shown to be reduced with sunitinib treatment (Finke et al., 2008). Also, renal cell carcinoma patients treated with sunitinib demonstrated a reduction of myeloid-derived suppressor cells (Ko et al., 2009), which are also considered to promote tumor survival and growth. However, in an in vitro study of renal cell carcinoma, sunitinib led to a decrease in IL-12 production by dendritic cells matured with TNF-

α, IFN-γ, and poly I:C (Alfaro et al., 2009). Patients with metastatic clear cell renal cancer that were treated with sunitinib showed depressed level of CD3pos and CD4pos T cells (Powles et al.,

2011). It would seem that the impact of sunitinib on the immune system is both immunostimulatory as well as immunosuppressive, depending on the cell type. These conflicting results of sunitinib having anti-tumor immunity is still unclear (Jaini et al., 2014). However, when sunitinib was administered in combination with targeted immunotherapeutic vaccination

(in murine model), it affected the priming of the antigen presenting cells (decrease in CD11bpos,

CD11cpos) and led to vaccination failure (Jaini et al., 2014). On the other hand, when sunitinib was administered after the priming phase of vaccination, there was an increase in vaccination mediated immune response (Jaini et al., 2014). These latter studies make clear that timing is an important issue. The drugs and the vaccine may work best if given sequentially rather than concurrently to avoid most of the problems of drug-induced immunosuppression. More studies need to be performed for breast cancer cell lines to examine whether combining sunitinib with vaccination would lead to immunostimulation or immunosuppression.

In the case of CDK4/6 inhibitors, fewer studies are available regarding its impact on the immune system. It one study that looked at the combination of checkpoint inhibitor drugs with small molecule inhibitors of CDK4/6, it was found that inhibiting CDK4/6 for a brief period of time leads to a significant enhancement of T-cell activation. This appears to be due to

suppression of the NFAT (nuclear factor of activated T cells) family proteins, which are important for regulating T-cell function. Nonetheless, CDK4/6 inhibition does decrease overall

T-cell proliferation. Even so, it appeared to also increase tumor infiltration and activation of effector T cells (Deng et al., 2018). So as in the case with sunitinib, available data with CDK4/6 inhibitors indicate that the drug may be a double-edged sword. More studies are needed to determine optimized treatment schedules for both drugs when they are used as adjuvants to vaccination.

In summary, these discoveries collectively demonstrate that small molecule inhibitor drugs, such as ibrance and sunitinib, work in combination with Th1 cytokines to maximize apoptotic cell death in different types of breast cancer cell lines. These preclinical findings provide the rationale for their further development as adjuvants to be included with anti-HER-2

DC1 vaccination therapy to increase its response and to enhance tumor death. Vaccination efficacy might be improved through these type of drugs that interfere with cell cycle check points and signaling pathways.

Chapter 5

Future Direction

Here we show that Th1 cytokines combined with either ibrance or sunitinib leads to breast cancer death via apoptosis. Yet, there are several ideas that are not explored in this thesis.

For example, even though treatment using Th1 cytokines (IFN-γ and TNF-α) for breast cancer results in high levels of cellular apoptosis, suppression of HER-2 growth factor, and mimics in vivo effects of vaccination (Namjoshi et al., 2016), we still have not ascertained the complete mechanism of action for the cytokines alone.

To examine the mechanism of these Th1 cytokines involved in apoptosis, the IFN-γ and

TNF-α pathway needs further investigation. IFN-γ has been shown by others to upregulate expression of p21 and p27, which leads to cell cycle arrest. Cells with Th1 cytokine treatment can be examined for p21 and p27 expression via western blot. Similarly, TNF-α is one of the ligands involved in the activation of extrinsic pathway for apoptosis. When TNF-α binds, it activates the DISC formation that is only formed by the activation of extrinsic pathway, which needs to be further investigated. This could be tested by performing co- immunoprecipitation/western blot studies to verify the formation of the disc complex.

Even though we demonstrated that ibrance leads to cell death via a probable apoptotic mechanism (Annexin V/PI assay), further experiments need to be performed to establish a complete apoptosis pathway. One such line of investigation would be to examine initiator caspase 8 (activate only through extrinsic apoptosis pathway) activity and perform a western blot to analyze whether both pro- and cleaved forms of caspase 8 can be detected. Potentially, Th1

cytokines and small molecule inhibitors may induce apoptosis by working on the extrinsic pathway. It is important to examine caspase 8 activity which could help distinguished between intrinsic and extrinsic pathway of apoptosis. Cleaved caspase 8 eventually leads to the release of cytochrome c from the mitochondria. Cytochrome c levels can be measured using flow cytometry. Another downstream effect of cleaved caspase 8 is activating executioner caspase 3.

Western blot analysis can be used to examine if the caspase 3 is cleaved. The experiments that we performed for this thesis only examined overall PARP activity. To examine if the cleaved caspase 3 leads to cleavage of PARP, western blot can be performed to measure cleaved PARP levels. Performing these experiments will help us design the actual mechanistic pathway of how the small molecule inhibitors and Th1 cytokines are cooperating to enhance apoptosis.

Several studies have shown that ibrance treatment leads to reduce phosphorylated Rb; however, we could only detect Rb in higher molecular weights. To confirm the higher molecular weight of Rb, western blot analysis needs to be followed by sequencing. This sequencing method will allow us to verify whether the observed high molecular weight protein is actually Rb or some other protein that cross-reacts with two individual anti-phospho-Rb antibodies.

Surprisingly in MDA-MB-468 cell lines that are Rb negative, cell death was increased with combination of Th1 cytokines and ibrance treatments, which could be due to Rb family pocket protein. These Rb family pocket proteins (p107/p130) need to be examined by performing western blot to observe if they are involved in the enhanced cell death with the combination treatment. If they are present, perhaps siRNA studies could knock down the expression of these proteins and modulate sensitivity of cells to drugs plus Th1 cytokines. These data will help us understand the mechanistic pathway.

For sunitinib studies, we restricted our observations to HER-2 expression; however, sunitinib inhibits other receptor tyrosine kinases receptor such as VEGF, PDGF, and Kit. It would be of interest to determine whether Th1 cytokines also inhibit the expression of these other RTKs, and whether addition of the drug enhances this suppression. Finally, studies should be performed in a mouse model to obtain preclinical data that could potentially support a clinical trial that adds ibrance or sunitinib to DC-based Th1-polarizing vaccines directed against HER-2 overexpressing tumors.

References

AL-NASIRY, S. et al. The use of Alamar Blue assay for quantitative analysis of viability, migration and invasion of choriocarcinoma cells. Human reproduction, v. 22, n. 5, p. 1304-1309, 2007. ISSN 1460-2350.

ALFARO, C. et al. Influence of bevacizumab, sunitinib and sorafenib as single agents or in combination on the inhibitory effects of VEGF on human dendritic cell differentiation from monocytes. British Journal of Cancer, v. 100, n. 7, p. 1111, 2009. ISSN 1532- 1827.

ANDERSON, W. F. et al. How many etiological subtypes of breast cancer: two, three, four, or more? Journal of the National Cancer Institute, v. 106, n. 8, p. dju165, 2014. ISSN 1460-2105.

APPELLA, E.; ANDERSON, C. W. Post‐translational modifications and activation of p53 by genotoxic stresses. The FEBS Journal, v. 268, n. 10, p. 2764-2772, 2001. ISSN 1432- 1033.

ASHKENAZI, A. Targeting death and decoy receptors of the tumour-necrosis factor superfamily. Nature Reviews Cancer, v. 2, n. 6, p. 420, 2002. ISSN 1474-1768.

AVELAR-FREITAS, B. A. et al. Trypan blue exclusion assay by flow cytometry. Brazilian journal of medical and biological research, v. 47, n. 4, p. 307-315, 2014. ISSN 0100- 879X.

BACH, E. A.; AGUET, M.; SCHREIBER, R. D. The IFNγ receptor: a paradigm for cytokine receptor signaling. Annual review of immunology, v. 15, n. 1, p. 563-591, 1997. ISSN 0732-0582.

BACHELOT, T. et al. Sunitinib in combination with trastuzumab for the treatment of advanced breast cancer: activity and safety results from a phase II study. BMC Cancer, v. 14, p. 166-166, 03/0712/09/received02/20/accepted 2014. ISSN 1471-2407. Available at: < http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3995914/ >.

BALACHANDRAN, S. et al. Activation of the dsRNA‐dependent protein kinase, PKR, induces apoptosis through FADD‐mediated death signaling. The EMBO journal, v. 17, n. 23, p. 6888-6902, 1998. ISSN 0261-4189.

BALASUBRAMANIAN, K.; MIRNIKJOO, B.; SCHROIT, A. J. Regulated externalization of phosphatidylserine at the cell surface implications for apoptosis. Journal of Biological Chemistry, v. 282, n. 25, p. 18357-18364, 2007. ISSN 0021-9258.

BERGH, J. et al. First-Line Treatment of Advanced Breast Cancer With Sunitinib in Combination With Docetaxel Versus Docetaxel Alone: Results of a Prospective, Randomized Phase III Study. Journal of Clinical Oncology, v. 30, n. 9, p. 921-929, 2012/03/20 2012. ISSN 0732-183X. Available at: < https://doi.org/10.1200/JCO.2011.35.7376 >. Accessed on: 2018/01/29.

BOATRIGHT, K. M.; SALVESEN, G. S. Mechanisms of caspase activation. Current Opinion in Cell Biology, v. 15, n. 6, p. 725-731, 2003/12/01/ 2003. ISSN 0955-0674. Available at: < http://www.sciencedirect.com/science/article/pii/S0955067403001364 >.

BOULARES, A. H. et al. Role of poly (ADP-ribose) polymerase (PARP) cleavage in apoptosis Caspase 3-resistant PARP mutant increases rates of apoptosis in transfected cells. Journal of Biological Chemistry, v. 274, n. 33, p. 22932-22940, 1999. ISSN 0021- 9258.

BURRIS, H. A. Dual kinase inhibition in the treatment of breast cancer: initial experience with the EGFR/ErbB-2 inhibitor lapatinib. The oncologist, v. 9, n. Supplement 3, p. 10-15, 2004. ISSN 1083-7159.

CADOO, K. A.; GUCALP, A.; TRAINA, T. A. Palbociclib: an evidence-based review of its potential in the treatment of breast cancer. Breast Cancer: Targets and Therapy, v. 6, p. 123, 2014.

CALDON, C. E. et al. Cell cycle control in breast cancer cells. Journal of cellular biochemistry, v. 97, n. 2, p. 261-274, 2006. ISSN 1097-4644.

CHIN, Y. E. et al. Cell growth arrest and induction of cyclin-dependent kinase inhibitor p21WAF1/CIP1 mediated by STAT1. Science, v. 272, n. 5262, p. 719-722, 1996. ISSN 0036-8075.

CHRISTENSEN, M. E. et al. Flow cytometry based assays for the measurement of apoptosis- associated mitochondrial membrane depolarisation and cytochrome c release. Methods, v. 61, n. 2, p. 138-145, 2013/06/01/ 2013. ISSN 1046-2023. Available at: < http://www.sciencedirect.com/science/article/pii/S1046202313000893 >.

COBRINIK, D. Pocket proteins and cell cycle control. Oncogene, v. 24, n. 17, p. 2796, 2005. ISSN 1476-5594.

CROWN, J. P. et al. Phase III Trial of Sunitinib in Combination With Capecitabine Versus Capecitabine Monotherapy for the Treatment of Patients With Pretreated Metastatic Breast Cancer. Journal of Clinical Oncology, v. 31, n. 23, p. 2870-2878, 2013/08/10 2013. ISSN 0732-183X. Available at: < https://doi.org/10.1200/JCO.2012.43.3391 >. Accessed on: 2018/01/29.

CZERNIECKI, B. J. et al. Targeting HER-2/neu in Early Breast Cancer Development Using Dendritic Cells with Staged Interleukin-12 Burst Secretion. Cancer Research, v. 67, n. 4, p. 1842, 2007. Available at: < http://cancerres.aacrjournals.org/content/67/4/1842.abstract >.

D'AMOURS, D. et al. Poly(ADP-ribosyl)ation reactions in the regulation of nuclear functions. Biochemical Journal, v. 342, n. Pt 2, p. 249-268, 1999. ISSN 0264-60211470-8728. Available at: < http://www.ncbi.nlm.nih.gov/pmc/articles/PMC1220459/ >.

DARLAND-RANSOM, M. et al. Role of C. elegans TAT-1 Protein in Maintaining Plasma Membrane Phosphatidylserine Asymmetry. Science, v. 320, n. 5875, p. 528, 2008. Available at: < http://science.sciencemag.org/content/320/5875/528.abstract >.

DARNELL, J. E.; KERR, I. M.; STARK, G. R. Jak-STAT pathways and transcriptional activation in response to IFNs and other extracellular signaling proteins. Science, v. 264, n. 5164, p. 1415-1421, 1994. ISSN 0036-8075.

DE ANDREA, M. et al. The interferon system: an overview. European Journal of Paediatric Neurology, v. 6, p. A41-A46, 2002. ISSN 1090-3798.

DEAN, J. L. et al. Therapeutic CDK4/6 inhibition in breast cancer: key mechanisms of response and failure. Oncogene, v. 29, n. 28, p. 4018-4032, 2010. ISSN 0950-9232.

DENG, J. et al. CDK4/6 Inhibition Augments Antitumor Immunity by Enhancing T-cell Activation. Cancer discovery, v. 8, n. 2, p. 216-233, 2018. ISSN 2159-8274.

DESANTIS, C. E. et al. Breast cancer statistics, 2015: Convergence of incidence rates between black and white women. CA: a cancer journal for clinicians, v. 66, n. 1, p. 31-42, 2016. ISSN 1542-4863.

DHILLON, S. Palbociclib: first global approval. Drugs, v. 75, n. 5, p. 543-551 %@ 0012-6667, 2015.

D’AMATO, V. et al. Mechanisms of lapatinib resistance in HER2-driven breast cancer. Cancer treatment reviews, v. 41, n. 10, p. 877-883, 2015. ISSN 0305-7372.

D’AMOURS, D. et al. Gain-of-function of poly (ADP-ribose) polymerase-1 upon cleavage by apoptotic proteases: implications for apoptosis. Journal of cell science, v. 114, n. 20, p. 3771-3778, 2001. ISSN 0021-9533.

EKHOLM, S. V.; REED, S. I. Regulation of G1 cyclin-dependent kinases in the mammalian cell cycle. Current opinion in cell biology, v. 12, n. 6, p. 676-684, 2000. ISSN 0955-0674.

FIELDS, R. D.; LANCASTER, M. V. Dual-attribute continuous monitoring of cell proliferation/cytotoxicity. American biotechnology laboratory, v. 11, n. 4, p. 48-50, 1993. ISSN 0749-3223.

FINKE, J. H. et al. Sunitinib reverses type-1 immune suppression and decreases T-regulatory cells in renal cell carcinoma patients. Clinical Cancer Research, v. 14, n. 20, p. 6674- 6682, 2008. ISSN 1078-0432.

FINKLE, D. et al. HER2-Targeted Therapy Reduces Incidence and Progression of Midlife Mammary Tumors in Female Murine Mammary Tumor Virus huHER2-Transgenic Mice. Clinical Cancer Research, v. 10, n. 7, p. 2499, 2004. Available at: < http://clincancerres.aacrjournals.org/content/10/7/2499.abstract >.

FINN, R. S. et al. The cyclin-dependent kinase 4/6 inhibitor palbociclib in combination with letrozole versus letrozole alone as first-line treatment of oestrogen receptor-positive, HER2-negative, advanced breast cancer (PALOMA-1/TRIO-18): a randomised phase 2 study. The lancet oncology, v. 16, n. 1, p. 25-35, 2015. ISSN 1470-2045.

FINN, R. S.; DERING, J.; CONKLIN, D., et al. PD 0332991, a selective cyclin D kinase 4/6 inhibitor, preferentially inhibits proliferation of luminal estrogen receptor-positive human breast cancer cell lines in vitro. Breast Cancer Research, v. 11, n. 5, p. R77, 2009. ISSN 1465-542X.

FLAISHON, L. et al. Autocrine secretion of interferon γ negatively regulates homing of immature B cells. Journal of Experimental Medicine, v. 192, n. 9, p. 1381-1388, 2000. ISSN 0022-1007.

FRUCHT, D. M. et al. IFN-γ production by antigen-presenting cells: mechanisms emerge. Trends in immunology, v. 22, n. 10, p. 556-560, 2001. ISSN 1471-4906.

FRY, D. W. et al. Specific inhibition of cyclin-dependent kinase 4/6 by PD 0332991 and associated antitumor activity in human tumor xenografts. Molecular cancer therapeutics, v. 3, n. 11, p. 1427-1438, 2004. ISSN 1535-7163.

GESSANI, S.; BELARDELLI, F. IFN-γ expression in macrophages and its possible biological significance. Cytokine & growth factor reviews, v. 9, n. 2, p. 117-123, 1998. ISSN 1359-6101.

GIACINTI, C.; GIORDANO, A. RB and cell cycle progression. Oncogene, v. 25, n. 38, p. 5220, 2006. ISSN 1476-5594.

GOOCH, J. L.; HERRERA, R. E.; YEE, D. The role of p21 in interferon gamma-mediated growth inhibition of human breast cancer cells. Cell growth & differentiation: the molecular biology journal of the American Association for Cancer Research, v. 11, n. 6, p. 335-342, 2000. ISSN 1044-9523.

GUDKOV, A. V.; KOMAROVA, E. A. The role of p53 in determining sensitivity to radiotherapy. Nature Reviews Cancer, v. 3, n. 2, p. 117, 2003. ISSN 1474-1768.

HARARI, D.; YARDEN, Y. Molecular mechanisms underlying ErbB2/HER2 action in breast cancer. Oncogene, v. 19, n. 53, p. 6102, 2000. ISSN 0950-9232.

HARRIS, S. L.; LEVINE, A. J. The p53 pathway: positive and negative feedback loops. Oncogene, v. 24, n. 17, p. 2899, 2005. ISSN 1476-5594.

IQBAL, N.; IQBAL, N. Human Epidermal Growth Factor Receptor 2 (HER2) in Cancers: Overexpression and Therapeutic Implications. Molecular Biology International, v. 2014, p. 852748, 09/07 05/28/received 08/18/revised 08/19/accepted 2014. ISSN 2090- 2182 2090-2190. Available at: .

JAINI, R. et al. Combination of sunitinib with anti‐tumor vaccination inhibits T cell priming and requires careful scheduling to achieve productive immunotherapy. International journal of cancer, v. 134, n. 7, p. 1695-1705, 2014. ISSN 1097-0215.

KAUFMANN, S. H. et al. Specific proteolytic cleavage of poly (ADP-ribose) polymerase: an early marker of chemotherapy-induced apoptosis. Cancer research, v. 53, n. 17, p. 3976- 3985, 1993. ISSN 0008-5472.

KLUCK, R. M.; BOSSY-WETZEL, E., et al. The release of cytochrome c from mitochondria: a primary site for Bcl-2 regulation of apoptosis. Science, v. 275, n. 5303, p. 1132-1136, 1997. ISSN 0036-8075.

KLUCK, R. M.; MARTIN S. J., et al. Cytochrome c activation of CPP32‐like proteolysis plays a critical role in a Xenopus cell‐free apoptosis system. The EMBO journal, v. 16, n. 15, p. 4639-4649, 1997. ISSN 0261-4189.

KNUDSEN, E. S.; KNUDSEN, K. E. Tailoring to RB: tumour suppressor status and therapeutic response. Nature reviews. Cancer, v. 8, n. 9, p. 714-724, 2008. ISSN 1474-175X1474- 1768. Available at: < http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2914856/ >.

KO, J. S. et al. Sunitinib mediates reversal of myeloid-derived suppressor cell accumulation in renal cell carcinoma patients. Clinical cancer research, v. 15, n. 6, p. 2148-2157, 2009. ISSN 1078-0432.

KOOPMAN, G. et al. Annexin V for flow cytometric detection of phosphatidylserine expression on B cells undergoing apoptosis. Blood, v. 84, n. 5, p. 1415-1420, 1994. ISSN 0006- 4971.

KOSKI, G. K. et al. A novel dendritic cell-based immunization approach for the induction of durable Th1-polarized anti-HER-2/neu responses in women with early breast cancer. Journal of immunotherapy (Hagerstown, Md. : 1997), v. 35, n. 1, p. 54-65, 2012. ISSN 1524-9557 1537-4513. Available at: < http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3241864/ >.

LEONARD, J. P. et al. Selective CDK4/6 inhibition with tumor responses by PD0332991 in patients with mantle cell lymphoma. Blood, v. 119, n. 20, p. 4597-4607, 2012. ISSN 0006-4971.

LEVENTIS, P. A.; GRINSTEIN, S. The distribution and function of phosphatidylserine in cellular membranes. Annual review of biophysics, v. 39, p. 407-427, 2010. ISSN 1936- 122X.

LI, J.; YUAN, J. Caspases in apoptosis and beyond. Oncogene, v. 27, n. 48, p. 6194, 2008. ISSN 1476-5594.

LOGAN, J. E. et al. PD-0332991, a potent and selective inhibitor of cyclin-dependent kinase 4/6, demonstrates inhibition of proliferation in renal cell carcinoma at nanomolar concentrations and molecular markers predict for sensitivity. Anticancer research, v. 33, n. 8, p. 2997-3004, 2013. ISSN 0250-7005.

LOUIS, K. S.; SIEGEL, A. C. Cell viability analysis using trypan blue: manual and automated methods. In: (Ed.). Mammalian Cell Viability: Springer, 2011. p.7-12.

LOWENFELD, L.; MICK, R., et al. Dendritic cell vaccination enhances immune responses and induces regression of HER2pos DCIS independent of route: results of randomized selection design trial. Clinical Cancer Research, v. 23, n. 12, p. 2961-2971, 2017. ISSN 1078-0432.

LOWENFELD, L.; ZAHEER, S.; OECHSLE, C., et al. Addition of anti-estrogen therapy to Anti-HER2 dendritic cell vaccination improves regional nodal immune response and pathologic complete response rate in patients with ERpos/HER2pos early breast cancer. Oncoimmunology, v. 6, n. 9, p. e1207032, 2017. ISSN 2162-402X.

LUTFUL KABIR, F.; ALVAREZ, C.; CURTIS BIRD, R. Canine Mammary Carcinomas: A Comparative Analysis of Altered Gene Expression. 2016. 1-21.

MAHMOUD, S. M. A. et al. Tumor-Infiltrating CD8+ Lymphocytes Predict Clinical Outcome in Breast Cancer. Journal of Clinical Oncology, v. 29, n. 15, p. 1949-1955, 2011/05/20 2011. ISSN 0732-183X. Available at: < https://doi.org/10.1200/JCO.2010.30.5037 >. Accessed on: 2018/03/10.

MANDAL, D. et al. Caspase 3 regulates phosphatidylserine externalization and phagocytosis of oxidatively stressed erythrocytes. FEBS letters, v. 513, n. 2-3, p. 184-188, 2002. ISSN 1873-3468.

MANDLEKAR, S.; KONG, A. N. Mechanisms of tamoxifen-induced apoptosis. Apoptosis, v. 6, n. 6, p. 469-477, 2001. ISSN 1360-8185.

MENDEL, D. B. et al. In vivo antitumor activity of SU11248, a novel tyrosine kinase inhibitor targeting vascular endothelial growth factor and platelet-derived growth factor receptors: determination of a pharmacokinetic/pharmacodynamic relationship. Clinical Cancer Research, v. 9, n. 1, p. 327-337, 2003. ISSN 1078-0432.

MICHAEL, D.; OREN, M. The p53–Mdm2 module and the ubiquitin system. 1: Elsevier, 2003. p.49-58.

MILLER, W. R. Aromatase inhibitors: mechanism of action and role in the treatment of breast cancer. Elsevier, 2003. p.3-11.

MOHAMMADIZADEH, F. et al. Role of cyclin D1 in breast carcinoma. Journal of Research in Medical Sciences : The Official Journal of Isfahan University of Medical Sciences, v. 18, n. 12, p. 1021-1025, 02/28/received 04/29/revised 08/28/accepted 2013. ISSN 1735-1995 1735-7136. Available at: .

MORALES, J. C. et al. Review of Poly (ADP-ribose) Polymerase (PARP) Mechanisms of Action and Rationale for Targeting in Cancer and Other Diseases. Critical reviews in eukaryotic gene expression, v. 24, n. 1, p. 15-28, 2014. ISSN 1045-4403. Available at: < http://www.ncbi.nlm.nih.gov/pmc/articles/PMC4806654/ >.

MORIMOTO, A. M. et al. Gene expression profiling of human colon xenograft tumors following treatment with SU11248, a multitargeted tyrosine kinase inhibitor. Oncogene, v. 23, n. 8, p. 1618, 2004. ISSN 1476-5594.

NAGATA, S.; HANAYAMA, R.; KAWANE, K. Autoimmunity and the clearance of dead cells. Cell, v. 140, n. 5, p. 619-630, 2010. ISSN 0092-8674.

NAMJOSHI, P. et al. T-helper 1-type cytokines induce apoptosis and loss of HER-family oncodriver expression in murine and human breast cancer cells. Oncotarget, 2016. ISSN 1949-2553.

NARITA, M. et al. Bax interacts with the permeability transition pore to induce permeability transition and cytochrome c release in isolated mitochondria. Proceedings of the National Academy of Sciences, v. 95, n. 25, p. 14681-14686, 1998. ISSN 0027-8424.

NEVINS, J. R. The Rb/E2F pathway and cancer. Human molecular genetics, v. 10, n. 7, p. 699-703, 2001. ISSN 0964-6906.

OSBORNE, C. K. et al. Crosstalk between estrogen receptor and growth factor receptor pathways as a cause for endocrine therapy resistance in breast cancer. Clinical cancer research, v. 11, n. 2, p. 865s-870s, 2005. ISSN 1078-0432.

PAN, Q. et al. CDK4/6 Inhibitors in Cancer Therapy: A Novel Treatement Strategy for Bladder Cancer. Bladder Cancer, v. 3, n. 2, p. 79-88, 2017. ISSN 2352-3727.

PAUL, M. K.; MUKHOPADHYAY, A. K. Tyrosine kinase–role and significance in cancer. International journal of medical sciences, v. 1, n. 2, p. 101, 2004.

PEOPLES, G. E. et al. Clinical Trial Results of a HER2/neu (E75) Vaccine to Prevent Recurrence in High-Risk Breast Cancer Patients. Journal of Clinical Oncology, v. 23, n. 30, p. 7536-7545, 2005/10/20 2005. ISSN 0732-183X. Available at: < https://doi.org/10.1200/JCO.2005.03.047 >. Accessed on: 2018/03/17.

PESTKA, S.; KRAUSE, C. D.; WALTER, M. R. Interferons, interferon‐like cytokines, and their receptors. Immunological reviews, v. 202, n. 1, p. 8-32, 2004. ISSN 1600-065X.

PEYRESSATRE, M. et al. Targeting cyclin-dependent kinases in human cancers: from small molecules to peptide inhibitors. Cancers, v. 7, n. 1, p. 179-237, 2015.

POWLES, T. et al. The effect of sunitinib on immune subsets in metastatic clear cell renal cancer. Urologia internationalis, v. 86, n. 1, p. 53-59, 2011. ISSN 0042-1138.

RICCARDI, C.; NICOLETTI, I. Analysis of apoptosis by propidium iodide staining and flow cytometry. Nature protocols, v. 1, n. 3, p. 1458, 2006. ISSN 1750-2799.

RIVADENEIRA, D. B. et al. Proliferative suppression by CDK4/6 inhibition: complex function of the retinoblastoma pathway in liver tissue and hepatoma cells. Gastroenterology, v. 138, n. 5, p. 1920-1930, 2010. ISSN 0016-5085.

ROBINSON, T. J. W. et al. RB1 status in triple negative breast cancer cells dictates response to radiation treatment and selective therapeutic drugs. PloS one, v. 8, n. 11, p. e78641, 2013. ISSN 1932-6203.

SAGE, J. et al. Targeted disruption of the three Rb-related genes leads to loss of G1 control and immortalization. Genes & development, v. 14, n. 23, p. 3037-3050, 2000. ISSN 0890- 9369.

SANGWAN, V.; PARK, M. Receptor tyrosine kinases: role in cancer progression. Current oncology, v. 13, n. 5, p. 191, 2006.

SARASTE, A.; PULKKI, K. Morphologic and biochemical hallmarks of apoptosis. Cardiovascular research, v. 45, n. 3, p. 528-537, 2000. ISSN 1755-3245.

SCHRODER, K. et al. Interferon‐γ: an overview of signals, mechanisms and functions. Journal of leukocyte biology, v. 75, n. 2, p. 163-189, 2004. ISSN 1938-3673.

SCHWARTZ, G. K. et al. Phase I study of PD 0332991, a cyclin-dependent kinase inhibitor, administered in 3-week cycles (Schedule 2/1). British journal of cancer, v. 104, n. 12, p. 1862-1868, 2011. ISSN 0007-0920.

SCOVASSI, A. I.; POIRIER, G. G. Poly (ADP-ribosylation) and apoptosis. Molecular and cellular biochemistry, v. 199, n. 1-2, p. 125-137, 1999. ISSN 0300-8177.

SEARS, A. K. et al. AE37: a novel T-cell-eliciting vaccine for breast cancer. Expert Opinion on Biological Therapy, v. 11, n. 11, p. 1543-1550, 2011/11/01 2011. ISSN 1471-2598. Available at: < https://doi.org/10.1517/14712598.2011.616889 >.

SEGAWA, K. et al. Caspase-mediated cleavage of phospholipid flippase for apoptotic phosphatidylserine exposure. Science, v. 344, n. 6188, p. 1164-1168, 2014. ISSN 0036- 8075.

SHARMA, A. et al. HER‐2 pulsed dendritic cell vaccine can eliminate HER‐2 expression and impact ductal carcinoma in situ. Cancer, v. 118, n. 17, p. 4354-4362, 2012. ISSN 1097- 0142.

SIEGEL, P. M.; MULLER, W. J. Mutations affecting conserved cysteine residues within the extracellular domain of Neu promote receptor dimerization and activation. Proceedings of the National Academy of Sciences of the United States of America, v. 93, n. 17, p. 8878-8883, 1996. ISSN 0027-8424 1091-6490. Available at: .

SLEE, E. A.; ADRAIN, C.; MARTIN, S. J. Executioner caspase-3,-6, and-7 perform distinct, non-redundant roles during the demolition phase of apoptosis. Journal of biological Chemistry, v. 276, n. 10, p. 7320-7326, 2001. ISSN 0021-9258.

SOH, J. et al. Identification and sequence of an accessory factor required for activation of the human interferon γ receptor. Cell, v. 76, n. 5, p. 793-802, 1994. ISSN 0092-8674.

SOLDANI, C. et al. Poly (ADP-ribose) polymerase cleavage during apoptosis: when and where? Experimental cell research, v. 269, n. 2, p. 193-201, 2001. ISSN 0014-4827.

SOLDANI, C.; SCOVASSI, A. I. Poly (ADP-ribose) polymerase-1 cleavage during apoptosis: an update. Apoptosis, v. 7, n. 4, p. 321-328, 2002. ISSN 1360-8185.

STENNICKE, H. R. et al. Caspase-9 can be activated without proteolytic processing. Journal of Biological Chemistry, v. 274, n. 13, p. 8359-8362, 1999. ISSN 0021-9258.

SUMIMOTO, H. et al. The BRAF–MAPK signaling pathway is essential for cancer-immune evasion in human melanoma cells. Journal of Experimental Medicine, v. 203, n. 7, p. 1651-1656, 2006. ISSN 0022-1007.

TAKAMI, K. et al. Interferon-γ Inhibits Hepatocyte Growth Factor-Stimulated Cell Proliferation of Human Bronchial Epithelial Cells: Upregulation of p27kip1 Cyclin- Dependent Kinase Inhibitor. American journal of respiratory cell and molecular biology, v. 26, n. 2, p. 231-238, 2002. ISSN 1535-4989.

TEWARI, M. et al. Yama/CPP32β, a mammalian homolog of CED-3, is a CrmA-inhibitable protease that cleaves the death substrate poly (ADP-ribose) polymerase. Cell, v. 81, n. 5, p. 801-809, 1995. ISSN 0092-8674.

VAN DER KUIP, H. et al. Mechanisms of clinical resistance to small molecule tyrosine kinase inhibitors targeting oncogenic tyrosine kinases. American Journal of Pharmacogenomics, v. 5, n. 2, p. 101-112, 2005. ISSN 1175-2203.

VAN ENGELAND, M. et al. A novel assay to measure loss of plasma membrane asymmetry during apoptosis of adherent cells in culture. Cytometry, v. 24, n. 2, p. 131-139, 1998. ISSN 0196-4763. Available at: < https://doi.org/10.1002/(SICI)1097- 0320(19960601)24:2<131::AID-CYTO5>3.0.CO >. Accessed on: 2018/03/22.

VILCEK, J. Novel interferons. Nature immunology, v. 4, n. 1, p. 8, 2003. ISSN 1529-2916.

VOGEL, C. L. et al. Efficacy and safety of trastuzumab as a single agent in first-line treatment of HER2-overexpressing metastatic breast cancer. Journal of Clinical Oncology, v. 20, n. 3, p. 719-726, 2002. ISSN 0732-183X.

VOGELSTEIN, B.; LANE, D.; LEVINE, A. J. Surfing the p53 network. Nature, v. 408, n. 6810, p. 307, 2000. ISSN 1476-4687.

WAJANT, H. The Fas signaling pathway: more than a paradigm. Science, v. 296, n. 5573, p. 1635-1636, 2002. ISSN 0036-8075.

WEINBERG, R. The Biology of Cancer, Second Edition.Taylor & Francis Group, 2013. ISBN 9781317963462. Available at: < https://books.google.com/books?id=MzMmAgAAQBAJ >.

WILEY, H. S.; BURKE, P. M. Regulation of receptor tyrosine kinase signaling by endocytic trafficking. Traffic, v. 2, n. 1, p. 12-18, 2001. ISSN 1600-0854.

YOUNG, H. A. Regulation of interferon-γ gene expression. Journal of interferon & cytokine research, v. 16, n. 8, p. 563-568, 1996. ISSN 1079-9907.

YUAN, C.-H.; FILIPPOVA, M.; DUERKSEN-HUGHES, P. Modulation of Apoptotic Pathways by Human Papillomaviruses (HPV): Mechanisms and Implications for Therapy. 2012. 3831-50.

ZIMMERMANN, K. C.; GREEN, D. R. How cells die: Apoptosis pathways. Journal of Allergy and Clinical Immunology, v. 108, n. 4, p. S99-S103, ISSN 0091-6749. Available at: < http://dx.doi.org/10.1067/mai.2001.117819 >. Accessed on: 2018/03/13.

ZOU, H. et al. Apaf-1, a Human Protein Homologous to C. elegans CED-4, Participates in Cytochrome c–Dependent Activation of Caspase-3. Cell, v. 90, n. 3, p. 405-413, 1997/08/08/ 1997. ISSN 0092-8674. Available at: < http://www.sciencedirect.com/science/article/pii/S0092867400805012 >.