Engineering Ca2+-Activated Control of Ras Family Gtpases by Janice Hoi

Engineering Ca2+-activated control of Ras family GTPases

Janice Hoi-Yee Wong

A thesis submitted in conformity with the requirements for the degree of Master of Applied Science

Institute of Biomaterials and Biomedical Engineering University of Toronto

Abstract Engineering Ca2+-activated control of Ras family GTPases Janice Hoi-Yee Wong

Master of Applied Science Graduate Department of Biomaterials and Biomedical Engineering University of Toronto 2019

Reprogrammed cells are capable of sensing specified biomarkers for targeting disease sites. My work aims to rewire mammalian cells to use Ca2+ signaling to respond to a broad spectrum of stimuli. Previously, a Ca2+-CaM system replaced the GTP/GDP switch in Rho-

GTPases such that increased intracellular Ca2+ leads to protein activation. This work applied the same strategy to a Rab and Ras GTPase, Rab5 and Ras, respectively. Ca2+-activated Rab5 (called

Raber) induces cell retraction, whereas, Ca2+-activated Ras (called Raser) induces focus formation. To demonstrate rewiring potential to Ca2+ mobilizing proteins, Raber and Raser were co-expressed with either ChR2 or VEGFR2 to indirectly activate them with light and VEGF, respectively. This work demonstrates the potential for these proteins to be activated by any external stimuli via Ca2+ rewiring. In regards to cell-based therapies, Raber can enable cellular detachment, encouraging migration, whereas Raser can enable cells to form foci, accumulating at target sites.

Acknowledgements

To my supervisor, Dr. Kevin Truong, for the support and guidance. To my parents, Eric and Emily Wong, for the unconditional love and encouragement. To Taylor Fox, for the motivation.

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Table of Contents

Abstract ...... ii Acknowledgements ...... iii List of Abbreviations ...... vi List of Figures ...... vii List of Tables ...... vii List of Appendices ...... viii Chapter 1. Introduction ...... 1 1.1. Background and Motivation ...... 1 1.2. Ca2+ Signaling ...... 2 1.3. Rab and Ras GTPases ...... 9 1.4. Research Objectives ...... 13 1.5. Organization ...... 13 Chapter 2. Raber: a Ca2+-activated Rab GTPase ...... 15 2.1. Chapter Aims and Motivation ...... 15 2.2. Literature Review of Rab GTPases ...... 16 2.3. Theory and Method for Creating Raber ...... 20 2.4. Experimental Procedures ...... 22 2.5. Results ...... 24 2.6. Discussion ...... 33 Chapter 3. Raser: a Ca2+-activated Ras GTPase ...... 35 3.1. Chapter Aims and Motivation ...... 35 3.2. Literature Review of Ras GTPases ...... 36 3.3. Theory and Method for Creating Raser ...... 39 3.4. Experimental Procedures ...... 41 3.5. Results ...... 43 3.6. Discussion ...... 48 Chapter 4. Summary and Conclusion ...... 51 4.1. Conclusion ...... 51 4.2. Future Directions ...... 52 References ...... 54

Appendices ...... 63

Appendix A ...... 63

Appendix B ...... 79

List of Abbreviations ATP adenosine 5′-triphosphate CaM calmodulin CaRQ engineered Ca2+-activated RhoA protein ChR2 channelrhodopsin 2 GAP GTPase-activating protein GEF guanine exchange factor GDP guanosine diphosphate GPCR G-protein coupled receptor GTP guanosine triphosphate GTPase guanosine triphosphatase HEK293 human embryonic kidney cell line IQ2p IQ peptide of myosin V MLCKp myosin light chain kinase peptide NIH 3T3 mouse embryo fibroblast cell line PLC phospholipase C PKC phosphokinase C RTK receptor tyrosine kinase VEGF-A vascular endothelial growth factor-A VEGFR2 vascular endothelial growth factor receptor 2

List of Figures Figure 1.1. Therapeutic applications of engineering protein networks Figure 1.2. Calcium signaling network Figure 1.3. Ca2+-mobilizing proteins and pathways Figure 1.4. Calmodulin and its binding peptides, IQ2p and MLCKp Figure 1.5. GTP/GDP switch mechanism Figure 1.6. Physical and chemical stimuli to induce Ca2+ release from ER Figure 2.1. Graphic map of the transfer vector CMVp_LynVenusRb5er_kb32 Figure 2.2. Raber expressing HEK293 cells with ATP stimulation Figure 2.3. Schematic of Ca2+-activated Rab5 protein with light stimulus Figure 2.4. Stimulation of cells co-expressing Raber and ChR2 with light Figure 2.5. Schematic of Ca2+-activated Rab5 protein with VEGF-A Figure 2.6. Stimulation of cells co-expressing Raber and ChR2 with VEGF-A Figure 2.7. Summary graph of protrusion retraction in cells co-expressing Raber and VEGFR2 Figure 3.1. Schematic of Ras-dependent transformation Figure 3.2. Graphic map of the transfer vector CMVp_LynVenusRASer_kb32 Figure 3.3. Comparison of normal and transformed morphologies of NIH3T3 cells at 100% confluency Figure 3.4. Schematic diagram and microscopy images of Raser rewired to ChR2 and VEGFR2 Figure 3.5. Summary graph for foci count in Raser cells rewired to ChR2 and VEGFR2

List of Tables Table 1.1. Ras Superfamily of Small GTPases

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List of Appendices Appendix A. Review on Cell-Based Senotherapies Appendix B. Arduino Software Sketch

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1. Introduction

This chapter provides the motivation for engineering calcium activated Rab and Ras proteins.

Backgrounds of protein engineering, cell-based therapies, calcium signaling, and Ras superfamily proteins are discussed. The research objectives of this thesis are introduced for

Raber, a calcium activated Rab5 protein, and Raser, a calcium activated Ras protein. This chapter also provides an organizational overview of this document.

1.1. Engineering genetic networks for therapeutic applications

Synthetic biology uses engineering concepts to create biomolecular networks and cellular mechanisms for various research and clinical applications (1). Synthetic biologists combine simple functional units, and defined inputs and outputs, to create complex circuit systems (2).

Advances in synthetic biology have led to the use of genetic engineering techniques to develop synthetic gene networks that can control cellular behaviour under context-specific conditions.

Protein engineering involves the creation of original and functional protein units through modification and merging of natural proteins. A protein sequence can be customized to have desired specifications including substrate specificity, regulatory mechanisms, fluorescence- labelling, pH and temperature stability (3). Because the way that they interact between and within cells can be designed, engineered proteins have enabled protein-based approaches in studying and manipulating biological processes such as rewiring cellular networks. As discoveries of protein structures and functions are made, researchers have used rational design and directed evolution approaches to make desired modifications to better suit industrial, medical and research applications.

Figure 1.1. Therapeutic applications of engineering protein networks. The genetically reprogrammed cell is capable of modulating cellular behaviour in the response to a specific biomarker.

Genetic and protein engineering have enabled the creation of synthetic gene and protein networks that are capable of reprogramming cells for therapeutic uses (4-6). First-generation engineered cells are already being used in the clinic and demonstrating a lot of promise as new treatment modalities (7). However, some of the issues with current engineered cell-based approaches include overexpression of a gene product that can lead to adverse effects, and lack of control in terms of their dosage, cellular context, localization, and timing (5). A safer and more effective method is to create engineered cells capable of sensing specified inputs such as disease and/or cell-specific biomarkers to control their behaviour (2) (Figure 1.1).

1.2. Calcium (Ca2+) Signaling

The following work will reprogram mammalian cells to use calcium (Ca2+) signaling to enable cells to respond to a broad spectrum of stimuli. Ca2+ is a ubiquitous second messenger important in nearly every aspect of cellular activity. When cytosolic Ca2+ concentrations are too high, however, it can be lethal to the cell as Ca2+ will precipitate phosphates. Hence, cells invest a lot of energy in chelating, compartmentalizing, and pumping out the ion to tightly regulate Ca2+

concentrations, maintaining the 10,000-fold electrochemical gradient between the cytosol and extracellular milieu (8).

The main functional units of the Ca2+ signaling toolkit include an external stimulus, Ca2+- mobilizing proteins, Ca2+ ions, Ca2+-sensitive proteins, and Ca2+-removing mechanisms (Figure

1.2). To mediate cellular processes with Ca2+ signaling, an external stimulus activates Ca2+- mobilizing proteins/pathways, which in turn increases the cytoplasmic Ca2+ concentrations via plasma membrane channels or release from intracellular stores (i.e. endoplasmic or sarcoplasmic reticulum) (8, 9). The resting cytoplasmic Ca2+ concentration in a typical cell is ~100 nM, which is approximately 10,000-fold and 100 to 500-fold lower than the extracellular milieu and endoplasmic (and sarcoplasmic) reticulum, respectively (10). A range of stimuli – such as extracellular and/or intracellular signaling molecules, and membrane depolarization, can cause a steep influx of Ca2+ to increase cytosolic levels from 100 nM to 1 M or more. In response,

Ca2+-sensitive proteins can directly, or indirectly by activating downstream effector proteins to lead to changes in cellular responses or physiological processes. Finally, Ca2+-removing proteins and homeostatic mechanisms restore cytoplasmic Ca2+ concentrations to resting (pre-stimulus) conditions either by pumping Ca2+ ions out of the cell or into intracellular stores.

Figure 1.2. Calcium signaling network used to modulate cellular behaviour and physiological processes.

The signaling toolkit of Ca2+-mobilizing proteins is diverse and vast, comprising of natural and engineered proteins. Ca2+-mobilizing proteins are responsible for controlling the entry of extracellular Ca2+ and/or release of Ca2+ from intracellular stores (9). The following work demonstrates the versatility of Ca2+ signaling by using a diverse set of external stimuli to control the engineered Ca2+ networks. The Ca2+-mobilizing proteins discussed in this work involve proteins that are endogenous, pre-existing proteins that translate external stimuli into a

Ca2+ signal (Figure 1.3A); and exogenous, proteins introduced via transgene expression to translate external stimuli into a Ca2+ signal (Figure 1.3B). Synthetic Ca2+-mobilizing proteins, which are engineered proteins introduced via transgene expression will not be further discussed as the following work does not involve this group of Ca2+-mobilizing proteins.

Figure 1.3. Ca2+-mobilizing proteins and pathways. A. Endogenous Ca2+-mobilizing proteins. B. Exogenous Ca2+-mobilizing proteins. P2X and P2Y are purinergic receptors. ATP (adenosine 5′- triphosphate) is represented by pink circles. P2X and ChR2 (activated by light) mobilizes Ca2+ by 2+ transporting Ca ions from the extracellular space, whereas P2Y and VEGFR2 utilizes the PLC-IP3 signaling pathway to release Ca2+ ions from the endoplasmic (for muscle cells, sarcoplasmic) reticulum. VEGF-A is represented by green circles.

The endogenous Ca2+-mobilizing proteins, P2X and P2Y are cell surface purinergic receptors that are activated by extracellular purines, with ATP (adenosine 5′-triphosphate) being the most common (Figure 1.3A) (11). P2X receptors are ligand-gated ion channels, and are preferably permeable to sodium, potassium, and calcium ions within milliseconds of binding to

ATP. P2Y receptors are G-protein coupled receptors (GPCRs) that when ATP-bound, cause the

Gq subunit to dissociate and activate phospholipase C (PLC) (12, 13). PLC then catalyzes the formation of DAG (diacylglycerol) and IP3 (inositol 1,4,5-triphosphate). This event is

2+ followed by Ca influx into the cytosol via IP3R (IP3 receptors) which are located on the endoplasmic (sarcoplasmic for muscles) reticulum.

The exogenous Ca2+-mobilizing proteins being discussed are VEGFR2 (vascular endothelial growth factor receptor 2) and ChR2 (channelrhodopsin 2) (Figure 1.3B). VEGFR2

(or Flk1) is a receptor tyrosine kinase (RTK) that is usually found as a monomer, and undergoes dimerization when VEGF-bound for signal transduction (14-16). Out of the 5 VEGF family genes that exist in the mammalian genome, VEGF-A binds to and activates VEGFR2 with the highest affinity (14, 16-18). VEGF-A binding leads to changes in the transmembrane domain conformation of VEGFR2 in order to increase kinase domain phosphorylation (16). This in turn enhances migration of inflammatory cells, hypervascular permeability, and endothelial proliferation (14, 17, 19). Unlike most RTKs, VEGFR2 stimulates ERK via the PLCγ-PKC-

MAPK pathway (20, 21). Phosphorylated PLCγ increases the levels of DAG which activates

2+ PKCs, and IP3 which activates IP3Rs to release Ca into the intracellular space (15).

ChR2 is a light-activated cation-selective channel widely used in optogenetics and biotechnology applications. ChR2 activation involves the photoisomerization of all-trans-retinal to 13-cis-retinal in order cause conformational change to the GPCR, and consequently, increased permeability to monovalent and divalent ions (22). In the context of Ca2+ signaling, light- activated ChR2 leads to Ca2+ entry into the cytosol and depolarization (23, 24). ChR2 is maximally activated at ~470 nm blue light with an open channel state lifetime of ~10 ms (22, 25-

27). Variants of ChR2 have been engineered for increased variety and effectiveness, and rewired to activate other second messengers like IP3 (25, 28, 29). ChR2 has been known to modulate

Ca2+ signals in neuronal cells, cardiomyocytes, and engineered protein networks in mammalian cell lines (25, 30-32). The following work relies on the ability of ChR2 to induce local Ca2+ signals that, with repeated illumination, causes global Ca2+ increases throughout the cytosol (32).

Although applying light as a control setting in therapeutic applications is not largely explored

because of its poor ability to penetrate tissues due to absorption and scattering, light-activated proteins are useful in a laboratory setting as they are easy to control spatially and temporally.

Once an engineered protein network has been characterized, the system’s control settings can be modified for other applications.

Once local and global cytosol Ca2+ signals are generated by the Ca2+-mobilizing proteins, a diverse set of proteins will respond with a range of affinities, cofactor requirements, responses, and complexity of signaling. Ca2+-sensitive (or Ca2+-activated) proteins can be affected directly, with Ca2+ acting as a simple switch via Ca2+ binding motifs, such as calmodulin (CaM), troponin

C (TnC), calcineurin, and calpain. Other downstream effector proteins and networks can be regulated in a Ca2+-dependent manner by associating with a Ca2+-activated protein or enzyme, such as Ca2+/CAM-dependent protein kinase (CaMK) (33). During this process, Ca2+ ions also

2+ bind to various Ca buffering proteins, such as parvalbumin, calbindin-D28K, and calretinin which act to influence the spatial spread, amplitude and duration of Ca2+ signals (34, 35).

The main Ca2+-activated protein network discussed in this work will be CaM and its binding peptides. CaM is a ubiquitously expressed protein has been well characterized for Ca2+ sensing. The ~150 amino acid long (16.7 kDa) protein has a stiff dumbbell shape comprised of a central α-helix connecting two pairs of (the Ca2+ binding) EF-hand motifs (33, 36-38). Like any other protein which relies on conformation and charge to function, Ca2+ affects the conformation of CaM. In resting state, Ca2+-free CaM (apoCaM) has an enclosed hydrophobic core with its

EF-hand motifs in parallel to each other. When Ca2+ bound, CaM undergoes a conformational change to open the EF-hands and expose its hydrophobic surfaces for Ca2+-dependent interactions, rendering it active by increasing its catalytic activity towards its downstream effectors. CaM is a versatile Ca2+ sensor due to having many known binding targets with or

without Ca2+, as well as its ability to respond to a wide range of Ca2+ concentrations (10-12 M to

10-6 M) (39). With CaM binding, target peptides have a natural propensity to adopt an α-helical structure (40). CaM can have Ca2+-independent interactions with its target proteins via IQ motifs.

IQ motifs, named after the first two conserved residues) have a sequence of IQXXXRGXXXR and provide binding sites for CaM and other EF-hand family proteins (41, 42). When cytosolic

Ca2+ levels are low, the interaction between CaM and the IQ motif peptide from myosin V

(IQ2p) forms a low affinity complex in the micromolar range (41). In comparison, when cytosolic Ca2+ levels are high, the Ca2+-dependent interaction between myosin light chain kinase

(MLCKp) and Ca2+-CaM is forms a high affinity complex (in the nanomolar range) (43, 44). The following work will be using a Ca2+-activated network involving CaM and its binding peptides,

IQ2p (from myosin V) and MLCKp (from myosin light chain kinase) (Figure 1.4) (45).

Figure 1.4. Schematic representation of a Ca2+-activated network using the CaM-MLCKp-IQ2p system. A. Primary protein sequence of CaM-MLCKp-IQ2p. The orange segments represent two fragments of the target protein. B. The binding events taking place under low and high Ca2+ concentrations. The CaM-MLCKp-IQ2p system involves replacing the natural regulatory/switch peptide with IQ2p of the target protein. The CaM and MLCKp peptide sequences are fused to the amino-

terminal of the IQ2p embedded sequence. Under low cytosolic Ca2+ levels, apoCaM has a Ca2+- independent binding with IQ2p. When Ca2+ levels increase, Ca2+-CaM disassociates from IQ2p due to its much higher affinity for MLCKp, rendering the target protein active.

1.3. Rab and Ras GTPases

This work will discuss the protein engineering of Ras superfamily proteins. The study of

RAS genes was initially in retrovirus research; it was later discovered that these viral genes were responsible for the highly oncogenic properties of RNA tumor viruses (46-48). The discovery of mutated constitutively active RAS genes in human cancer sparked intensive research efforts to understand the proteins encoded by these genes.

Ras superfamily proteins and their functionalities have been extensively studied for several decades due to their diverse and essential roles in signaling networks, proliferation, differentiation and survival (Table 1.1). In addition, dysregulation of Ras superfamily proteins have been implicated in having critical roles in several human diseases, including cancer and developmental syndromes (49-52). Ras and Ras-related genes are highly evolutionarily conserved among vertebrates and invertebrates (47). In humans, the Ras superfamily can be divided into five main families based on their sequences and functionalities: Ras, Rab, Rho, Arf, and Ran (Table 1.1) (53, 54).

Table 1.1. Ras superfamily of small GTPases present in humans.

Family Subfamilies Function(s) Ras • Ras • Cell proliferation, • Ral differentiation, survival, • Rap apoptosis, gene expression • Rad • GTP-dependent exocytosis • Rheb • Cell-cell and cell-matrix • Rit adhesion • Cell shape remodelling • mTOR pathway • Cell-cycle checkpoint and/or progression • Neuronal differentiation and survival Rab • Membrane and protein trafficking via endocytic and secretory pathways • Membrane organization Rho • Cytoskeleton organization • Cell shape, polarity, adhesion, movement • Cell-cycle progression • Gene expression Arf • Vesicular trafficking • Endocytosis • Exocytosis Ran • Nucleocytoplasmic transport • Mitotic spindle organization

Ras superfamily proteins are small guanosine triphosphatases (GTPases) which are monomeric proteins that are regulated by a GTP/GDP binary switch system. The GTP/GDP switch mainly involves the GTPase fold, composed of a six-stranded β-sheet flanked by five α- helices, is common to all Ras superfamily proteins (55). The GTP-bound form of the switch adopts a specific folded conformation that is capable of interacting with effector proteins, whereas the GDP-bound form causes an unfolding of the switch regions. The process of small

GTPases converting between their active (GTP-bound) and inactive (GDP-bound) states is tightly regulated via regulatory proteins called guanine nucleotide exchange factors (GEFs) and

GTPase-activating proteins (GAPs), respectively (Figure 1.5) (56).

Figure 1.5. The GTP/GDP switch mechanism. GTPase-activating proteins (GAPs) turn GTPases off by catalyzing the hydrolysis of GTP to GDP and inorganic phosphate (Pi). Guanine-nucleotide-exchange factors (GEFs) activate GTPases by catalyzing the exchange of GDP for GTP.

Previously, the subfamily of Rho GTPases (i.e. RhoA, cdc42 and Rac1) were engineered to become Ca2+-activated by replacing the GDP/GTP switch system with a Ca2+-responsive protein calmodulin (CaM) and two binding peptides: IQ2p (myosin V) and MLCKp (myosin light chain kinase) (57-60). Here, we apply the same genetic modification in replacement of the

GDP/GTP switch system, demonstrated with a Rab and Ras protein

The engineered Rab and Ras proteins are designed to be Ca2+-activated to allow rewiring to other protein networks. Ca2+-sensitive proteins are very versatile as calcium signals can be generated by different classes of stimuli such as physical (e.g. light and magnets) and chemical

(e.g. VEGF-A, IL-6) (61-64) (Figure 1.6).

Figure 1.6. Physical and chemical stimuli to induce Ca2+ release from the endoplasmic reticulum.

The Rab proteins form the largest family within the Ras superfamily. By recruiting specific effector proteins, Rab proteins are able to regulate diverse functions with several processes in vesicular trafficking pathways and membrane organization (65-67). Rab proteins have geranylated tails that allow for regulated membrane insertion (55). The Rab protein that will be further discussed and genetically engineered is Rab5, a human Rab subfamily protein commonly known for receptor-mediated endocytosis and pinocytosis (68). Rab5 has also shown to induce actin cytoskeleton reorganization and cell migration (68). Here, we engineered a dominant positive Rab5 protein, called Raber, which is conditionally activated by Ca2+ signalling and capable of protrusion retraction upon activation. The lentiviral transfer vector

CMVp_LynVenusRb5er_kb32 was used to create stable cell lines expressing membrane labelled

Venus fused to the engineered protein.

The Ras proteins participate in complex signaling pathways controlling cell division, migration, adhesion, cytoskeletal integrity, survival and differentiation. In normal cells, Ras activation is tightly regulated. In a mouse fibroblast (NIH3T3) cell line known for strong contact inhibition, dysregulation or overexpression of the Ras signal transduction pathway is known to trigger many events known to occur in malignant transformation such as loss of contact

inhibition, increased proliferation, reduced dependence on serum, acquisition of anchorage- independent growth potential, and ability to generate foci. With protein engineering, the Ras protein can be modified to undergo malignant transformation upon Ca2+ activation. Here, we engineered a dominant positive Ras protein that is conditionally activated by calcium signalling.

1.4. Research Objectives

The superfamily of Ras GTPases are versatile and important regulators in virtually all cellular processes. The goal of this research is to engineer proteins from the Rab and Ras subfamily (specifically Rab5 and Ras, respectively) to show that it is possible to directly control this superfamily of small GTPases with calcium signaling. Since the GTPase fold, composed of a six-stranded β-sheet flanked by five α-helices, is shared among all Ras superfamily proteins (55), it is hypothesized that the genetic modification of a replacing the GTPase fold with a calcium activated sequence – as done previously in Rho GTPases, can be made to replace the GTP/GDP switch system with a calcium signaling system. This thesis will achieve the following goals in an objective-based manner:

1. Determine if the engineered Rab5 protein (Raber) causes actin cytoskeleton

reorganization (e.g. cell detachment, membrane ruffling), and demonstrate rewiring of

Raber to physical and chemical stimuli.

2. Demonstrate rewiring of the engineered Ras protein (Raser) to exhibit cellular

transformation (e.g. generate foci) with physical and chemical stimuli.

1.5. Organization

The remaining chapters of this thesis are organised as follows:

• Chapter 2 provides a literature review of Rab GTPases, discussing the natural functions

and previous applications of Rab5 protein. This chapter also provides the rationale and

design of the engineered Ca2+-activated Ras GTPase, followed by its characterization and

rewiring to ChR2 and VEGFR2. The clinical relevance of engineering a Ca2+-activated

Ras GTPase to encourage cellular migration via cellular actin cytoskeleton reorganization

is also discussed.

• Chapter 3 provides a literature review of Ras GTPases, and a discussion of how its role

in cellular transformation, specifically focus formation, can be applied to genetic circuits

and cell-based therapies. The theory and method of engineering a Ca2+-activated Ras

GTPase is discussed, followed by its characterization and rewiring to ChR2 and

VEGFR2.

• Chapter 4 summarizes the findings, significance, and future directions for engineering

Ca2+-activated Ras family proteins.

• Appendix includes the Arduino script used for the LED dot matrix used in ChR2

stimulation. The section also includes a review article I wrote about a possible context

where engineered gene circuits could be applied (e.g. senotherapy).

Chapter 2

This chapter will provide a literature review related to the role of the Rab5 in cellular actin cytoskeleton reorganization and motility, and upstream regulation and downstream targets of

Rab5. In previous work, our lab has applied a Ca2+-CaM switch system in replacement of the

GTP/GDP switch in Rho GTPases. Upon an increase in intracellular calcium, the engineered fusion protein is activated. This work applied the same strategy to a Rab GTPase, Rab5. An engineered Ca2+-activated Rab5 (called Raber) plasmid was transfected into HEK293 cells, and activated to induce cell actin reorganization via cell retraction. To show its rewiring potential to other Ca2+ mobilizing proteins, Raber was co-expressed with either ChR2 and VEGFR2 to indirectly activate them with light and VEGF, respectively. This work demonstrates the potential for Raber to be activated by any external stimuli via Ca2+ rewiring.

2.1. Chapter Aims and Motivation

Rab5 was chosen to be engineered to become Ca2+-activated as there is no current studies that show Rab5 being naturally regulated by Ca2+-calmodulin. Additionally, an engineered Ca2+- activated Rab5 protein would allow rewiring to other protein networks, enabling cellular detachment, and subsequently, encouraging cellular migration.

The specific aims of this chapter that were drawn from the overall research goals:

1. Design a fusion protein that enables Ca2+-calmodulin control over Rab5.

2. Determine if the Ca2+-activated Rab5 fusion protein could regulate cellular morphology

in response to changes in intracellular Ca2+.

3. Determine if Ca2+ signals could be generated by Ca+-mobilizing proteins to regulate

cellular morphology via the Ca2+-activated Rab5 fusion protein.

The lentiviral transfer vectors CMVp_chr2_Cherry_rtz32 (producing ChR2 fused with mCherry under the CMV promoter), CMVp_VEGFR2_CMVp_Ceru3NES_ktz32 (producing nuclear excluded Cerulean and VEGFR2), and CMVp_LynCherry_CMVp_VEGF_kb32 (producing membrane labelled mCherry and the VEGF cytokine) were used to create stable cell lines in this chapter during the characterization of the Ca2+-activated Rab5 fusion protein.

2.2. Literature review of Rab GTPases

Rab (ras‐related in brain) GTPases constitute the largest subfamily of small GTPases with almost 70 different members encoded in the human genome (69). Rab GTPases can be further divided into subfamilies: early endocytic (Rab5, Rab21, Rab22)(70), late endocytic (Rab7,

Rab9)(71), recycling (Rab4, Rab11)(72), and secretory (Rab3, Rab27)(67, 73). Rab GTPases can exist as in both soluble and membrane-bound forms as well as several isoforms to perform similar functions, differing in regulation and differential expression dependent on cell type (53).

During post-translational modifications, Rab escort protein (REP) associates newly synthesized

Rab to a Rab-geranylgeranyltransferase (Rab-GGT) to enable membrane localization through a carboxy-terminal prenylation motif (74). Rab GTPases are well known for their essential roles in regulating vesicular transport, early endosome biogenesis and maturation, phosphatidylinositol

3-phosphate (PI3P) kinase activity, and membrane organization (66, 67). The regulation of Rab proteins are highly complexed since there are so many signaling molecules associated with their function. Hence, Rab proteins are involved in multiple cellular functions in addition to the initially proposed endosome trafficking. Other roles Rab GTPases have been associated with are regulation of cellular proliferation, apoptosis, adhesion, and motility (75). In recent years, there has been increased interest on the role of Rab GTPases in migration and invasiveness of

metastatic tumour cells as it has been shown that these processes are regulated by components of the endocytic machinery (75).

The natural regulation of Rab5 proteins by calcium is rather indirect. Intracellular calcium levels tightly control the endocytic capacity at many synapses and other non-neuronal cells (76). Dependent on cell-type, intracellular calcium transients may trigger and/or speed up classical endocytosis. Calcium may also exhibit opposite effects by inhibiting or slowing down endocytosis, although the mechanism is still heavily debated (77). A proposed mechanism involves the calcium/calmodulin-activated phosphatase, calcineurin, to dephosphorylate many endocytic proteins including Rab GTPases; however, calcineurin is not considered a key molecule in mediating endocytosis.

2.2.1. Isoforms and Main Effectors of Rab5

Rab5 is ubiquitously expressed but is primarily localized to the plasma membrane, clathrin coated vesicles, and early endosomes (78, 79). A common feature of Rab family of small

GTPases is the existence of structurally related isoforms with high sequence identity (80). That being said, Rab5 exists as three isoforms in human and mouse genomes – Rab5a, Rab5b, and

Rab5c (81). Although sharing 90% sequence homology, Rab5 isoforms are expressed by different genes, phosphorylated differently, and serving overlapping but non-redundant functions

(78).

Rab5a: The main functions that have been described for Rab5a include Rac1 activation, actin cytoskeleton remodeling, and migration (82). While all Rab5 proteins are responsive to epidermal growth factor (EGF) stimulation, Rab5a has higher sensitivity to EGF. Epidermal growth factor receptor (EGFR) activation stimulates signal transduction pathways (i.e. Raf-

Erk1/2 pathway) that mediate a spectrum of cellular processes including cell proliferation, differentiation, and apoptosis (83). As a negative feedback mechanism, EGF stimulations also leads to internalization of EGF receptor (EFGR). Activated EGF-EGFR complexes also translocate to the nucleus and binds to specific AT-rich DNA sequences, either directly or as a co-activator, to activate gene transcription that includes cyclin D1 and NFkB (83-85). Moreover,

Rin1 (Ras interaction/interference 1), a Rab5 GEF, has been also found to preferably associate with Rab5a, selectively blocking EGF activation of the Raf-Erk1/2 pathway (86). Another role

Rab5a has been found to be involved with is phagosome maturation, with upregulated transcription levels following treatment with cytokines such as interleukin-4 and interferon-γ

(87-89). Rab5 and other Rab effector proteins, including Rin1, promote the formation of enlarged early endocytic (EEE) compartment (90). Retroviral expression of dominant negative

Rab5:S34N mutant and siRNA Rab5a silencing studies demonstrated that Rab5a is essential for the large endosome phenotype and phagocytic receptor localization in macrophages(88). In migrating T-cells, chemokine receptors and integrins on the cell surface will activate PKC (82).

PKC phosphorylates Rab5a at Thr-7, which is crucial for activation of Rac1, actin cytoskeleton remodeling, and migration (82).

Rab5b: In primary hippocampal cultures, Rab5b plays a neuroprotective role (91-93). DHPG

((S)-3,5-dihydroxyphenylglycine) is an agonist of a metabotropic glutamate receptor and is known for protection against NMDA receptor-mediated cell death (91, 94). DHPG induces upregulation of Rab5b synthesis. Rab5b facilitates the clathrin-dependent internalization of

NMDA and AMPA receptors, decreasing their availability on the cell membrane (92).

Rab5c: Unlike Rab5a and Rab5b isoforms, Rab5c is not involved in EGF-EGFR signal transduction pathways (83). Instead, Rab5c has been shown to be involved in cell migration

during zebrafish granulation, cell invasion via integrin recycling, and cell motility via Rac1 (95-

97). By growth factor activation of Rac1, Rab5c can modulate membrane ruffle formation and cell motility (97). Mediated by Rab5, Rac1 activation occurs on early endosomes where the

RacGEF Tiam1 is also recruited. Subsequent recycling of Rac1 to the plasma membrane ensures efficient and localized signaling (98). Rac1 stimulates the formation of actin-based migratory protrusions through interaction with the WAVE complex (98, 99).

2.2.2. Role of Rab5 in Cell Actin Cytoskeleton Reorganization

Cell migration is understood as a multi-step process involving cell polarization, plasma membrane protrusion, formation of new adhesion sites, cell body contraction and turnover of adhesion complexes (75, 100). At the early endosome level, an increasing amount of Rab proteins have been associated with cell migration. Rab5 is a Rab GTPase primarily known for its canonical role in early endosome dynamics. Rab5 has been of particular interest to researchers due to its numerous binding partners and effectors. Consequently, there has been increased discoveries and understanding of Rab5 in non-canonical roles. It has been shown that Rab5 promotes cellular migration in vitro and in vivo by several mechanisms: integrin trafficking, Rho

GTPase signaling, and adhesion complex dynamics (101-104).

Integrins are both adhesion molecules and the main cell surface receptors, which mediate the interactions between the actin cytoskeleton and extracellular matrix. Integrins are capable of forming a variety of heterodimers resulting from a combination of 18 α and 8 β subunits (105).

Clusters of integrins and adaptor proteins form complexes known as integrin-based focal adhesions (FAs) (106). FAs are also considered signaling platforms as they initiate their own assembly and disassembly during cell migration (107). It has been proposed that a circulating

flux of integrins rather than steady state levels at the cell surface are essential in coordinating cell migration (102). Integrin availability at the cell surface is tightly regulated by different levels of endocytic routes via internalization and recycling. Since a fraction of integrins are not degraded and remain in endosomal compartments, this means there is a readily available pool of integrins for recycling and creating new FAs. Mobile fractions of integrins and other FA molecules are regulated by Rab5 in an activity-dependent manner (108). Integrins were already known to traffic through Rab5 and Rab21 positive endosomes (104, 109). In addition, Rab5-GTP activity was shown to increase at the leading edge of migrating cells, and the localized activation promoted integrin internalization and consequently, FA disassembly (103). Rab5 promotes disassembly of focal adhesions and modulates downstream pathways of integrin signaling, involving other proteins such as Ras and Rho family GTPases.

In endothelial cells, Rab5 GEF Rin2 localises to adhesion complexes at the leading edge of the cell, and tis involved in endocytosis of active β1 integrins (110). Rin2 recruits Ras proteins to further promote integrin internalization, and activates Rac via Tiam1. The Rab5 GAP inactivates Rab5 activity to suppress integrin internalization of β1 integrins (111).

2.3. Theory and method for creating Raber

Rab5 proteins are implicated in regulating actin cytoskeleton reorganization. This main interest of this study is the relationship between Rab5 and cellular morphology, specifically cellular detachment; hence, the output monitored and characterized will be protrusion and edge detachment. Naturally, Rab5 is regulated by a GTP-GDP switch system. By replacing the GTP-

GDP switch system with calmodulin and calmodulin binding peptides, Rab5 can become calcium activated.

Figure 2.1. Graphic map of the transfer vector CMVp_LynVenusRb5er_kb32 . The proposed Ca2+- dependent Rab5 switch system comprises of several main components within its transfer vector: promoter sequence (CMVp), membrane-labeled fluorescent protein/probe (LynVenus), dominant positive Rab5 protein (Rab5-DP), calmodulin (CaM) and associated binding peptides (MLCKp, IQ2p), and blastcidin resistance (kb32).

The sequence of Raber has 660 amino acids:

MGCIKSKGKDSAPRVSKGEELFTGVVPILVELDGDVNGHKFSVSGEGEGDATYGKLTLKLICTTGKLPVPWPTLVTTLG YGLQCFARYPDHMKQHDFFKSAMPEGYVQERTIFFKDDGNYKTRAEVKFEGDTLVNRIELKGIDFKEDGNILGHKLEY NYNSHNVYITADKQKNGIKANFKIRHNIEDGGVQLADHYQQNTPIGDGPVLLPDNHYLSYQSKLSKDPNEKRDHMVLL EFVTAAGITLGMDELYKPREQIAEFKEAFSLFDKDGDGTITTKELGTVMRSLGQNPTEAELQDMINEVDADGNGTIYFPE FLTMMARKMKDTDSEEEIREAFRVFDKDGNGYISAAELRHVMTNLGEKLTDEEVDEMIREADIDGDGQVNYEEFVQM MTAKSRKRRWKKNFIAVSAANRYKKISSSGALTSRGATRPNGPNTGNKICQFKLVLLGESAVGKSSLVLRFVKGQFHEF QESTIGAAFLTQTVCLDASGSAITVQRYVRGIQARAYARFLASDTTVKFEIWDTAGLERYHSLAPMYYRGAQAAIVVYD ITNEESFARAKNWVKELQRQASPNIVIALSGNKADLANKRAVDFQEAQSYADDNSLLFMETSAKTSMNVNEIFMAIAK KLPKNEPQNPGANSARGRGVDLTEPTQPTRNQ

The amino acids that are highlighted yellow, is the Lyn peptide for membrane localization; green is Venus for fluorescent labeling; cyan is CaM; purple is MLCKp; red is

Rab5; grey is IQ2p; and uncoloured are linker amino acids.

An IQ-motif peptide (IQ2p) was embedded between residues 62 and 63 of Rab5. There is

Ca2+-independent association between calmodulin (CaM) and myosin V (IQ2p) peptide (affinity in the micromolar range). This association between CaM and IQ2p prevents the folding of the functional Rab5 protein sequence, as well as Rab5 interaction with effector proteins. CaM-

MLCKp was fused to the amino-terminal of Rab5 and IQ2p. When there is an increase in intracellular calcium, myosin light chain kinase (MLCKp) acquires a higher affinity for Ca2+-

CaM, leading to dissociation of CaM to IQ2p, and allowing the Rab5 protein sequence within

Raber to fold into its active, functional form.

2.4. Experimental Procedure

2.4.1. Plasmid construction

The transfer vectors CMVp_LynVenusRb5er_kb32, CMVp_chr2_Cherry_rtz32,

CMVp_VEGFR2_CMVp_Ceru3NES_rtz32, and CMVp_LynCherry_CMVp_VEGF_kb32 were synthesized by Genscript (Pescataway, NY) and subcloned into the pUC57-Simple vector using

EcoRV. LynVenus and LynCherry, were synthesized as the tandem fusion of the signal sequence of Lyn (1MGCIKSKGKDSA12) with mVenus and mCherry, respectively. CeruNES was synthesized as the tandem fusion of mCerulean and the nuclear export signal

(1LQLPPLERLTLD12) from the HIV-1 Rev protein. All genes were highly expressed with the cytomegalovirus promoter (CMVp) in mammalian cells. All plasmid manipulations were performed by Genscript. All plasmids were transformed in E. coli DH5-α and isolated using a

Mini-prep kit (Invitrogen).

2.4.2. Cell lines

Human embryonic kidney (HEK293) cells were maintained in Dulbecco's modified Eagle's medium containing 25 mM D-glucose, 1 mM sodium pyruvate and 4 mM L-glutamine

(Invitrogen), supplemented with 10% fetal bovine serum (Sigma-Aldrich), in T25 flasks (37°C and 5% CO2). All cell lines were authenticated and tested for contamination. Cells were passaged at 90% confluency using 0.05% TrypLE with Phenol Red (Invitrogen) and seeded onto

24-well plates (Corning) at 1:20 dilution. Cells were transiently transfected using Lipofectamine

3000, according to the manufacturer's protocols (Invitrogen). Then, 24 h post-transfection, cells were treated with 0.05% TrypLE with Phenol Red (Invitrogen) and plated in six-well tissue culture plates (Corning) at a serial dilution, 1:2, 1:4, 1:8, 1:16 and 1:32. All stable HEK 293 cell lines were generated by lentiviral infection using the respective SINp transfer vector and subsequent selection with blasticidin (10 μg/ml), puromycin (1 μg/ml) or zeocin (200 µg/ml) for

2 weeks. Colonies were plated in 96-well tissue culture plates (Sarstedt) for subsequent experiments and imaging.

2.4.3. Cell stimulation

Cells were plated in 96-well tissue culture plates prewashed with and imaged in PBS (Thermo

Fisher Scientific). For light and VEGF-A stimulation, cells were imaged in culture media.

2.4.4. Live cell imaging

Imaging was performed using an inverted IX81 microscope with a Lambda DG4 xenon lamp source and a QuantEM 512SC CCD camera with a 10× or 40× objective (Olympus). Filter excitation (EX) and emission (EM) bandpass specifications (in nm) were as follows: CFP (EX,

438/24; EM, 482/32), YFP (EX, 500/24; EM, 542/27), RFP (EX, 580/20; EM, 630/60)

(Semrock). Image acquisition and analysis were performed with µManager and ImageJ software, respectively (112, 113).

2.4.5. Data analysis

All experiments with n=3 were conducted with at least ten cells evaluated in each trial. All data with normal distribution and similar variance were analyzed for statistical significance using

two-tailed, unpaired Student's t-tests, unless otherwise indicated. Multiple group comparisons were made with two-factor ANOVA with replication, followed by Tukey-Kramer HSD post-hoc test, unless otherwise indicated. For all tests, α was set at 0.05. P<0.05 was considered significant. Data are mean ± standard deviation unless otherwise stated. Data was analyzed using the Data Analysis ToolPak for Excel (Microsoft).

2.5. Results

The experimental results for this chapter will be discussed below. The CaM-Rab5 fusion protein was shown to mediate cellular/protrusion detachment upon ATP stimulation. Light and VEGF protein were also used with ChR2 and VEGFR2, respectively, to rewire with CaM-Rab5 fusion protein to induce actin reorganization via cellular/protrusion detachment.

2.5.1. Ca2+-dependent Protrusion and Edge Retraction by Raber

Retraction of cellular protrusions and/or cell edge can be used as an output signal for the

Raber fusion protein. Raber was infected into human embryonic kidney (HEK293) cell line and selected with blastcidin. The stable cell line of HEK293 cells expressing the Raber vector have a

Ca2+-activated sequence, functional Rab5 sequence, and Venus which labeled to the plasma membrane through tyrosine kinase Lyn (1MGCIKSKGKDSA12). With LynVenus, the cell periphery is outlined to show plasma membrane localization, and is visible under fluorescent

(YFP) microscopy (Figure 2.2B). Stable cell lines were infected by lentiviral infection to obtain homogenous cell colonies.

HEK293 cells have endogenous purinoreceptors. Hence, a stable RCaMP1.07 HEK 293 cell line was used to report the occurrence of a Ca2+ signal upon ATP stimulation, visible by red

(RFP) channel (Figure 2.2A) (114). No significant retraction occurred in RCaMP1.07 cells (1.7 ±

1.7%) (Figure 2.2C). Addition on a 10μM bolus of ATP triggered a Ca2+. When a bolus of 10µM

ATP was added, Raber cells showed retraction of cellular protrusions (Figure 2.2B). Retraction was quantified by calculating the displacement of the outermost point on a cellular protrusion, or edge. The average retracted distance in Raber cells upon ATP stimulation after 15 mins was 30.7

± 8.2 μm (Figure 2.2B). The percentage of Raber cells that demonstrated edge and/or protrusion retraction following ATP treatment was 92.6 ± 3.8% (Figure 2.2C). Raber cells with PBS added was used as a control, with 12.8 ± 6.2% of cells retracting.

Figure 2.2. (A) RCaMP expressing cells upon ATP stimulation show increase in intensity upon ATP stimulation. Scale bar is 30 μm (left). ATP was added at 30 seconds (right). (B) HEK 293 cells expressing Raber over 5 mins upon addition of 10M bolus of ATP (left) and average change in protrusion length after ATP addition (15 mins) (right). Scale bar is 20 μm. The average retracted distance before and after ATP stimulation was 30.7 ± 8.2 µm. Data is displayed as mean ± standard error. Matching markers for

same cells. Average protrusion length indicated by black line between the groups. n = 7, with at least 3 protrusions measured per cell. p < 0.05 (p = 0.000140542). Statistical analysis performed with single factor ANOVA, followed by paired student t-test assuming unequal variances. (C) Comparison of percentage of cells retracting. 10M bolus of ATP was added to stimulate Raber and RCaMP cells and imaged for a minimum of 10 mins. Raber cells were recorded before ATP addition. RCaMP cells increase in fluorescent intensity when stimulated with ATP to indicate the presence of a generated calcium signal, however, do not display retraction. One way ANOVA with replication. Followed by post hoc analysis using paired student t-test and Tukey-Kramer HSD. Data is displayed as mean ± standard deviation from three independent trials (n = 3) with at least 10 cells per trial,  = 0.05.

2.5.2. Controlling Cell Detachment via a Physical Stimuli: Rewiring of Raber

to Channelrhodopsin 2 (ChR2)

To examine control of Ca2+-activated Rab5 fusion protein, Raber, with a physical stimulus, HEK293 cells were co-infected with CMVp_LynVenusRb5er_kb32 and

CMVp_chr2_Cherry_rtz32 vectors, and selected using blastcidin and zeocin, respectively. The result was a stable cell line expressing the fusion proteins Raber, and channelrhodopsin 2 (ChR2) which is in tandem fusion to a membrane labeled red fluorescent protein, mCherry (Figure 2.3).

ChR2 is a light-activated cation channel, maximally activated at 440 nm light, and induces a

Ca2+ influx, increasing the local membrane Ca2+ concentration by 1-9 µM (115, 116). From previous studies, ChR2 has been applied to modulate Ca2+-responsive behaviour using light.

Cells that co-expressed yellow and red fluorescence were used to verify the presence of Raber and ChR2, respectively. A second stable cell line expressing only ChR2 (verified by red fluorescence from mCherry) was infected with CMVp_chr2_Cherry_rtz32. The two stable cell lines were stimulated by 10 second pulses of blue light every 30 seconds for 60 mins in a 96 well culture plate.

Figure 2.3. (A) Schematic of light stimulation to induce a Ca2+ signal and subsequent Raber activity. (B) HEK293 cells under YFP (left) and RFP (right) channel to verify presence of fusion proteins Raber and ChR2, respectively. Scale bar is 20 μm.

Without light, Raber ChR2 cells had a 1.4 ± 2.5% basal rate of protrusion/edge retraction

(Figure 2.4C). After 40 mins of blue light stimulus, 82.6 ± 16.0% of Raber ChR2 cells showed edge/protrusion retraction (Figure 2.4C). Cells expressing only Raber or ChR2 were also subjected to blue light stimulus, with only 13.1 ± 14.2% and 0.5 ± 0.9% displaying edge/protrusion retraction, respectively (Figure 2.4C). As a positive control, 10 μM ATP

stimulus resulted in 75.6 ± 16.3% of cells showing detachment and subsequent retraction of protrusions.

Figure 2.4. (A) Time lapse of Raber ChR2 cells showing actin reorganization via protrusions retracting under blue light stimulation. Imaged under YFP channel. (B) Control ChR2 cells under blue light stimulation. Imaged under RFP channel. (C) Summary graph of Raber ChR2 cells exhibiting retraction upon 40 mins of blue light (hv) and/or 10M bolus of ATP stimulus. * denotes p < 0.01 by unpaired t- test. Data is displayed as mean ± standard deviation from three independent trials (n = 3) with at least 10 cells per trial, α = 0.05.

2.5.3. Controlling Cell Detachment via a Chemical Stimuli: Rewiring of Raber

to VEGFR2

To continue observation of the versatility of rewiring and controlling Raber, a stable cell line was infected to control the fusion protein with a chemical stimulus. HEK293 cells were co- infected with CMVp_LynVenusRb5er_kb32 and CMVp_VEGFR2_CMVp_Ceru3NES_ktz32, and selected using blastcidin and zeocin, respectively. The result was a stable cell line expressing the fusion proteins Raber, vascular endothelial growth factor receptor 2 (VEGFR2) and a cerulean fluorescent protein with a C-terminal nuclear export signal, named CeruNES (Figure

2.5) (117). Hence, HEK293 cells co-expressing Raber and VEGFR2 are yellow fluorescent on the cell periphery and cerulean fluorescent in the cellular cytoplasm (Figure 2.5B).

CMVp_LynCherry_CMVp_VEGF_kb32 was infected into HEK293 cells, and selected with blastcidin to obtain a stable cell line of VEGF-A secreting cells that are membrane labeled by

LynCherry to have a red fluorescence. To stimulate cells expressing Raber and VEGFR2, supernatant from VEGF-A secreting cells was used. Dilutions containing 17%, 33%, and 67% of supernatant were measured for percentage of cells retracting to obtain a dose curve (Figure

2.6B).

Figure 2.5. (A) Schematic of how VEGF-A induces Ca2+ signal and subsequent Raber activity. Scale bar is 20 μm. (B) Verification of cells expressing Raber and VEGFR2 by observance of yellow (left) and cerulean (right) fluorescence. Scale bar is 20 μm.

Figure 2.7 summarizes the average retraction rate for Raber and VEGFR2 expressing cells. Upon addition of VEGF-A source cell supernatant, 75.2 ± 5.2% of cells expressing Raber and VEGFR2 displayed detachment of protrusions and subsequent retraction. Cells that only expressed Raber had 17.6 ± 5.1% of cells retracting. As a control, 10 μM ATP stimulus caused

81.7 ± 6.3% of Raber and VEGFR2 expressing cells to show cellular detachment and retraction.

Figure 2.6. (A) Time lapse images of Raber VEGFR2 cells upon addition of VEGF-A source cell supernatant. (B) Dose response of CMVp_LynCherry_CMVp_VEGF_kb32 (VEGF-A source) cell supernatant. The dilution is described as the percentage of VEGF-A source cell supernatant in the total diluted solution. Data represented as mean ± standard deviation, n=3 with at least 8 cells in each trial.

Figure 2.7. Summary graph showing percentage of cells exhibiting protrusion/edge retraction upon addition of either VEGF-A source cell supernatant, or 10 M bolus of ATP stimulus. Asterisk (*) denotes P < 0.01 by unpaired t-test. Data is displayed as mean ± standard deviation from three independent trials (n = 3) with at least 10 cells per trial, α = 0.05.

2.6. Discussion

Rab5 is part of a group of endocytic proteins that have shown to mediate cell actin organization and motility (75). The natural regulatory network of Rab5 is very complex. The aim of the work was to engineer a Ca2+-activated Rab5 protein. By doing so allows direct control of

Rab5 functionality using calcium, and allows efficient rewiring to other synthetic protein networks. It was shown that the engineered Ca2+-activated Rab5 fusion protein was capable of cellular detachment via protrusion/edge retraction. Cells were observed for their ability to undergo cell actin reorganization through cellular detachment of protrusions. To characterize

Raber, the transfer vector CMVp_LynVenusRb5er_kb32 was infected in HEK293 cells to create a stable cell line. Since HEK293 cells endogenously express purinoreceptors, they are capable of inducing an intracellular Ca2+ transient with ATP (as seen in RCaMP1.07 cells). In Raber cells, a

bolus addition of ATP was sufficient to induce a Ca2+ transient and activate Raber activity.

Calcium enables dissociation of the calmodulin (CaM) sequence from IQ2p sequence embedded within the engineered Rab5 protein to associate with MLCKp instead, thus the Rab5 protein sequence is free to fold properly into its active and functional form. For further evidence, using

Rab5 dominant positive and dominant negative mutants, Rab5:Q79L and Rab5 S34N (or

Rab5:N133I) (118, 119), respectively, could be used as additional controls to supplement

RCaMP1.07 cells.

Since it was shown that Raber was Ca2+ responsive, the next step was to add it to a protein network to show rewiring capabilities. Channelrhodopsin 2 (ChR2) and vascular endothelial growth factor receptor 2 (VEGFR2) were chosen as calcium mobilizing proteins to demonstrate rewiring of Raber to a physical (i.e. light) and chemical (i.e. VEGF-A ligand) stimulus (120). In cells expressing Raber and ChR2, cells were stimulated by blue light over a time course of 40 mins. In cells expressing Raber and VEGFR2, cells were stimulated by adding supernatant from VEGF-A secreting cells. Cells that expressed Raber but neither ChR2 nor

VEGFR2 were incapable of showing significant cellular detachment despite undergoing stimulation by light and VEGF-A, respectively. This exhibits the specificity of inputs required to activate Raber. In the future, Rab5 mutants as previously mentioned should be used to further support the findings. In addition, using inhibitors such as CAS 1177865-17-6 (121), a Rac1 inhibitor that interferes with the interaction between Rac1 and RacGEFs Trio and Tiam1 could further support that Raber has a role upstream of Rac activity and thus, cell actin reorganization.

That being said, with Raber shown to rewire with Ca2+-mobilizing proteins, Raber could be incorporated into protein networks to enable cell detachment and encourage cell migration.

Chapter 3

This chapter will provide a literature review of Ras GTPases, in regards to its regulatory

networks, downstream targets, and role in cellular transformation. This work will focus on

replacing the GTP/GDP switch in the Ras GTPase, Ras, with the Ca2+-CaM switch system to

engineer a Ca2+-activated Ras protein (called Raser) to enable focus formation. Upon an increase

in intracellular calcium, the engineered fusion protein is activated. The Raser transfer vector was

transfected into NIH3T3 cells, and activated to induce cellular transformation via focus

formation. To show its rewiring potential to other Ca2+ mobilizing proteins, Raser was co-

expressed with either ChR2 and VEGFR2 to indirectly activate them with light and VEGF,

respectively. This work demonstrates the potential for Raser to be activated by any external

stimuli via Ca2+ rewiring.

3.1. Chapter Aims and Motivation

Ras was chosen to be engineered to become Ca2+-activated as there is no current studies that

show Ras being naturally regulated by Ca2+-calmodulin. Additionally, an engineered Ca2+-

activated Ras protein would allow rewiring to other protein networks, cellular transformation via

focus formation.

The specific aims of this chapter that were drawn from the overall research goals:

4. Design a fusion protein that enables Ca2+-calmodulin control over Ras.

5. Determine if the Ca2+-activated Ras fusion protein could induce cellular transformation

by focus formation in response to increases in intracellular Ca2+.

6. Determine if Ca2+ signals could be generated by Ca+-mobilizing proteins to induce focus

transformation via the Ca2+-activated Ras fusion protein.

The fusion proteins: CMVp_chr2_Cherry_rtz32, CMVp_VEGFR2_CMVp_Ceru3NES_ktz32, and CMVp_LynCherry_CMVp_VEGF_kb32 were used in this chapter during the characterization of the Ca2+-activated Ras fusion protein.

3.2 Literature review of Ras GTPases

Sharing the same name as its superfamily classification, the Ras protein family is a class of small GTPases that participate in complex signaling pathways controlling cell division, migration, adhesion, cytoskeletal integrity, survival and differentiation (122). Ras activation is tightly regulated in normal cells. When mutations in Ras proteins and/or their regulatory network occur, constitutively active Ras-GTP proteins may result in malignant transformation, and play crucial roles in the development and progression of cancer (123, 124).

3.2.1 Ras-dependent Cellular Transformation

Malignant transformation is typically associated with cytoskeleton and cell adhesion rearrangement, increased proliferation, reduced dependence on serum, loss of contact inhibition, acquisition of anchorage-independent growth potential, and ability to generate foci (a cluster of cells that have grown in a multilayered manner) (Figure 3.1). In mouse fibroblast (NIH3T3) cells, a cell line known for strong contact inhibition, the Ras signal transduction pathway is known to trigger many of these events (known as Ras or Ras-dependent transformation) (125,

126). Instead of dividing normally into confluent monolayers, transformed NIH3T3 cells grow into foci. NIH3T3 cells have been used extensively in focus formation assays to study the oncogenic potential of a gene.

Figure 3.1. Schematic of Ras-dependent transformation. (A) Normal fibroblast cells grow in monolayers in a regular pattern. (B) Mutations in Ras proteins or their regulatory networks may lead to high and constitutive levels of active Ras-GTP activity. Transformed cells with increased Ras-GTP activity may lose or have reduced contact inhibition, anchorage-dependent growth, and serum dependence. In addition, transformed cells can acquire the ability to generate foci.

A diverse set of extracellular stimuli can lead to activation of the Ras-mediated pathway

(127, 128). Ras is known to be a downstream effector of RTKs such as EGF and PDGF receptors, and PKC, via RasGEFs that exchange between GDP and GTP (129-134). It involves activated RTKs and the association of adaptor proteins (e.g. Grb2) to recruit RasGEFs (e.g. SOS,

RasGRF) to the plasma membrane which consequently leads to formation of active GTP-bound

Ras (Ras-GTP) and its effector proteins (135, 136).

Phospholipase C (PLC) is an isozyme that is also a Ras activator. PLC runs downstream of distinct signal transduction pathways such as RTKs (PLCγ), GPCRs (PLCβ), Ras/Rho small

GTPases (PLCε) (137). PLC is a catalyst for converting membrane phosphoinositides into IPs and DAG. DAG is capable of recruiting the RasGEF called RasGRP to the plasma membrane to activate Ras. The N-terminus of PLCε has a RasGEF domain which makes PLCε capable of activating the Ras-MAPK-ERK pathway. PLCε is an effector for human H-Ras, binding of H-

Ras directly to the Ras association (RA) domains of PLCε led to translocation to the plasma membrane, and stimulation of IP3 and DAG production, and calcium release (138-140).

There has also been evidence for calcium signaling being upstream and regulating Ras activity (141). A group of RasGEFs called Ras guanine nucleotide-releasing factors (GRFs) and

Ras guanine nucleotide-releasing proteins (GRPs) (or CalDAG-GEFII) have binding motifs for one or both of the second messengers, calcium and DAG. Ras GRPs have EF-hand and C1 domains that allow binding of calcium and DAG, respectively (142). In the context of calcium signaling, several things are not clear about this class of RasGEFs – whether these domains serve as positive or negative regulators of Ras, the binding affinities of the EF hands for calcium, and whether they are a regular sensor of calcium due to the short spacer sequence between the EF hand motifs (143, 144). That is to say, the role and degree of direct Ras regulation by calcium is complex and not yet fully understood.

Effectors of Ras are characterized by Ras binding (RBD) or Ras association (RA or

RalGDS/AF-6) domains that preferentially bind to Ras-GTP (145, 146). The Raf family members are cytoplasmic serine/threonine kinases that are also one of the best characterized class of Ras effectors (47). The Raf-MEK-ERK cascade is involved in turning on transcription factors for growth and development, and oncogenic growth and transformation. The Raf-MAPK-

ERK cascade is not enough for Ras-induced transformation and requires activation of other effectors such as phosphoinositide 3-OH kinase (PI3K) (147). The p110 (α, β, γ, and δ) catalytic subunit of PI3K is an AKT serine/threonine kinase and involved in actin cytoskeleton regulation by growth factors such as PDGF and insulin (148-153). Its role in Ras-induced transformation is mediated by Rac activation (147). However, PI3K pathway is not sufficient for Ras-induced transformation, and works cooperatively with Raf (154).

Ras-dependent transformation is also accompanied by changes in actin cytoskeleton. The activation of Cdc42, Rac1, and RhoA, are essential for Ras-induced malignant transformation

(155). Cdc42 is involved in filipodia extension. Rac1 controls lamellipodia and ruffling behaviour, and RhoA regulates focal adhesions (156, 157). Cdc42 mutants that spontaneously exchange GDP for GTP have shown to transform NIH3T3 cells because of their ability to interfere with EGFR down-regulation (158). Cdc42 activates Rac, which in turn activates

Rac/Rho GTPases (155).

In previous studies of Ras-dependent cellular transformation, mutant H-Ras alone was not able to transform primary rodent fibroblasts, but required activation of an oncogene (e.g.

Myc), or inactivation of a tumour suppressor (p53) (159, 160). That is to say, in some cases, many cooperating events must occur due to the tight regulation of cellular Ras activity. Many cases of Ras-dependent transformation are due to mutant Ras proteins that are insensitive to

GAP, and consequent accumulation of active Ras-GTP (47). The genetic engineering of Ras proteins could potentially make calcium signaling sufficient to induce Ras-dependent processes.

In the context of genetically engineered cell-based therapies, a Ca2+-activated Ras protein could enable cells to accumulate at disease sites more effectively and be rewired to different Ca2+- mobilizing protein networks.

3.3. Theory and method for creating Raser

This main interest of this study is to engineer a protein to directly connect Ras and cellular transformation. With that said, the output monitored and characterized will be focus formation. Naturally, Ras is regulated by a GTP-GDP switch system with several upstream regulators. By replacing the GTP-GDP switch system with calmodulin and calmodulin binding peptides, Ras can become calcium activated. Figure 3.2 illustrates the transfer vector that will be

infected into NIH3T3 cells to result in a stable cell line expressing the proposed engineered Ca2+- activated Ras protein.

Figure 3.2. Graphic map of the transfer vector CMVp_LynVenusRASer_kb32.

The sequence of Raser is consists of 635 amino acids as follows:

MGCIKSKGKDSAPRVSKGEELFTGVVPILVELDGDVNGHKFSVSGEGEGDATYGKLTLKLICTTGKLPVPWPTLVTTLG YGLQCFARYPDHMKQHDFFKSAMPEGYVQERTIFFKDDGNYKTRAEVKFEGDTLVNRIELKGIDFKEDGNILGHKLEY NYNSHNVYITADKQKNGIKANFKIRHNIEDGGVQLADHYQQNTPIGDGPVLLPDNHYLSYQSKLSKDPNEKRDHMVLL EFVTAAGITLGMDELYKPREQIAEFKEAFSLFDKDGDGTITTKELGTVMRSLGQNPTEAELQDMINEVDADGNGTIYFPE FLTMMARKMKDTDSEEEIREAFRVFDKDGNGYISAAELRHVMTNLGEKLTDEEVDEMIREADIDGDGQVNYEEFVQM MTAKSRKRRWKKNFIAVSAANRYKKISSSGALTSTEYKLVVVGAGGVGKSALTIQLIQNHFVDEYDPTIEDSYRKQVVI DASGSAITVQRYVRGIQARAYARFLASGETCLLDILDTAGLEEYSAMRDQYMRTGEGFLCVFAINNTKSFEDIHHYREQI KRVKDSEDVPMVLVGNKCDLPSRTVDTKQAQDLARSYGIPFIETSAKTRQGVDDAFYTLVREIRKHKEKMSKDGKKK KKKSKTK

The amino acids that are highlighted yellow is the Lyn peptide for membrane localization; green is Venus for fluorescent labeling; cyan is CaM; purple is MLCKp; red is Ras; grey is IQ2p; and uncoloured are linker amino acids.

Similar to the design of Raber, IQ2p was embedded between residues 46 and 47 of Ras.

There is Ca2+-independent association between CaM and myosin V (IQ2p) peptide (affinity in the micromolar range). This association between CaM and IQ2p prevents the folding of the

functional Ras protein sequence, as well as Ras interaction with its effector proteins. CaM-

MLCKp was fused to the amino-terminal of Ras and IQ2p. When there is an increase in intracellular calcium, MLCKp acquires a higher affinity for Ca2+-CaM, leading to dissociation of

CaM to IQ2p, and allowing the Ras protein sequence within Raser to fold into its active, functional form.

3.4. Experimental Procedure

3.4.1. Plasmid construction

The transfer vectors CMVp_LynVenusRASer_kb32, CMVp_chr2_Cherry_rtz32,

CMVp_VEGFR2_CMVp_Ceru3NES_rtz32, and CMVp_LynCherry_CMVp_VEGF_kb32 were synthesized by Genscript (Pescataway, NY) and subcloned into the pUC57-Simple vector using

Mini-prep kit (Invitrogen).

3.4.2. Cell lines

Mouse embryonic fibroblast (NIH3T3) cells were maintained in Dulbecco's modified Eagle's medium containing 25 mM D-glucose, 1 mM sodium pyruvate and 4 mM L-glutamine

(Invitrogen), supplemented with 10% fetal bovine serum (Sigma-Aldrich), in T25 flasks (37°C and 5% CO2). All cell lines were authenticated and tested for contamination. Cells were

passaged at 90% confluency using 0.05% TrypLE with Phenol Red (Invitrogen) and seeded onto

24-well plates (Corning) at 1:20 dilution. Cells were transiently transfected using Lipofectamine

2 weeks. Colonies were plated in 96-well tissue culture plates (Sarstedt) for subsequent experiments and imaging.

3.4.3. Cell stimulation

The dot matrix was an 8 × 8 Matrix Dot LEDs MAX 7219 Display Module (White Color) connected to an Arduino uno board. Each LED measured at 2 mm in diameter. The LED matrix was programmed using the Arduino IDE Software. For overnight stimulation using the dot matrix, an LED was programmed to flicker white light for 10 ms flashes every 50 ms for 30 s, with a 30 s rest period of no activity. This constituted one cycle, which was repeated indefinitely to produce a stationary flickering light source.

3.4.4. Live cell imaging

438/24; EM, 482/32), YFP (EX, 500/24; EM, 542/27), RFP (EX, 580/20; EM, 630/60)

(Semrock). Image acquisition and analysis were performed with µManager and ImageJ software,

respectively (112, 113). Microscopic images underwent an image processing method that involved applying a bandpass filter and gaussian blur, followed by thresholding and labelling.

3.4.5. Data analysis

All experiments with n=3 were conducted with at least ten cells evaluated in each trial. All data with normal distribution and similar variance were analyzed for statistical significance using two-tailed, unpaired Student's t-tests, unless otherwise indicated. Multiple group comparisons were made with one-factor ANOVA with Tukey-Kramer post-hoc test, unless otherwise indicated. For all tests, α was set at 0.05. P<0.05 was considered significant. Data are mean ± standard deviation, unless otherwise stated. Data was analyzed using the Data Analysis ToolPak for Excel (Microsoft).

3.5. Results

3.5.1. Ca2+-dependent Focus Formation by Raser

Following infection, a stable cell line of NIH3T3 cells expressing Raser was examined under brightfield and fluorescent microscopy for evidence of transformed morphology. To determine whether cells were transformed and focus formation had occurred, transformed colonies were selected based on being more refractive under brightfield microscopy, high relative fluorescence, and an irregular growth pattern compared to normal fibroblasts. A representative colony of morphologically transformed cells is shown in Figure 3.3.

Figure 3.3. Comparison of normal and transformed morphologies of NIH3T3 cells at 100% confluency. (A) Normal NIH3T3 cells using a 10x objective. (B) Transformed NIH3T3 cells using a 10x objective. (C) Brightfield microscopy of a focus from (B) using a 40x objective. (D) Observation under YFP channel of a focus from (B) using a 40x objective. Raser expressing cells are membrane labeled with LynVenus to verify expression.

Infection and subsequent blastcidin selection achieved a stable NIH3T3 cell line expressing Raser. NIH3T3 cells have endogenous purinoreceptors (161), hence, 10 μM ATP was added to Raser expressing cells to stimulate Raser activity. To encourage cellular transformation, cells were stimulated at 100% confluency in a 96 well plate. In Raser expressing cells, addition

of ATP and overnight incubation resulted in 0.33 ± 0.58 foci (Figure 3.5). Normal NIH3T3 cells did not form foci at 100% confluency.

3.5.2. Controlling Focus Formation via a Physical Stimuli: Rewiring of Raser to

Channelrhodopsin 2 (ChR2)

Raser fusion protein was co-expressed with ChR2 to examine rewiring of Raser to light activation. Raser and ChR2 expressing cells were obtained by infecting NIH3T3 cells with

CMVp_LynVenusRASer_kb32 and CMVp_chr2_Cherry_rtz32 vectors under blastcidin and zeocin selection. To verify expression of Raser and ChR2, cells fluoresce yellow and red along the membrane, respectively (Figure 3.4B). To stimulate Raser and ChR2 expressing cells, a flickering LED light matrix was placed underneath the well. At 100% confluency, Raser and

ChR2 expressing cells were incubated overnight with a LED light source. After 16 hours of light stimulation, 12.33 ± 1.53 foci formed (Figure 3.6).

Figure 3.4. (A) Schematic representation of Raser activity with Ca2+-mobilizing activity from CMVp_chr2_Cherry_rtz32 (ChR2). (B) NIH 3T3 cells expressing Raser and ChR2 after 16 hours LED light stimulation. Imaged under brightfield (left), YFP channel (center) and RFP channel (right) under 4x objective. Scale bar is 200 μm. 3.5.3. Controlling Focus Formation via a Chemical Stimuli: Rewiring of Raser

to VEGFR2

To extend Raser rewiring a chemical stimulus, a stable cell line was obtained by co- infection of the transfer vectors CMVp_LynVenusRASer_kb32 and

CMVp_VEGFR2_CMVp_Ceru3NES_ktz32, followed by selection with blastcidin and zeocin.

Expression of Raser and VEGFR2 were verified by presence of yellow fluorescence along the membrane and cerulean fluorescence in the cytoplasm, respectively (Figure 3.5B). To stimulate

Raser and VEGFR2 expressing cells, supernatant from VEGF-A secreting HEK293 cells was used (from Section 2.5.3), followed by overnight incubation. 2.67 ± 0.58 foci were counted following overnight incubation in VEGF-A supernatant (Figure 3.6).

Figure 3.5. (A) Schematic representation of Raser activity with Ca2+-mobilizing activity from CMVp_VEGFR2_CMVp_Ceru3NES_ktz32 (VEGFR2). (B) NIH 3T3 cell expressing Raser and VEGFR2 with addition of supernatant from HEK 293 cells secreting VEGF-A, and incubation for 24 hours. Cells were cultured in 96 well plates. Imaged under brightfield (left), YFP channel (center), and CFP channel (right) under 10x objective. Scale bar is 30µm.

Figure 3.6. Combined results for foci count in NIH3T3 cells expressing Raser, VEGFR2 and CeruNES following bolus addition of VEGF-A media, and NIH3T3 cells expressing Raser and ChR2 following 16 hours of LED light stimulation. Results are based on 3 trials (n=3) per transfected cell line. Focus count for each cell line are as follows: Raser and ChR2 (11, 12, 14), Raser and VEGFR2 (2, 3, 3), and Raser (0, 0, 1). Data that is not represented in graph are NIH3T3 cells expressing: Raser stimulated by VEGF-A, Raser stimulated with 16 hours of LED light (foci counts were zero). Mean and standard deviation as error bars are represented, α = 0.05. Significance for * and ** are p > 0.001.

3.6. Discussion

The aim of this work was to engineer a Ca2+-activated Ras protein. When constitutively

GTP-bound, active Ras is known for its critical role in cellular transformation, as a result of multiple downstream effectors working cooperatively including Raf-MAPK-ERK, PI3K, RhoA, and Rac1 pathways (162). However, the only occurrence when Ras-dependent transformation occurs is in disease states. Additionally, Ras GTPase in terms of cellular transformation is extremely complex in terms of upstream regulation, with many pathways remaining largely unknown. Ras GTPase proteins are naturally directly controlled by the exchange of GTP and

GDP, mediated by GEFs and GAPs. This work replaced the GTP/GDP switch system for a Ca2+- calmodulin switch system, enabling Ras to become controlled by Ca2+ instead. The ability to

form foci was the chosen output to show that the engineered Ca2+-activated Ras protein, Raser was able to accomplish cellular transformation. With the ability to form foci upon Ca2+ signaling, cells expressing Raser could be applied to protein networks in which it serves to accumulate at target sites.

NIH3T3 cells were cultured until 100% confluency in order to encourage transformation.

Because NIH3T3 cells have endogenous purinoreceptors, ATP was used to stimulate cells expressing Raser. In one of three cases, foci formed for Raser expressing cells. It is recommended in future work to compare the engineered Ras fusion protein to Ras mutants, such as dominant negative Ras:S17N (163), to further support the findings.

To demonstrate rewiring of the engineered Ras fusion protein to a chemical stimulus,

NIH3T3 cells were co-infected with CMVp_LynVenusRASer_kb32 and

CMVp_VEGFR2_CMVp_Ceru3NES_ktz32 to expressRaser and VEGFR2, respectively.

Additionally, another stable cell line of HEK293 cells infected with

CMVp_LynCherry_CMVp_VEGF_kb32 was used to obtain a VEGF-A source. Supernatant from the VEGF-A source cells, followed by overnight incubation was able to stimulate Raser and VEGFR2 expressing cells to form foci. To further demonstrate the ability to directly control focus formation capabilities of Ras, NIH3T3 cells were co-transfected with

CMVp_LynVenusRASer_kb32 and CMVp_chr2_Cherry_rtz32 to express Raser and ChR2, respectively. To stimulate cells, a flickering LED light matrix was placed underneath the plated cells overnight. Interestingly, compared to the cases using ATP and VEGF-A supernatant, the number of foci formed by Raser and ChR2 expressing cells was significantly higher. This may have been due to higher frequency and stimulation of the light stimulus as compared to the other treatments. Thus, this work has been able to demonstrate that Ras GTPase proteins could be

engineered to become directly controlled by calcium signaling. However, for further support of this work, it is recommended to use different Ras mutant cell lines (such as Ras:S17N), or inhibitors for downstream effectors of Ras (e.g. Rac1 inhibitor CAS 1177865-17-6, ROCK inhibitor Y-27632)(164).

Chapter 4. Conclusion and Future Directions

4.1. Conclusion

The goal of this work was to engineer Ca2+-activated fusion proteins from the Rab and

Ras GTPase family proteins. In this work, the GTP/GDP switch system of the original proteins and replaced with a Ca2+-calmodulin system. This involved insertion of the IQ2p peptide within the functional sequence of the protein, in which the CaM sequence binds to in low intracellular

Ca2+ concentrations. When intracellular concentrations increase, Ca2+-CaM associates with

MLCKp instead, resulting in dissociation with IQ2p. Due to release of IQ2p, the functional protein is able to fold properly into its active form.

A Ca2+-activatable Rab5 protein (named Raber) capable of cellular detachment was engineered. The fusion protein was comprised of a constitutively membrane labeling tag, YFP,

CaM, and MLCKp on the amino terminal of the Rab5 protein. Between amino acids 62 and 63,

IQ2p sequence was embedded. Upon stimulation, Raber expressing cells exhibited cell actin reorganization by protrusion and edge retraction. Cells co-expressing Raber and ChR2 resulted in protrusion and edge retraction following light stimulation. Similarly, cells co-expressing

Raber and VEGFR2 that were stimulated by VEGF-A cell culture supernatant resulted in retraction.

A Ca2+-activatable Ras protein (named Raser) was also engineered. The fusion protein was comprised of a constitutively membrane labeling tag, YFP, CaM, and MLCKp on the amino terminal of the Ras protein. Between amino acids 46 and 47, IQ2p sequence was embedded.

Cells co-expressing Raser and ChR2 were subjected to overnight stimulation of light, and

resulted in focus formation. Likewise, cells co-expressing Raser and VEGFR2 resulted in focus formation following overnight incubation with VEGF-A cell supernatant.

In regards to the research objectives outlined in Chapter 1, this work addresses the two objectives: (1) a Ca2+-activatable Rab5 protein was engineered to induce cellular actin reorganization via protrusion retraction, and (2) similarly, a Ca2+-activatable Ras protein was engineered to induce focus formation in NIH3T3 cells upon stimulation. In both cases for the engineered Rab5 and Ras fusion proteins, their activity was activated under specific conditions

(i.e. light via ChR2 and VEGF-A via VEGFR2) to modulate its behaviour. Meeting these research objectives illustrates that the strategy of replacing the GTP/GDP switch system with

Ca2+-calmodulin can be further applied to other Ras superfamily GTPases. It is important to note that, this work could be further supported by the use of Rab5 and Ras mutants, as well as chemical inhibitors as controls.

4.2. Future Directions

In the context of cell-based therapeutics, these engineered Rab5 and Ras fusion proteins could be incorporated in synthetic Ca2+ signaling networks used to reprogram cells. The ability of Raber to enable cellular reorganization by detachment of protrusion and cell edge could have potential uses in enabling migratory behaviour. Raber could be coupled with CaRQ, an engineered Ca2+-activated RhoA protein that enables cells to adopt a migratory morphology via blebbing (58). Biomarkers could serve as migratory cues, enabling reprogrammed cells to respond proportionally to concentration of the biomarker. To add to the combination of Raber and CaRQ, Raser, with its ability to form foci, could serve to accumulate reprogrammed cells as they approach the targeted disease site. This work has demonstrated the versatility of these Ca2+-

activated proteins via ChR2 and VEGFR2. With that being said, rewiring these fusion proteins to the broad library of existing engineered chimeric receptors would allow control of Raber and

Raser functions to nearly any extracellular stimuli.

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Appendix

Appendix A.

The content from this section has been modified from:

Wong J., Qudrat A., Mosabbir A., Truong K. Advances in senotherapies. To appear as a book chapter in Molecular basis and emerging strategies for anti-aging interventions.

Introduction Senescence is defined as the gradual deterioration of function leading to increased mortality from disease and injury, also referred to as “biological aging”. Although the concept of death may seem as an inevitability, reducing the rate of senescence has shown the possibility of delaying death or even reduced to a negligible factor. For example, as early as 1934, rats undergoing dietary restriction have shown to increase their lifespan by 14-45% (Swindell, 2012). Alternatively, some fish, turtles, and invertebrates achieve a state of “negligible senescence”, which occurs when there is no measurable decline in function or reproductive capabilities with age (Guerin, 2004). In humans, changes to senescence come in the form of diseases that hijack natural senescent processes, such as the rare accelerated aging syndrome (Dreesen and Stewart, 2011) or “Syndrome X”, a disease in which a person remains physically and mentally an infant throughout their life (Walker et al., 2015). Since age is the major risk factor for prevalent diseases in the developed world (i.e. cancer, cardiovascular disease, neurodegeneration etc.), as well as the fact that the aging process can be hijacked, the possibility of reversing the aging process through manipulating cellular senescence pathways has been key research interest. Cellular senescence, which leads to organismal senescence, specifically refers to the irreversible arrest of cell proliferation that starts due to stressful stimuli (Campisi and d'Adda di Fagagna, 2007). These can include DNA damage, dysfunctional telomeres, disrupted chromatin or oncogenes (Campisi et al., 2001, Serrano and Blasco, 2001). Not only are senescent cells irreversibly arrested, but they also secrete an assortment of cytokines, chemokines, growth factors, and proteinases that are collectively termed the senescence-associated secretory phenotype (SASP) (Campisi, 2013). The SASP act as paracrine signals that produce a myriad of effects depending on the physiological context. With age, the number of senescent cells increases and consequently leads to an accumulation of SASP cytokines and proteins which in turn can accelerate pathology. This condition in cells is thought to be irreversible due to the absence of any known physiological conditions that can reverse senescence. However, biological manipulations targeting cellular senescence pathways have shown a regeneration of proliferative function. For example, inactivation of the p53 gene in senescent fibroblasts caused a return to robust growth (Beausejour et al., 2003). This finding is one of many that attempt to uncover

therapeutic strategies for age-related diseases by selectively eliminating the disease-causing features of senescent cells, collectively termed senotherapies. Senolytics are a class of senotherapies that uses molecular compounds to selectively and efficiently induce cell death in senescent cells. For example, Baker et al. (2011) showed that the removal of p16(Ink4a)-positive senescent cells in mice delayed age-associated diseases such as osteoporosis. In this chapter, we review the literature contributing to the development of senolytic strategies, specifically therapeutic agents that target cellular senescence pathways. Senolytics targeting pro-survival networks Cellular senescence is described as having a double edged influence on cellular proliferation. On one hand, senescence can be seen as an anti-cancer response that turns potentially cancerous cells into benign tumours. Conversely, senescence of healthy cells or large amounts of cells can reduce the ability for tissue to regenerate and repair itself. Thus, most senolytic agents are currently being developed to target pro-survival networks due to the observation that cellular senescence is preceded by some form of tumorigenic stress such as DNA damage. Senescent cells, although harboring DNA damage and being immersed in local SASP, have the remarkable ability to withstand stress. Common markers of cellular senescence include decreased cellular proliferation, and increased cell size and volume (Fuhrmann- Stroissnigg et al., 2017). Senescent cells also tend to have increased expression of cell cycle inhibitors p21(Cip1) and p16(Ink4a), and the SASP factor interleukin-6 (IL-6) (Fuhrmann- Stroissnigg et al., 2017). Consequently, it is hypothesized that they have upregulated pro- survival/anti-apoptotic networks. Senolytics targeting pro-survival networks have shown efficacy against atherosclerosis, osteoporosis, cancer, and other age-related disorders (Campisi and d'Adda di Fagagna, 2007). The major pro-survival/anti-apoptotic pathways to be discussed in this chapter include p53/p21, Bcl-2/Bcl-xL, PI3K/Akt, and serpine pathways (Figure 1).

Figure 1. The p53/p21, Bcl-2, and PI3K/Akt pathways all have regulatory roles in cell survival. Although it is known that PAI-1 upregulates p53/p21 activity, the mechanism of the serpine pathway still remains unknown. p53/p21 pathway: FOXO4-interacting peptide Permanent growth arrest is initiated with the p53/p21 pathway. Activated p53 leads to the induction of p21, which in turn inhibits the cyclin/cyclin dependent kinase complexes involved in cell cycle progression (Harris and Levine, 2005). Activation of p53 also leads to cell death as p53 is translocated to the mitochondria, which is important in the release of cytochrome c and protease activation (Harris and Levine, 2005). However, in senescent cells, while cyclin/cyclin dependent kinase activity is inhibited, cells do not undergo apoptosis. In a study with human IMR90 fibroblast cells, despite elevated levels of pro-apoptotic initiators and reduced levels of anti-apoptotic factors, cells resisted death (Baar et al., 2017). The group also observed that senescent cells had elevated mRNA and protein expression of FOXO4, a protein that had not been previously linked to senescent cell death. Interestingly, FOXO4 was able to induce IMR90 cells to senesce rather than apoptose, despite the high levels of pro-apoptotic and low levels of anti-apoptotic factors priming the cells. FOXO4’s mechanism involves its association with p53 in the nucleus, preventing nuclear exclusion and p53-mediated apoptosis. By inhibiting FOXO4, there was release of cytochrome c

and caspase activity in pre-senescent cells, and decreased cell viability and density in senescent cell cultures. In response, Baar et al. (2017), aimed to disrupt the FOXO4-p53 interaction, designing a D-retro inverso (DRI)-modified peptide containing part of the p53-interaction region found in FOXO4 which they called FOXO4-DRI. DRI-modified peptides can increase peptide potency in vitro and in vivo (Borsello et al., 2003). FOXO4-DRI was capable of binding to p53 with higher affinity than FOXO4, enabling p53 nuclear exclusion and translocating it to the mitochondria to induce p53-mediated apoptosis. In fast-aging mice, FOXO4-DRI treatment reduced the effects of doxorubicin-induced senescence, counteracted hair loss, improved renal function, and improved fitness such as increased voluntary running wheel activity. Bcl-2/Bcl-xL pathway inhibitors The B-cell lymphoma-2 (Bcl-2)-related family constitutes important apoptosis-regulatory genes that usually act on the mitochondrial and nuclear membrane, and endoplasmic reticulum due to a carboxy-terminal transmembrane (TM) region limiting their subcellular distribution (Muchmore et al., 1996, Wang et al., 2001). While most Bcl-2 proteins are death-inhibiting (e.g. Bcl-2, Bcl-xL, Bcl-w, Mcl-1, Bfl1/A-1, Bcl-B), containing all four Bcl-2 homology domains; there are also Bcl-2 homologs that comprise of death-inducers, subdivided into proteins containing Bcl-2 homology 1-3 domains (e.g. Bax, Bak, Bok) (Wolter et al., 1997, Chittenden et al., 1995, Hsu et al., 1997), and proteins containing only the BH3 domain (e.g. Bid, Bim, Bad) (Oltersdorf et al., 2005, O'Connor et al., 1998, Yang et al., 1995). Notably, the ratio of pro- and anti-apoptotic Bcl-2 family proteins influences the fate of a cell. When the expression of anti- apoptotic Bcl-2 proteins overwhelms the levels of pro-apoptotic Bcl-2 proteins, the cell is able to escape apoptosis, thus resisting drugs and therapeutic agents (Del Poeta et al., 2003, Minn et al., 1995). Constitutively high levels of the pro-survival Bcl-2 proteins have been associated with aggressive malignancies, drug resistance towards chemotherapeutic agents, and senescent cells (Reed, 2008, Davis et al., 2003). Hence, there has been a significant effort in targeting Bcl-2 family proteins with senolytic agents, such as TW-37, which is a nonpeptide Bcl-2 inhibitor (Zhu et al., 2016); Navitoclax (ABT-263), which has shown preferential elimination of senescent cells by inducing apoptosis via caspase 3 and 7 activity (Zhu et al., 2016); and ABT-737, a Bcl-2/Bcl- w/Bcl-xL inhibitor, which has shown in vivo preferential elimination of senescent cells and increased hair-follicle stem cell proliferation in the epidermis (Yosef et al., 2016). However, the effectiveness of Bcl-2 inhibitors is cell-type dependent. For instance, Navitoclax (ABT-263) has shown to selectively induce apoptosis in radiation-induced senescent human umbilical vein endothelial cells (HUVECs) and IMR90 cells, whereas TW-37 has no senolytic activity in these cell types (Zhu et al., 2017). Another issue posed by inhibitors like ABT-263 and ABT-737 is their cause of severe thrombocytopenia (Schoenwaelder et al., 2011, Schoenwaelder and Jackson, 2012). PI3K/Akt pathway: HSP90 inhibitors, fisetin

Phosphatidylinositol-3 kinases, PI3Ks, are lipid kinases capable of phosphorylating inositol ring 3’-OH group found in inositol phospholipids (Fruman et al., 1998). Through one of their SH2 domains in the adaptor subunit, PI3Ks are recruited to the membrane, binding to phosphotyrosine residues on growth factor receptors/adaptor proteins. Activation of PI3K leads to the conversion of phosphatidylinositol-4,5-bisphosphate (PIP2) to the second messenger phosphatidylinositol-3,4,5-trisphosphate (PIP3), in which PIP3 recruits signalling proteins with the pleckstrin homology (PH) domains to the inner membrane such as PDK1 and Akt (Pawson and Nash, 2000). PDK1 (3′-phosphoinositide-dependent kinase 1) is thought to be a constitutively active protein that phosphorylates Akt at T308, enabling stabilization of phosphorylated Akt (p-Akt) (Alessi et al., 1996). Through several mechanisms, active p-Akt activates and inhibits several substrates involved in regulating cell survival, cell cycle progression and growth (Fresno Vara et al., 2004). The PI3K/Akt pathway has been associated with inducing an apoptosis-resistant phenotype and senescence in several cell types (Lorenzini et al., 2002, Astle et al., 2012). It has been suggested that heat shock protein 90 (HSP90) binds to p-Akt and apoptosis signaling regulating kinase 1 (ASK1) which stabilize p-Akt, encouraging cellular survival and senescence. This binding prevents ASK1 from forming an interaction with p38 to induce signalling for apoptosis (Watanabe et al., 2015, Zhang et al., 2005). It has also been suggested that HSP90 and Akt need to function together in order to inhibit ASK1-p38 signalling. With that being said, disruption of the HSP90-Akt interaction would lead to destabilization of active/phosphorylated Akt and subsequent apoptosis. Several HSP90 inhibitors have been identified as potential senolytics including tanespimycin (17-AAG), geldanamycin, 17-AAD, and 17-DMAG (Fuhrmann-Stroissnigg et al., 2017). The group considered chemical compounds to have senolytic potential if they significantly reduced senescent cells. In the same study, 17-DMAG was able to downregulate the level of p-Akt in senescent Ercc1−/− mouse embryonic fibroblast (MEF) cells in vitro, while another HSP90 inhibitor, namely ganetespib, showed senolytic activity specifically in HUVECs. This illustrates that not all HSP90 inhibitors work in a similar fashion on all cell types. Using a human progeroid syndrome mice model, Fuhrmann-Stroissnigg et al. (2017) found that 17-DMAG extended healthspan by assessing reduction in age-related symptoms such as kyphosis, dystonia, tremor, loss of forelimb grip strength, coat condition, ataxia, gait disorder, and body condition. Another chemical that functions through the PI3K/Akt pathway specifically is a naturally occurring flavone called fisetin (3,3’,4’,7-tetrahydroxyflavone), which is found in high concentrations in strawberries (160 µg/g) (Khan et al., 2013). Fisetin is a hydrophobic molecule that accumulates in cells, and has shown selective apoptosis induction of human breast cancer MCF-7 cells via caspases 7, 8, and 9 (Yang et al., 2012). In both in vitro and in vivo studies, fisetin demonstrated senolytic activity in HUVECs, but not IMR90 and primary human preadipocytes, as shown by caspase 3 and 7 activity assays (Zhu et al., 2017). Fisetin is a widely available nutritional supplement and has very little known side effects, demonstrating its potential to act as an orally-administered senolytic agent (Zhu et al., 2017).

Serpine pathway inhibitors Serpine genes encode for serine protease inhibitor (serpin) superfamily proteins that include serpin B2 (PAI-2), serpine E1 (PAI-1), and serpine E2 (Potempa et al., 1994, Kortlever et al., 2006). Among these, there has been considerable interest in plasminogen activator inhibitor 1 (PAI-1) as it has been identified as a senescence-associated gene given its increased expression in senescent cells (Kortlever et al., 2006, Suzuki et al., 2001, Dimri et al., 2000, Elzi et al., 2012). There has been increasing evidence to support that PAI-1 is not only a biomarker but also a mediator of cellular senescence (Perez et al., 2010, Eren et al., 2014a, Eren et al., 2014b, Ghosh et al., 2016). In rat idiopathic pulmonary fibrosis (IPF) alveolar type II (ATII) cells, treatment with bleomycin caused an increase in PAI-1, as well as other senescence biomarkers p53, p21, and senescence associated beta-galactosidase (SA-β-gal) (Jiang et al., 2017). On the other hand, silencing PAI-1 using siRNA reduced p53 and p21 expressions (Jiang et al., 2017). This suggests that PAI-1 positively regulates p53 and p21 levels in the p53-p21 pathway. However, the mechanism PAI-1 uses to regulate p53 activity is still unknown. Screening for senolytic agents β-galactosidase assay The β-galactosidase assay is a widely used screening platform to identify senotherapeutic drugs and combinations in vitro and in vivo due to its simplistic method and apparent specificity towards senescent cells (Krishnamurthy et al., 2004, Cao et al., 2003, Castro et al., 2003, Itahana et al., 2007). The assay measures the expression levels of senescence-associated β-galactosidase activity (SA-β-gal) which is expressed predominantly by senescent cells, occurring at higher frequency in older tissues (Dimri et al., 1995). SA-β-gal is detectable by colorimetric X-gal (5- bromo-4-chloro-3-indolyl-β-D-galactopyranoside) staining at pH 6.0 and/or by using the fluorescent substrate C12FDG (5-dodecanoylaminofluorescein-di-b-D-galactopyranoside) staining (Dimri et al., 1995). There are, however, a few criticisms of using SA-β-gal as a surrogate marker of senescent cells. It is important to consider that SA-β-gal has also been highly expressed in non-senescent states (Severino et al., 2000, Untergasser et al., 2003) and in confluent cultures maintained for prolonged periods in vitro (Dimri et al., 1995). Additionally, despite mRNA knockdown of the gene GLB1, which codes for SA-β-gal expression, cells still entered senescence (Lee et al., 2006). This suggests that SA-β-gal is not required for senescence. Hence, measuring SA-β-gal is sometimes coupled with measuring other biomarkers of senescence such as p16 gene products and the SASP inflammatory cytokine IL-6 (Kuilman et al., 2008, Capparelli et al., 2012, Marcoux et al., 2013). Animals models for senescent studies: Engineered by Demaria et al. (2014), the p16-3MR mouse model allows observation and manipulation of senescent cells in vivo. Using their p16-3MR model, the group demonstrated that although senescent cells have mostly inflammatory and detrimental effects through SASP,

the total elimination of senescent cells can hinder wound healing and tissue differentiation (Demaria et al., 2014). The p16-3MR mouse strain expresses a trimodal reporter protein (3MR) that is under control by the p16(Ink4a) promoter, and contains the functional domains of a synthetic Renilla luciferase (LUC), monomeric red fluorescent protein (mRFP), and truncated herpes simplex virus 1 (HSV-1) thymidine kinase (HSV-TK) (Demaria et al., 2014, Ray et al., 2004). Senescent cells, both in vivo and in vitro, often express p16(Ink4a), a cyclin-dependent kinase inhibitor that is also known as CDKN2A. Expression of p16(Ink4a) causes the growth arrest associated with irreversible senescence (Coppe et al., 2011, Baker et al., 2011). Hence, the amount of p16 gene products and the SASP (the most common cytokine being IL-6) would reflect the number of senescent cells. Cells were also engineered to be fluorescent via mRFP to allow distinguishability of senescent cells from tissue samples and to be bioluminescent via LUC to allow traceability in vivo (Chang et al., 2016). HSV-TK converts ganciclovir prodrug into a toxic form that subsequently induces apoptosis (Laberge et al., 2013, Gao et al., 1999). Assessing current senolytic agents Developing drugs for targeted senolytics is currently being researched as a viable treatment modality to alleviate disease symptoms. Markers of drug effectiveness include expression levels of survival gene networks together with apoptotic resistance. Small molecule drugs would significantly inform current medicinal approaches aimed to relieve the harmful effects of aging diseases as senolytics drugs have the capacity to selectively target and kill senescent cells. Common target genes converge through various pathways to cell cycle inhibitors such as p21, which in turn inhibits cyclins/cyclin dependent kinases eventually leading to senescence. One key pathway that current tested drugs target for senescence are the ephrin dependent receptor ligands, called EFNB1 or EFNB3 (Hwang et al., 2018). These are the largest receptor tyrosine kinases and coordinate cell survival during development. Like other ligand-receptor interactions, the Ephrin receptors can interact with like receptors on adjacent cells to stimulate downstream cell signalling. Specifically, EFNB3 has been known to induce the SASP when the gene is overexpressed (Hwang et al., 2018). Table 1 highlights the pro-survival pathways together with their targeted senolytics for reference. In vitro testing of drugs targets gene products that protect senescent cells. Both Dasatinib and Quercetin are two drugs that are known to clear senescent cells (Hwang et al., 2018). The drug, Dasatinib, is an inhibitor of multiple tyrosine kinases which was originally used for treating cancers. Specifically, it is known to inhibit the suppression of apoptosis in human fat cell progenitors. Similarly, Quercetin inhibits another class of kinases called PI3K and serpines. This drug was particularly effective against heart and umbilical vein endothelial cells (HUVEC). This presents evidence for cell specific targeted therapy. In addition, combinations of both drugs showed the selective killing of both senescent fat cell progenitors as well as HUVECs. Post in vitro testing, the drugs were tested in reducing the viability of senescent murine cells. The murine cells specifically tested were mouse embryonic fibroblasts (MEFs) which

showed a significant reduction in number post treatment with both Dasatinib and Quercetin (Demaria et al., 2014). In these mice, when the cardiovascular function was tested, there was substantial impairment in vascular reactivity seen in aged mice. This further uncovered a link between senescent cells and cardiovascular dysfunction in humans. However, not all formulations were successful in drug testing. For example, a selective pro-drug in targeting senescent cells is the quercetin derivative, quercetin 3D galactoside (Q3G; hyperoside). Hyperoside is a natural derivative of quercetin and structurally identical except that it contains a cleavable galactoside group. However, it was found to be ineffective in targeting senescent endothelial cells in vitro (Hwang et al., 2018). Navitoclax, a Bcl-2 inhibitor, was similar in action to Quercetin and Dasatinib, and also eliminated cells via apoptosis in similar human and mouse cell types (Zhu et al., 2017). In addition to targeting multiple Bcl-2 family target proteins, Navitoclax acted non-specifically. To illustrate, administration of Navitoclax in mice led to the effective depletion of senescent bone marrow hematopoietic stem cells (HSCs) as well as senescent muscle stem cells (MuSCs) (Chang et al., 2016). Similarly, Bcl-xL inhibitors, namely A1331852 and A1155463 were found to be senolytic in HUVECs and IMR90 cells, but not pre-adipoctypes (Zhu et al., 2017). This activity occurred through apoptosis as tested by caspase 3/7 activity in vitro. Piperlongumine is a natural product, isolated from the genus Piper and demonstrated to have senolytic properties (Wang et al., 2016). Piperlongumine was shown to selectively kill human WI-38 fibroblasts by several means: reducing viability in IR-induced as well as Ras- induced WI-38 senescent cells. It was uncovered that the selective killing occurred through apoptosis, through a reactive-oxygen species (ROS)-independent mechanism. Further, a synergistic effect of Piperlongumine was seen when administered together with Navitoclax. Other drugs on the market such as metformin, rapamycin and ruxolitinib have shown promising effects to suppress SASP, specifically alleviating symptoms of age-related disorders causing metabolic dysfunction (Huffman et al., 2016). For example, ruxolitinib, a JAK 1/2 inhibitor, alleviates insulin resistance and tissue dysfunction (Xu et al., 2015). The intermittent administration of these drugs have shown to mitigate effects of cellular senescence. However, since the specific mechanism of action remain unknown, it is difficult to identify exactly how these SASP inhibitors function. Moreover, administering them in conjunction with the pathway- specific drugs described above may prove a difficult, if not impossible task. Table 1. Mapping drugs to potential targeted pathways

Potential Targeted senolytics References pathway p53/p21 Quercetin, Dasatinib + Quercetin, (Hwang et al., 2018, Wang et al., Piperlongumine, FOXO4-related peptide 2016, Baar et al., 2017)

Bcl-2/Bcl-xL Quercetin, Navitoclax (ABT-263), ABT- (Hwang et al., 2018, Wang et al., 737, Piperlongumine, A1331852, 2016, Baar et al., 2017, Zhu et al., A1155463 2017)

PI3K/Akt Quercetin, Fisetin, Geldanamycin, (Hwang et al., 2018, Khan et al., Tanespimycin, Ganetespib 2013, Fuhrmann-Stroissnigg et al., 2017)

Serpine Quercetin, Dasatinib + Quercetin (Hwang et al., 2018)

Future direction of senolytics Although the potential to translate senolytics into clinical treatments shows promise, there are some concerns moving forward. Perhaps the biggest challenge lies in the fact that cellular senescence has a dual nature, and creating anti-aging treatments may not be as simple as accentuating or attenuating core senescence pathways. The inhibition of growth for example, acts as a natural anti-cancer mechanism that prevents tumours from progressing past benign stages. Given that tumorigenic effects largely begin by mutations in the genome, removing the senescence capabilities of rapidly dividing cells altogether would simultaneously increase the chances of developing tumours. This is a major risk with regards to “off-target” effects of senolytic drugs. On the other hand, the benefits of removing senescent cells in animal models have clearly shown promise. For example, the removal of p16(Ink4a)-positive senescent cells in mice delays age-associated diseases such as osteoporosis (Baker et al., 2011). One strategy to approach dealing with issues regarding the duality of senescent effects is to engineer therapies to target specific cellular contexts. A recent innovative approach involved using engineered proteins expressed in cells to target senescent cells releasing IL-6 cytokines (Figure 2) (Qudrat et al., 2017).

Figure 2. Concept of IL-6 targeting cells responding and migrating towards IL-6 sources (e.g. senescent cells in SASP). Understanding the specific contexts of harmful senescence states and targeting therapeutics to that context may provide a better approach to selectively remove harmful senescent cells. This cellular-based approach to senotherapies involves the creation of a stable cell line expressing a synthetic chimeric receptor for a SASP cytokine, and calcium activated RhoA (CarQ) to enable migration towards SASP sources (Qudrat et al., 2017). In Qudrat et al., (2017), IL-6 targeting cells were engineered to express a chimeric receptor called IL6Rchi, which is comprised of the extracellular portion of the IL-6 receptor to bind to the SASP cytokine IL-6, and the transmembrane and cytoplasmic domains of VEGFR2 to generate calcium signalling. The calcium signals generated by IL-6 binding to IL6Rchi activates the activity of calcium activated RhoA (CarQ), enabling the IL-6 targeting cell to migrate towards areas/sources of high IL-6 expression (i.e. senescent cells expressing the SASP). Once at the targeted SASP site, the HSV-TK system can be used to convert ganciclovir into its toxic form to induce apoptosis (Gao Ding et al., 1999, Qudrat et al., 2017). The results have been promising in vitro in targeting directed SASP engineered cells. Although the potential to translate senolytics into clinical treatments is present, there are clear obstacles. For one, the main difficulty is in determining potential endpoints in clinical trials (Kirkland and Tchkonia, 2017). Further, treatments that appear effective in mice may prove altogether ineffective in humans. This is particularly true when attempting to translate from genetically-induced mice models to humans. And since not all senescent cells are harmful as they play an important role in wound healing and tissue repair, the need for targeted therapeutics is ever present to eliminate harmful and maintain beneficial effects of cellular senescence. On the contrary, generating targeted therapeutics to remove senescent cells may have decreased rates of drug resistance and recurrence since these cells no longer divide (Kirkland and Tchkonia, 2017). Looking ahead, there are many questions that still need to be addressed before senolytics find their way to the market. First, there is a need for characterizing potential drug side effects, in addition to delays in wound healing, as well as off-target effects (Demaria et al., 2014). Secondly, to optimize the frequency of drug administration, rates of senescent cell re- accumulation need to be explored. Thirdly, the additive and synergistic effects of drugs, specifically in conjunction with SASP inhibitors, need to be uncovered. Fourthly, definitive studies need to be performed to expound the realized effect of senolytics on lifespan. Lastly, a multiplex approach needs to be evaluated coupling drug delivery and cell-based therapeutics to maximize proficiency. Conclusion Age is a condition associated with decreasing physiological and psychological capabilities in humans. This deterioration gives rise to vulnerability to a host of diseases as well as a lower quality way of life. Over the past century, decreased mortality rates and increased life expectancy has been a major advancement of the human condition, followed closely however by a domination of age-related diseases. The discovery of cellular senescence pathways and their

link to aging, as well as the development of animal models that show little signs of accelerated aging have been a major step to a potential treatment for aging conditions. Understanding that cellular senescence can be a beneficial mechanism for cancer protection has also deepened our understanding of the benefits of aging and the complexity of human physiology. Senolytic therapies have attempted to reduce age-related pathologies by the removal of harmful senescent cells. Senolytics targeting pro-survival networks have shown efficacy against atherosclerosis, osteoporosis, cancer, and other age-related disorders in cell and animal based studies. Developing targeted approaches to remove harmful senescence while maintaining the body’s natural defence against uncontrolled proliferation may be the next major challenge. Greater testing in clinical trials and well defined outcomes are also needed but nevertheless, the current outlook in senolytics seems bright. The past century has seen a decrease in mortality rates and increased life expectancy, and although more research is needed, significant steps are being made towards understanding the complexity of aging to move forward into the next century.

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Appendix B.

Arduino IDE Software sketch for 8 x 8 LED Dot Matrix (Arduino uno board)

//We always have to include the library #include "LedControl.h"

/* Now we need a LedControl to work with. ***** These pin numbers will probably not work with your hardware ***** pin 12 is connected to the DataIn pin 11 is connected to the CLK pin 10 is connected to LOAD We have only a single MAX72XX. */ //LedControl lc=LedControl(12,11,10,1); LedControl lc=LedControl(8,10,9,1);

/* we always wait a bit between updates of the display */ unsigned long delaytime=100; void setup() { /* The MAX72XX is in power-saving mode on startup, we have to do a wakeup call */ lc.shutdown(0,false); /* Set the brightness to a medium values */ //lc.setIntensity(0,8); lc.setIntensity(0,15); /* and clear the display */ lc.clearDisplay(0); }

/* This method will display the characters for the word "Arduino" one after the other on the matrix. (you need at least 5x7 leds to see the whole chars) */ void writeArduinoOnMatrix() { /* here is the data for the characters */ byte a[5]={B01111110,B10001000,B10001000,B10001000,B01111110}; byte r[5]={B00111110,B00010000,B00100000,B00100000,B00010000}; byte d[5]={B00011100,B00100010,B00100010,B00010010,B11111110}; byte u[5]={B00111100,B00000010,B00000010,B00000100,B00111110}; byte i[5]={B00000000,B00100010,B10111110,B00000010,B00000000}; byte n[5]={B00111110,B00010000,B00100000,B00100000,B00011110}; byte o[5]={B00011100,B00100010,B00100010,B00100010,B00011100};

/* now display them one by one with a small delay */

lc.setRow(0,0,a[0]); lc.setRow(0,1,a[1]); lc.setRow(0,2,a[2]); lc.setRow(0,3,a[3]); lc.setRow(0,4,a[4]); delay(delaytime); lc.setRow(0,0,r[0]); lc.setRow(0,1,r[1]); lc.setRow(0,2,r[2]); lc.setRow(0,3,r[3]); lc.setRow(0,4,r[4]); delay(delaytime); lc.setRow(0,0,d[0]); lc.setRow(0,1,d[1]); lc.setRow(0,2,d[2]); lc.setRow(0,3,d[3]); lc.setRow(0,4,d[4]); delay(delaytime); lc.setRow(0,0,u[0]); lc.setRow(0,1,u[1]); lc.setRow(0,2,u[2]); lc.setRow(0,3,u[3]); lc.setRow(0,4,u[4]); delay(delaytime); lc.setRow(0,0,i[0]); lc.setRow(0,1,i[1]); lc.setRow(0,2,i[2]); lc.setRow(0,3,i[3]); lc.setRow(0,4,i[4]); delay(delaytime); lc.setRow(0,0,n[0]); lc.setRow(0,1,n[1]); lc.setRow(0,2,n[2]); lc.setRow(0,3,n[3]); lc.setRow(0,4,n[4]); delay(delaytime); lc.setRow(0,0,o[0]); lc.setRow(0,1,o[1]); lc.setRow(0,2,o[2]); lc.setRow(0,3,o[3]); lc.setRow(0,4,o[4]); delay(delaytime); lc.setRow(0,0,0); lc.setRow(0,1,0); lc.setRow(0,2,0); lc.setRow(0,3,0); lc.setRow(0,4,0); delay(delaytime); }

/* This function lights up a some Leds in a row.

The pattern will be repeated on every row. The pattern will blink along with the row-number. row number 4 (index==3) will blink 4 times etc. */ void rows() { for(int row=0;row<8;row++) { delay(delaytime); lc.setRow(0,row,B10100000); delay(delaytime); lc.setRow(0,row,(byte)0); for(int i=0;i

/* This function lights up a some Leds in a column. The pattern will be repeated on every column. The pattern will blink along with the column-number. column number 4 (index==3) will blink 4 times etc. */ void columns() { for(int col=0;col<8;col++) { delay(delaytime); lc.setColumn(0,col,B10100000); delay(delaytime); lc.setColumn(0,col,(byte)0); for(int i=0;i

/* This function will light up every Led on the matrix. The led will blink along with the row-number. row number 4 (index==3) will blink 4 times etc. */ void single() { for(int row=0;row<8;row++) { for(int col=0;col<8;col++) { delay(delaytime); lc.setLed(0,row,col,true); delay(delaytime); for(int i=0;i

lc.setLed(0,row,col,false); delay(delaytime); lc.setLed(0,row,col,true); delay(delaytime); } } } }

/* This function lights up a some Leds in a column. The pattern will be repeated on every column. The pattern will blink along with the column-number. column number 4 (index==3) will blink 4 times etc. */ void movingDot(int blinkPeriod, int blinkDuration, int blinkRepeat) { for(int col=0;col<8;col++) { /* delay(100); // Shift delay. Just make it fast lc.setColumn(0,col,B00100000); delay(100); // Shift delay lc.setColumn(0,col,(byte)0); */ for(int i=0;i

delay(recoveryTime); restCount = restCount + 1;

if ((restCount % skipPeriodAfterRests) == 0) { col = col +1; } } }

void wholeBlink(int blinkPeriod, int blinkDuration) { for(int col=0;col<8;col++) { lc.setColumn(0,col,B11111111); } delay(blinkDuration); // How many secs on? for(int col=0;col<8;col++) { lc.setColumn(0,col,(byte)0); } delay(blinkPeriod); // Period of blink } void dotBlink(int blinkPeriod, int blinkDuration) { for(int col=0;col<8;col++) { lc.setColumn(0,col,(byte)0); } lc.setColumn(0,4,B00010000); delay(blinkDuration); // How many secs on? for(int col=0;col<8;col++) { lc.setColumn(0,col,(byte)0); } delay(blinkPeriod); // Period of blink } void dotBlinkWait(int blinkPeriod, int blinkDuration, int blinkRepeat, int recoveryTime) { for(int i=0;i

delay(recoveryTime); }

void imageBlink(int blinkPeriod, int blinkDuration, int blinkRepeat) {

for(int i=0;i

for(int col=0;col<8;col++) { lc.setColumn(0,col,(byte)0); } delay(blinkPeriod); // Period of blink }

for(int i=0;i

lc.setColumn(0,0,B00000000); lc.setColumn(0,1,B01111110); lc.setColumn(0,2,B01000010); lc.setColumn(0,3,B01000010); lc.setColumn(0,4,B01000010); lc.setColumn(0,5,B01000010); lc.setColumn(0,6,B01111110); lc.setColumn(0,7,B00000000); delay(blinkDuration); // How many secs on?

for(int col=0;col<8;col++) { lc.setColumn(0,col,(byte)0); } delay(blinkPeriod); // Period of blink }

} void loop() { //writeArduinoOnMatrix(); //rows(); //columns(); //single(); //movingDot(3000,200,1200); //movingDotWait(50,10,20,3000,1); //period, exposure, repeats, wait b4 next cycle, skipPeriodAfterRests wholeBlink(5000,200); //dotBlink(3000,200); //dotBlinkWait(50,10,150,30000); //imageBlink(1000,100,5); } 84