A Master's Thesis Entitled Gβγ Mediated

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A Master’s Thesis

entitled

Gβγ mediated calcium mobilization and subsequent calcium-calmodulin (CaM)

signaling in the trailing edge retraction during cell migration

Praneeth Siripurapu

Submitted to the Graduate Faculty as partial fulfillment of the requirements for

The Master of Science Degree in Chemistry

______Dr. Ajith Karunarathne, Committee Chair

______Dr. Jon R. Kirchhoff, Committee Member

______Dr. Donald R. Ronning, Committee Member

______Dr. Amanda Bryant-Friedrich, Dean College of Graduate Studies

The University of Toledo

May 2017

This document is copyrighted material. Under copyright law, no parts of this document may be reproduced without the expressed permission of the author.

An abstract of

Gβγ mediated calcium release and subsequent calcium- calmodulin (CaM) signaling in the trailing edge retraction during cell migration

Praneeth Siripurapu

Submitted to the Graduate Faculty as partial fulfillment of the requirements for The Master of Science Degree in Chemistry

The University of Toledo

May 2017

Cell migration is a multi-step process, which plays a vital role in embryonic morphogenesis and contributes to tissue repair and regeneration. Cell migration starts with the formation of leading edge (LE) and retraction of trailing edge (TE). The LE activities include polymerization of actin and formation of nascent focal adhesions generating traction forces. Simultaneously, disassembly of cytoskeleton and detachment of mature focal adhesions supported by actomyosin contractility can be seen at the TE. G protein coupled receptors (GPCRs) have been shown to regulate a wide range of effectors through the activated subtypes of heterotrimeric () G proteins (Gαs, Gαi, Gαq, G12/13 and

Gβγ). In a migratory cell, the activated GPCRs at the leading edge result in the formation of lamellepodia, primarily through Gmediated PI3K activation. However, it is not clear how the activated of GPCRs in the leading edge induce the retraction of the trailing edge.

While the leading edge activities are attributed to Gβγ, retraction of the TE is assigned to

G12/13 induced activation of Rho guanine-nucleotide-exchange factors (RhoGEF) resulting in the activation of RhoA. Here, we demonstrate a novel mechanism for the trailing edge retraction that is independent of G12/13, which is mediated by Gβγ induced

iii calcium. The data show that free Gβγ extensively results in not only PI3K activation but also increases cytosolic calcium through PLCIP3 pathway. However, the role of free

Gβγ or intracellular calcium in the retraction of trailing edge in migrating cells has not been investigated. The work described in this thesis demonstrate the existence of the pathway;

GβγPLCβ PIP2 IP3 calcium calcium-calmodulin complex myosin light chain kinase myosin phosphorylation actomyosin contraction, as a mandatory requirement for the trailing edge retraction in cell migration.

Acknowledgements

I would like to express my highest gratitude to my advisor and mentor, Dr. Ajith

Karunarathne, for his support, guidance and encouragement during the past two years of research.

I would like to extend my deepest gratefulness to my committee members Dr.

Donald R. Ronning, and Dr. Jon R. Kirchhoff. I would like to thank the Department of

Chemistry and Biochemistry, the University of Toledo for providing me the opportunity to pursue my studies.

Lastly, I would like to thank all my lab members for all their support, advice and words of encouragement, and for all the insightful scientific discussions.

Table of Contents

Abstract……………………………………………………………………………….... iii

Acknowledgements………………………………………………………………….…...v

Table of Contents……………………………………………………………………...... vi

List of Figures……………………………………………………………………..….....ix

List of Tables…………………………………………………………………………….xi

List of Abbreviations………………………………………………………………...…xii

List of Symbols ...... xiv

1 Introduction …………………………………………………………………... 1

1.1 G protein coupled receptors (GPCRs)…………...…………………………....1

1.2 Gα subunits …………...……………………………………………………....2

1.3 Gβγ subunits ………………………………………………………………….3

1.4 G protein signaling ……………………………...……………………………5

1.5 Role of GPCRs in diseases ……...………………………………………..…..6

1.5.1 Cancer metastasis ……………………………………..……..…... ..6

1.5.2 Heart diseases ……………………………………….………...... 7

1.6 Cell migration ………………………………………………….…...... …….8

1.7 GPCR mediated cell migration ……………………………...…………...... 9

1.8 Perturbation of GPCR signaling through photo-sensitive GPCRs.………….12

1.9 Thesis objective… ……………………………...…………..………………..13

2 Does free G control both elongation of the LE and retraction of the TE

during cell migration?...... …….…...……………………...... ….……..15

2.1 Introduction…………………..……………………………………………... 15

2.2 Goal……….. ………………………………………………………...... 16

2.3 Materials and methods…………..………………………….…………...... 17

2.3.1 Cell culture and Transfections ……….……………………………..17

2.3.2 DNA constructs and Reagents …….……….….………..…………..18

2.3.3 Cytosolic calcium measurements..…………………..………………19

2.3.4 Live cell imaging and Data analysis..……………… ……………....19

2.3.5 RT-PCR Assay..……………….………………..………………...... 20

2.4 Results ……………….…………………………..………….…………….…21

2.4.1 OA of blue opsin to control Subcellular G protein activation and

directional cell migration …………………………...... 21

2.4.2 Can Gαs and Gαq coupled GPCR activation result in result in a

similar free G generation response?...... 25

2.4.3 Can Gαs coupled GPCR activation result in cell migration similar to

Gαi?...... 28

2.4.4 Cell migration is governed by Gαi-G and calcium……………….30

2.4.5 Will gallein, the G inhibitor interfere with the heterotrimer

dissociation and activity of G?...... 34

2.4.6 Is G mediated PIP3 alone sufficient to govern cell

migration?...... 36

vii

2.4.7 Does intracellular calcium regulate the G mediated PIP3.……..41

2.4.8 Regulation of Ca-CaM mediated phosphorylation………………..43

2.4.9 G mediated PIP2 hydrolysis……………………………………44

2.4.10 Calcium mobilization in RAW cells is steered by Gi coupled

GPCRs…………………………………………………………...49

2.5 Discussion……………………..………………………...... ……...54

2.6 Conclusion………………………………………………………………...... 58

References……………………………………………………………..………...... 60

A Appendix……………………………………....………………………………..70

Table 1……………………………………....…………………...70

viii

List of Figures

1-1 Schematic of G protein–coupled receptors. ……………………………………....2

1-2 Schematic representation of GPCR activation.……………….……...... 3

1-3 Steps involved in cell migration...... ……………………………………..9

1-4 Conventional GPCR signaling pathway depicting the events at the LE…………10

1-5 Conventional GPCR signaling pathway depicting the events at the TE…………11

1-6 Schematic representation of blue opsin ………………………………………....13

2-1 Schematic representation of GPCR translocation assay…………………………22

2-2 Images showing a HeLa cell expressing blue opsin and mCh 9………………..23

2-3 Representative images of HeLa cell transiently transfected with mCh γ9, before

and after activation of CXCR4 ……………………………………………….…25

2-4 Images showing translocation of GFP in HeLa cells before and after activation

with norepinephrine ……………………………………………………………..26

2-5 Gq coupled GPCR activation induces PIP2 hydrolysis…………………...... 28

2-6 Gαs coupled GPCR activation can induce cell migration………………………..30

2-7 Gαi coupled receptors can induce complete cell migration in RAW cells...... 33

2-8 Gallein does not interfere with heterotrimer dissociation and GGTP

activity……………………………………………………………………….…..35

2-9 Schematic representation of mCh-CRY-iSH2 construct and its activation….…..36

2-10 A RAW cell expressing mCh-CRY-iSH2, CIBN-CaaX and AKT-PH-Venus was

optically activated…………………………………………………………….…37

2-11 The images show RAW cells transfected with blue opsin-mCh and their responses

to localized OA…………………………………………………………………..40

2-12 RNA-seq data showing the relative expression of Geffectors in RAW cells...41

2-13 RAW cells expressing blue opsin and AKT-PH-mCh, untreated and incubated

with cell permeable calcium chelator BAPTA-AM…………………………...... 42

2-14 Calcium mediated actomyosin contractility…………………………………...…43

2-15 The images show RAW cell transfected with blue opsin-mCh incubated with A- 7

hydrochloride…………………………………………………………………….44

2-16 RT-PCR mapping of genes required for Gβγ to induce MLCK activation in RAW

cells………………………………………………………………………………45

2-17 Schematic representation of mCh-PH sensor activity……………………………46

2-18 The images show PIP2 dynamics in HeLa cells expressing mCh-PH……..…….48

2-19 Representative images of RAW cells showing calcium mobilization……………52

2-20 Representative images of RAW cells showing calcium mobilization……………53

2-21 Plot shows the overlap between the absorption spectrum of blue opsin and the

emission spectrum of the EGFP……………………………………………...….53

2-22 Proposed signaling pathway…………………………………………………….58

List of Tables

1-1 Gβγ effectors and their physiological functions ...... 5

1 RAW 264.7 gene specific primers for RT-PCR…………………………………71

List of Abbreviations

ACs …………….Adenylyl cyclase BAPTA………...1, 2-bis (O-aminophenoxy) ethane-N, N, N′, N′-tetra acetic acid BAPTA-AM……1,2-bis(2-aminophenoxy)-ethane-N,N,N'N'-tetra acetic acid tetrakis (acetoxymethyl) ester Ca-CaM………..Calcium-calmodulin complex C5aR …………..Complement component C5 receptor C5a…………….Complement component C5 DAG……………Diacylglycerol EMT……………Epithelial-mesenchymal transition ER……………... Endoplasmic reticulum FA………………Focal adhesions GAP……………GTPase-activating proteins GDP……………Guanosine diphosphate GFP……………Green fluorescent protein GIRK…………. G protein-coupled inwardly rectifying potassium channel GPCRs………...G protein coupled receptors GRPR…………Gastrin releasing peptide receptor GRK………… ...G protein coupled receptor kinase GTP…………... Guanosine triphosphate HBSS…………...Hanks balanced salt solution IM……………. . Internal membranes IP3……………..Inositol 1, 4, 5 trisphosphate IP3R……………Inositol 1, 4, 5 trisphosphate receptor mAbs…………..Monoclonal antibodies mCh…………...mCherry min…………….Minutes MLC…………..Myosin light chain MLCK………...Myosin light chain kinase MLCP…………Myosin light chain phosphatase MYTP1………..Myosin phosphatase target subunit 1 NE……………..Norepinephrine OA……………..Optical activation PIP2…………...Phosphatidylinositol-4, 5-bisphosphate PIP3…………...Phosphatidylinositol-3, 4, 5-bisphosphate PI3K…………..Phosphoinositide 3-kinase PKC…………...Protein kinase C PLC…………...Phospholipase C

xii

PLCβ……………Phospholipase Cβ PM……………....Plasma membrane PTx…………...... Pertussis toxin RhoGEF……...... Rho family guanine nucleotide exchange factor ROCK………….Rho kinase RAW 264.7……..RAW s…………………seconds SERCA…………Sarcoplasmic/ER calcium-ATPase pump TE………………Trailing edge TM……………...Transmembrane Wt………………Wild type YFP…………….Yellow fluorescent protein α2-AR…………..Alpha2-adrenergic receptors 1-AR…………..eta1-adrenergic receptor 2-AR…………..eta2-adrenergic receptor 2APB……………2-Aminoethoxydiphenyl borate

xiii

List of Symbols

% ...... Percentage µ ...... Micro α ...... Alpha β ...... Beta γ ...... Gama  ...... Delta

xiv

Chapter 1

Introduction

1.1 G protein coupled Receptors (GPCRs)

Of the vast families of receptors, by far the largest, most versatile and ubiquitous is the G protein coupled seven-transmembrane receptors, GPCRs. These receptors control most of the physiological processes in cells. GPCRs are classified into five families: rhodopsin, secretion, glutamate, adhesion and taste/frizzled depending upon their sequence homology and functions1-3. Rhodopsin receptors are considered as the family among GPCRs and are the first set of receptors to be studied to determine the structure of GPCRs3, 4. All GPCRs have an extracellular N-terminus, an intracellular C terminus, and seven transmembrane domains (TM 1-7) in the PM5, 6 (Fig. 1-1). All the seven TM domains are linked together by 3 intracellular and 3 extracellular loops5 and have a highly organized structure. The first

X-ray crystal structure studied was that of bovine rhodopsin bound to its inverse agonist

11-cis retinal4. Furthermore, from the structure of rhodopsin in its non-ligand bound form it was revealed that the structure of rhodopsin varied with that of the opsin, where, opsin in its dry form showed structural deviations in the TM helix regions7. Structure of β2-AR bound to its inverse agonist carazolol has also been studied 8. As of today, more than 35 crystal structures of GPCRs have been studied for their structural similarities which

1 unraveled the complexities involved in different types of ligand binding and conformational changes in the TM regions upon ligand binding.

Figure 1-1. Schematic of G protein–coupled receptors. G protein–coupled receptor with its extracellular N-terminus segment and an intracellular C-terminus segment. The seven transmembrane (TM) spanning domains (TM 1–7) are linked by alternating intracellular (i1–i3) and extracellular (e1–e3) loops9.

1.2 Gα subunits

GPCRs are coupled to heterotrimeric G proteins composed of an α subunit bound to GDP and a Gβγ dimer 10. Upon activation of the receptor by the agonist, receptors undergo conformational changes causing GDP to GTP exchange on the Gα subunit which results in the heterotrimer dissociation into G GTPand Gβγ subunits11 (Fig. 1-2). In humans, 16 genes encode for 21 Gα subunits12. These have been classified into four groups based on their sequence similarity and functions13. These include Gαs, which stimulates AC, Gαi which inhibits AC, Gαq which activates PLC and Gα12/13 which activates RhoGEFs13.

The crystal structures of Gα in its GDP bound form and as well, GTP bound forms have been resolved 14.

Figure 1-2. Schematic representation of GPCR/G-protein interactions during signaling. In the inactive state, GPCR couples with heterotrimeric G proteins (Gαβγ). Upon agonist binding, the receptor undergoes a conformational change that catalyzes the exchange of GDP for GTP on the Gα subunit. GTP-bound Gα and the βγ complex dissociate and activate downstream signaling.

1.3 Gβγ subunits

The constitutive dimer of G and G forms the second half of the G protein heterotrimeric complex. In humans 5 Gβ subunits and 12 Gγ subunits have been identified 15. On studying the sequence homology among the Gβ, it was revealed that Gβ1-4 are highly homologous

36 KDa proteins with more than 80% amino acid sequence similarity between each other.

The homology of Gβ5 with other Gβ subunits is <50% 16. In contrast, all Gγ subunits are small (7 KDa ) proteins, which are significantly different from each other 17. With 4 Gβ’s

(excluding β5) and 12 Gγ’s, 48 different combinations of Gβγ dimers are possible17. The

3 structures of Gβγ in complex with Gα subunits as well as individual subunits have been solved18. These crystal structure studies have helped understanding of the heterotrimer dissociation-association process, as well as the identification of effector interaction sites19,

20. However, the functional roles of diverse combinations of Gβγ isoforms has not been established.

During the posttranslational modifications all Gγ subunits are prenylated at the C terminus specifically at the CaaX motif, where‘C’ represents the cysteine and ‘a’ represents the aliphatic amino acids. The isoprenoids are first attached to the cysteine by a thioester bond followed by cleavage of the aaX amino acids and subsequent carboxymethylation of the C terminus. ‘X’ amino acid dictates the nature of the prenylation, with the amino acids Serine,

Methionine, Glutamine and Alanine, leading to attachment of a 15-carbon farnesyl group and leucine leading to attachment of a 20-carbon geranylgeranyl moiety21-23. The N terminus Gβ subunit interacts with the N terminus of the  subunit. Unlike the Gα subunits, no studies have reported about the conformational changes in Gβγ subunit either in free or heterotrimer24. The free Gβγ subunits have been shown to interact with a variety of effector molecules, known to be involved in signaling pathways governing the cell survival (Table

1-1).

Type of G effectors Function Inwardly rectifying K+ channel Channels permeable to potassium ions25 (GIRK1,GIRK2, GIRK4) GPCR kinase 2 and 3 Regulate the activity of GPCRs26 PLC 1, 2, 3 Catalyzes the formation of IP3 and DAG27-29 AC2, AC4, AC7 Activation of adenylyl cyclase and cAMP generation30, 31 AC1, AC3, AC5, AC6 Inhibition of adenylyl cyclase30- 32 N type Ca2+ channels Facilitates calcium movement33 P/Q type Ca2+ channels Facilitates calcium movement34 PI3K Conversion of PIP2 to PIP3, lamellepodia at LE35 SNAP-25 Protein responsible for membrane fusion36 P114RhoGEF, FLJ00018, P-Rex1 GEF activity towards RhoA, Rac1 and Cdc4237-39

Table 1-1. Gβγ effectors and their physiological functions.

1.4 G protein signaling

GPCRs are crucial for maintaining basic biological processes such as tissue homeostasis, development and immunity40. G-proteins interact with effector proteins, resulting in rapid changes in the downstream effector molecules. However, activation of G-proteins have also been associated with several tyrosine kinase receptors, including the receptors for epidermal growth factor (EGF) and insulin40. Primary importance and significance of

GPCRs is more evident in the surplus of drugs that target these receptors. Typically, G protein signaling pathway is composed of GPCRs, G proteins and intracellular effector molecules. The ability of GPCRs to mediate the signaling to intracellular effectors is attributed to response mechanisms evoked by external sensory stimuli 41. Activation of

GPCR signaling proceeds with the activity of G proteins which act as a bridge between the receptors and downstream signaling molecules. The stimuli, which activate the GPCRs,

5 include diverse biomolecules such as neurotransmitters (serotonin for 5HT1B), chemoattractants (cAMP for cAR1), hormones (glucagon and epinephrine), peptides

(SDF1), small molecules (isoproterenol) and light. Upon stimuli/agonist binding, the receptor undergoes a series of conformational changes within the TM region leading to the conversion of GDP to GTP on the  subunit42. This GTP bound form of Gα has a lower affinity for Gβγ, thus leading to heterotrimer dissociation. After the receptor activation, the intrinsic GAP activity which negatively regulate the promotion of GDP to GTP exchange on the Gα subunit results in the heterotrimer reassociation43. Collectively, G protein signaling pathways regulate many physiological and pathological functions ranging from cell migration to apoptosis.

1.5 Role of GPCRs in disease

GPCR activation results in the generation of GGTP and free G subunits. These G subunits interact with many different effector proteins as described earlier. As G signaling is involved in the activation of a large and diverse group of effector molecules, abnormalities in their signaling result in a variety of pathological conditions. For example, it has been proved that G dependent regulation of PI3K plays a central role in leukocyte migration in response to receptor activation which result in cancer cell migration44.

1.5.1 Cancer and Metastasis

The disease condition cancer is defined as the condition involving the abnormal proliferation of cells and leading to tumor development. The transformation of a normal cell into a tumor cell is a multistage process, typically the progression starts from a pre- cancerous stage to generate malignant tumors. These malignant tumors have the ability to

6 invade other tissues as a result of metastasis45. Metastasis constitutes of several sequential steps (i) local tumor cell invasion (ii) penetration of the walls of blood vessels and circulation through the bloodstream (iii) reach other sites in the body and proliferation46.

Recent statistics reveal that cancer metastasis is mostly responsible for cancer related deaths over the primary tumors47. In many cases, it is possible to cure the tumor in its early stage by using chemotherapy, surgery or employing irradiation if it is restricted to the location of origin. However, if the tumor is detected after it has metastasized to other organs, it becomes difficult to treat. Metastasis can show an organ-specific pattern of spread. For example, breast and prostate cancer often metastasize to bone, liver and lungs.

Similarly, if the primary tumor is originated in the lungs, it can metastasize to the other organs including brain, liver and kidneys48. Metastasis is difficult to detect because it might take a long time for the tumor to develop even after the primary tumor site is cured. In this thesis we will focus on understanding the cell migration mediated by calcium signaling, thereby paving the way to identify molecular targets that may be useful in therapy development for metastatic conditions.

1.5.2 Heart diseases

Recent studies indicate that G signaling plays a vital role in heart related disorders49.

Heart muscle contraction is positively controlled by the activation of -AR50. Similarly, muscarinic acetylcholine receptors (M-ChR) modulate the pacemaker activity, atrioventricular conduction, and directly (in atrium) or indirectly (in ventricles) force of contraction51. It has also been reported that regulation of Gs and Gi protein signaling might result in cardiac hypertrophy and heart failure52. During disease progression, higher

7 concentrations of catecholamine can lead to excessive stimulation of -AR resulting in generation of an elevated amount of free G Gis known to interact and control the activity of GRK2, which is responsible for the phosphorylation of GPCRs leading to receptor desensitization54. Furthermore, the Greleased upon activation of Gq coupled

GPCRs has been shown to interact with extracellular signal-regulated kinase (ERK) resulting in the phosphorylation of substrates such as mitogen and stress-activated protein kinase-1 (MSK1) leading to cardiac hypertrophy55.

1.6 Cell Migration

Cell migration involves the translocation of cells from one location to another. Directional cell migration is a highly coordinated multistep process that plays a central role in wide variety of biological processes including embryonic morphogenesis, tissue repair and regeneration and is mainly responsible in the pathogenesis of cancer through metastasis56,

57. In a migrating cell, a complex set of regulatory pathways act simultaneously coordinating the migration process58, 59. When a cell senses extracellular cues (ligands) it starts migrating towards the signal. During this process the cell polarizes forming a distinct

LE and TE. At the LE cell undergo polarization and extend in the direction of external cues forming protrusions called lamellepodia, which is driven by actin-polymerization. These extensions are stabilized by focal adhesions to the extracellular matrix and serve as traction sites for the migrating cell as it moves forward. As the cell moves forward, the focal adhesions at the TE disassemble resulting in TE retraction and complete cell migration (Fig

1-3). At the TE, the actin filament disassembly/destabilization occurs and the old adhesions

8 will be detached60. Failure to retract the TE results in mere elongation of the cell but not complete migration.

Figure 1-3 Steps involved in cell migration. 1. Elongation 2. Extension of a lamellepodia 3.Formation of a new focal adhesion. 4. Translocation of the cell 5. Retraction of the TE60.

1.7 GPCR mediated cell migration

GPCRs are cell surface receptors which play an important role in cell migration. Gαi coupled GPCRs induce chemotaxis through the regulation signaling molecules including

Rho family of small GTPases such as RhoA, Cdc42, and Rac61-64. Additionally, free Gβγ subunits activate PI3K producing PIP3 at the LE65.

Figure 1-4 Conventional GPCR signaling pathway depicting the events at the LE. GPCR activation leads to the generation of free Gβγ. Gβγ activates PI3K resulting in the lamellepodia formation at the LE via downstream signaling proteins.

Gβγ maintains the cell polarity and lamellepodia formation by regulating the actin cytoskeleton remodeling in mammalian cells through the control of downstream signaling molecules including Rac and Cdc42 at the LE66-68. Rac and Cdc42 control the lamellepodia formation by activating the suppressor of cAMP receptor (SCAR), WASP-family verprolin-homologous protein (WAVE) and Wiskott–Aldrich syndrome protein (WASP) proteins. These scaffold proteins activate the actin regulatory protein (Arp2/3) complex, which nucleates branching of actin filaments by binding to the barbed ends of the filamentous actin and remodel of the actin cytoskeleton. (Fig 1-4). The lamellepodia is a

10 result of this actin nucleation at the LE of the cell. While lamellepodia formation is seen at the LE, the retraction of the TE has been assigned to the activity of G12/13 subunits69

(Fig 1-5).

Figure 1-5 Conventional GPCR signaling pathway depicting the events for TE retraction. G12/13 mediated activation of RhoGEF followed by RhoA activation resulting in RhoA/ROCK mediated contraction at the TE.

The G12/13 subunits have been involved in Rho activation and signaling via the activation of a RhoGTPase through RhoGEFs (p114-RhoGEF and p115-RhoGEF)69, 70.

Further, RhoA/ROCK mediated pathway is linked to invasion and metastasis of cancer cells71. The GTPase, RhoA, is initially present in the inactive GDP form, which is

11 converted to the active GTP-bound form. Upon this conversion, RhoA induces the activity of a ROCK, which plays a critical role in cell adhesion and contraction72. ROCK phosphorylates MLCP thereby inhibiting its activity (Fig 1-5)73, 74. This results in an increase in MLC phosphorylation, consequently increasing actomyosin-based contractility75.

1.8 Perturbation of GPCR signaling through photo-sensitive GPCRs

The activity of GPCRs can be studied by monitoring their downstream signaling activity.

Conventionally, genetic, chemical or pharmacological methods have been used to study cellular activities. The main drawback of these methods is that they provide no spatial and temporal control. Recent studies and developments in the field of optogenetics have presented researchers with tools to investigate and study the signaling and cellular activities with precise spatial and temporal control76, 77. In GPCR optogenetics, opsins are genetically engineered to be used as light sensitive GPCRs. Opsins are photoreceptors and they undergo conformational change from inactive state to signal activating state upon absorption of light. In opsins, the GPCR ligand is a form of retinal bound to the opsin via a Schiff base linkage. For our studies we employed a recently reported light sensitive

GPCR, human cone photoreceptor blue opsin78 (Fig 1-6 A). Blue opsin (absorption max.

414 nm) can activate Gi pathway upon exposure to blue light (445 nm) once it is supplemented with 11-cis retinal as the chromophore. Retinal chromophore bound blue opsin absorbs blue light causing a conformational change with the conversion of 11-cis retinal to all-trans retinal. This induces the subsequent dissociation of the heterotrimer.

Here, we show that genetically encoded human cone blue opsin can activate endogenous

Gi/o G proteins in subcellular regions of a cell (Fig 1-6B). By employing blue opsin, spatial and temporal control of G protein can be achieved to direct cell migration. Further, to monitor the intracellular processes like the G protein activation and PIP3 accumulation we have used different fluorescently tagged sensors. We have used fluorescent proteins including GFP (488 nm), YFP (515 nm) and mCh (594 nm) for this purpose.

Figure 1-6 (A) Schematic representation of blue opsin used, with its λ-max and the type of G protein it is coupled with. (B) Illustration of blue opsin activation upon retinal chromophore binding using 445 nm blue light. Upon OA blue opsin undergoes conformational changes and result in the heterotrimer dissociation.

1.9 Thesis objectives

The main objective is to identify how G protein Gαi and Gβγ subunits mediate the activation of effector molecules at the LE to govern the retraction of the TE. To achieve

13 required confined G protein activation at the LE, we have used a human blue opsin (Gαi coupled) which was activated with 445 nm blue light while, longer wavelengths including

515 nm and 594 nm were used to image molecular and cellular responses. In our preliminary studies, we observed that when opsins are subjected to OA at spatially confined regions of a migratory cell, the cell migrates towards the optical stimulus. This confined activation is confirmed by monitoring free Gβγ subunits translocation from the activated area to IMs of the cell. Here, we focused on understanding how cells achieve complete migration in response to a localized activation of GPCRs, without activating the Gα12/13 subunits as well. We also hypothesized that the free Gβγ subunits are responsible for the

TE retraction and play a broader role in controlling of the entire cell migration process.

Experiments were carried out using subcellular optogenetics together with pharmacological perturbations to study and understand the mechanisms and factors that govern cell migration.

Chapter 2

Does free Gcontrol both elongation of the LE and retraction of the TE during cell migration?

2.1. Introduction

Cell migration is associated with activity of receptors on the cell surface. These cell surface receptors are stimulated by chemoattractants and initiate the intracellular signaling. GPCRs make up for the majority of the cell surface receptors. GPCRs upon activation transmit the signal through heterotrimeric G proteins Gα and Gβγ. Upon activation by the ligand binding, conformational changes are induced in the GPCR that causes G protein activation5. The Gα subunit of the G protein releases GDP in exchange for GTP and once activated the Gα subunit dissociates from the Gβγ subunits, Gα and Gβγ subunits relay signals to a wide range of downstream effectors, including AC isoforms79, PLC80, ion channels81, protein tyrosine kinases, and MAP kinases82, among other to induce biochemical modifications that play a vital role in various cellular processes22.

In metastasis, cells migrate through dense extra cellular matrix (ECM) by developing protrusions. These protrusions have the ability to alter and degrade the ECM83. This type of protrusion are particularly seen in the cells near blood vessels, which helps the cancer cells to invade through the basement membrane. Invasive cancer cells form actin-rich

15 membrane protrusions extended vertically from the ventral cell membrane. These structures have been termed invadopodia84. Invadopodia possesses enhanced levels of actin regulatory proteins, adhesion molecules, signaling/adaptor proteins, membrane remodeling proteins and matrix degrading proteases, that are required for matrix degradation activities85.

One such family of GPCRs known to regulate cell migration are chemokine receptors including C-X-C chemokine receptor type 4 (CXCR4)86. Chemokines are small proteins that bind to their cognate receptors to stimulate directional migration also called as chemotaxis. Dysregulations in chemokine pathways have been associated with numerous pathological conditions such as metastatic advancement during tumor progression. For example, the enhanced CXCR4 expression increases the migratory capacity of non-small cell lung cancer (NSCLC) cells and melanoma cells87, 88. This suggests that they are involved in metastatic dissemination. Further studies to establish the role of CXCR4 in metastasis conditions revealed that, during cancer treatment, in patients treated with anti-

CXCR4 mAbs showed inhibition of the metastatic spread to target organs in vivo89.

2.2. Goal

The current study focusses on understanding how free Gβγ induces complete cell migration in response to receptor activation at the LE, without activating the Gα12/13 subunits, which is known to induce retraction at the TE. We assumed that the free Gβγ subunits are responsible for the TE retraction and play a broader role in controlling the formation of

LE-TE polarity in migrating cells. Using mouse macrophage cells, RAW, we investigated potential pathway through which Gβγ activates PLC and induce the release of cytosolic

16 calcium from the ER. Cells contain a various intracellular calcium-binding proteins and one of the major calcium-binding protein is calmodulin (CAM)90. Ca-CaM complex activates downstream targets including the MLCK and induce the phosphorylation of MLC resulting in actomyosin contraction. Here, we present evidence that Gβγ interaction with

PLC, is sufficient for the TE retraction and does not require the activation of Gα12/13 subunits. The data presented here can provide an insight into the involvement of calcium in cell migration and can be treated as an important therapeutic target for anti-metastatic drug development.

2.3. Materials and methods

2.3.1. Cell culture and Transfections

HeLa cells (ATCC, Manassas, VA) were cultured in minimum essential medium (CellGro) supplemented with 10% dialyzed fetal bovine serum (Atlanta Biologicals), in the presence of 1% penicillin−streptomycin in 60 mm tissue culture dishes at 37 °C in a 5% CO2 humidified incubator and sub cultured every 2 or 3 days. At 75% confluency, adherent cells were detached after incubating with versene-EDTA (CellGro), centrifuged at 1000×g for

3 min, and versene-EDTA was aspirated before resuspending the cell pellet in the regular culture media at a cell density of 1 × 106 cells/mL. RAW 264.7 mouse macrophage cell line (ATCC, Manassas, VA) were cultured in RPMI 1640 (10-041-CV; CORNING,

Manassas,VA) with 10% dialysed fetal bovine serum (Atlanta Biologicals, Flowery

Branch, GA), L-glutamine and 1% penicillin–streptomycin in a humidified incubator at 37

°C and 5% CO2. Cells were sub cultured every 2–3 days when they achieved 70–80% confluency and for experiments the cells were transferred to 35 mm glass bottomed dishes

(8×104 ml/dish) (In Vitro Scientific) RPMI 1640 with10% dialyzed fetal bovine serum is used on the glass bottom dishes. RAW cells ranging from passage 3 to passage 25 were used for experiments. RAW cells were transfected by a biodegradable polymer based DNA transfection reagent (PolyJet™) according to manufactures protocol. Transfection was performed on 8×104 million cells plated on a glass bottom dish. After 5 h of transfection, the transfection media is aspirated and 1 mL of fresh warm culture medium was added to the dishes, placed in an incubator at 37 °C and 5% CO2, and imaged 12-14 h after transfection. RAW cells were transfected by electroporation using the D-032 on an Amaxa

Nucleofector device (Lonza, Basel, Switzerland). Each electroporation was performed on

1.25 million cells in 200 μL of Nucleofector solution, immediately followed by addition of

800 μL of warm culture medium. The cells were then plated on glass bottom dishes, placed in the incubator at 37 °C and 5% CO2, and imaged 12-14 h after electroporation.

2.3.2. DNA constructs and reagents

Engineering of blue opsin-mCh, blue opsin-mTurquoise, AKT-PH-mCh constructs have been described previously91,104. pCAGGS Raichu-RhoA-CR (plasmid 40258) was obtained from addgene. HBSS (10x) (Gibco laboratories), pertussis toxin (PTx),

SDF1isoproterenol hydrochloride, norepinephrine (SigmAldrich), gallein (TCI

AMERICA), fluo-4 (Molecular probes, Eugene, Oregon), BAPTA, BAPTA-AM, thapsigargin, wortmannin, A-7 Hydrochloride, 2APB (Cayman Chemical, Ann Arbor, MI), c5a (Eurogentec, Belgium), GSK 269962 (AdooQ Bioscience, Irvine, CA), ML-7 hydrochloride (AdooQ Bioscience, Irvine, CA), bombesin (Tocris), 11-cis retinal (National eye institute). 11-cis retinal was reconstituted in 200 proof ethanol and added to the cells.

All the reagents except PTx were dissolved in DMSO. HBSS was used to dilute these reagents to the required concentrations and then added to the cells.

2.3.3. Cytosolic calcium measurements

For intracellular calcium measurements, cells were cultured at on cell culture dishes in a

37 °C incubator with 5% CO2. Experiments were performed 12 to 24 h after seeding. Cells were then first washed twice with an external solution (1x HBSS with Ca2+, (pH 7.2) before calcium measurements. Then, cells, and were incubated for 25 min with the fluorescent calcium indicator, fluo-4. Fluo-4 fluorescence intensity (488 nm) was continuously imaged at 1 s intervals, using confocal microscopy. Fluo-4 fluorescence intensity obtained from regions of interest was normalized to the baseline intensity.

2.3.4. Live cell imaging and Data analysis

These experiments were performed with a 60x, 1.4 NA oil objective or a 10x, 0.3 NA objective in a spinning-disk XD confocal TIRF imaging system that is composed of a

Nikon Ti-R/B inverted microscope, a Yokogawa CSU-X1 spinning disk unit (5000 rpm), an Andor FRAPPA unit for photoactivation of manually selected regions of the opsins in real time, all controlled using Andor iQ 3.1 software (Andor Technologies, Belfast, United

Kingdom). 445, 488,515, and 594 nm solid-state lasers and iXon ULTRA 897BVback- illuminated deep-cooled EMCCD camera were used. Blue opsin-mCh, AKT-PH-mCh were imaged using 594 nm excitation−630 nm emission settings, blue opsin-mTurquoise was imaged using 445 excitation and 478 emission and fluo-4 was imaged using 488 excitation and 515 emission respectively. For OA (white box) of opsins, the 445-nm laser

19 was used at 5 μW across the selected region. This was performed in 3 s interval. Mean pixel fluorescence intensity changes in the entire cell or in selected areas of the cell were determined using Andor iQ 3.1 software time-lapse images were analyzed using the analytical tools accompanied by Andor iQ 3.1 software, smoothed graphs and histograms were generated using OrignPro computer program for interactive scientific graphing and data analysis (OriginLab Corporation).

2.3.5. RT-PCR Assay

RAW cells were homogenized and mRNA was isolated with GeneJET RNA purification kit according to the manufacturer protocol. Gene expression was quantified by real-time quantitative PCR (RT-PCR) using the RADIANT Green LO-ROX qPCR Kit (Alkali scientific, Pompano Beach, FL). DNA amplification was carried out using Icycler (BIO-

RAD, Hercules, CA), and measured the binding of the fluorescence dye SYBR Green to double-stranded DNA. All the primer combinations were provided by IDT (see Appendix).

The relative quantities of target gene mRNA against an internal control, beta-actin, was possible by following a ΔCT method. The difference (ΔCT) between the mean values in the replicated samples of target gene and those of beta-actin were calculated by Microsoft

Excel, OriginPro computer program. The relative quantified value (RQV) was expressed as 2-ΔΔCT.

2.4. Results

2.4.1. OA of blue opsin to control Subcellular G protein activation and directional cell migration.

It has been previously reported that human cone photoreceptor, blue opsin can be activated asymmetrically to induce directional migration92. Using this approach, we examined if the

OA can be confined exclusively to the LE. For this, we used a recently developed Gγ9 assay that employs GPCR induced translocation of mCh γ9 from the PM to IMs91 (Fig 2-

1). Here, the mCh γ9 (mCh fluorescent tag attached to the N-terminus of Gγ9) acts as a sensor for detecting the spatial and temporal activation of receptor in single cells91. The expressed mCh γ9 forms a dimer with the endogenous Gβ to form the heterotrimer

(GαGDPβγ).

Figure 2-1 Schematic representation of GPCR translocation assay. (1) GPCR bound to the heterotrimer in the inactive state. (2) GPCR activation upon ligand addition triggers the G protein activation followed by heterotrimer dissociation and translocation of Gβγ to IM’s. (3) Shuttling of the βγ subunits between the PM and IM’s. An equilibrium exists between the βγ subunits on the PM and IM’s. (4) With the removal of ligand, the concentration of αGDP is increased and the βγ subunits relocate to the PM. (5) Complete reversal of the translocation and the heterotrimer is associated back on the PM.

Using ethanol-reconstituted 11-cis retinal (50 µM) and single pulse of 445 nm blue light

(White box), blue opsin is optically activated globally to induce the receptor activation.

Due to the conformational changes upon receptor activation, the heterotrimer dissociates into GαGTP and Gβγ. The resultant free Gβγ shuttles between the PM and IMs (Fig 2-2A).

Plot shows the mCh γ9 translocation from the PM (black curve) and IMs (red curve) with

22 a translocation half time (t1/2) of ~7.5 s (Fig 2.2B). With the termination of OA (Blue light), complete reversal of the translocation process was observed.

Figure 2-2. (A) Images showing a HeLa cell expressing blue opsin and mCh 9. White box indicates the region of the OA, using 445 nm laser light pulsed at 0.5 Hz. Addition of 50 μM 11-cis retinal and upon global OA (445 nm, 1 W/m2) white box region), mCh 9 on the PM translocate to the IMs resulting in a transient mCh 9 loss on the PM (yellow arrow) and a gain in the IMs (white arrow) indicating efficient G protein activation. Upon termination of OA, mCh 9 returns to the PM. (B) Plot shows the normalized mCh γ9 fluorescence intensity changes on the PM and IMs. (C) Local OA resulted in translocation of mCh 9 on the PM to the IMs resulting in a transient mCh 9 loss on the PM (yellow arrow) and a gain in the IMs (blue arrow) indicating efficient G protein activation. (D) Plot shows the normalized mCh γ9 fluorescence intensity changes on the front and back PM and IMs. Scale bar, 10 μM.

Next, we examined if the localized subcellular activation of blue opsin can be achieved in a single cell if we confine the light exposure to a region of interest (ROI) on the cell. Using computer steered galvanometer device, a selected 2 × 10 µm2 region of interest (ROI-white

23 box) was exposed to a 445 nm blue light (Fig 2-2C). This resulted in the disappearance of mCh 9 on the PM only in the ROI. The plot shows the reduction of fluorescence intensity on the activated front-PM (solid black) while an increase in adjacent-IMs (solid red) (Fig

2-2D). To check if localized OA resulted in confined receptor activation, we examined the

PM and IM area on the opposite side from the optically activated region and found that neither IM nor the PM regions (dashed lines) showed any fluorescence intensity change.

It has been previously reported that in breast cancer cells, signaling through CXCR4 or

CCR7 mediates actin polymerization, subsequently inducing chemotactic and invasive responses. As both CXCR4 and blue opsin are Gi coupled GPCRs, we comparatively tested if they possess similar G proteins activation abilities. Both blue opsin supplemented with 11-cis retinal and exposed to blue light (Fig 2-2A) and endogenous CXCR4 activated with SDF1α (Fig 2-3 A, B) in HeLa cells co-expressing mCh γ9 resulted in robust G protein activation. Comparative analysis shows that the magnitude of responses induced by both the receptors are approximately equal, indicating that blue opsin is an optically controllable substitute for CXCR4 and therefore can be used as an alternative strategy to control cell migration.

Figure 2-3. (A) Representative images of HeLa cell transiently transfected with mCh γ9, before and after activation of CXCR4 (Gαi) receptor with 100 ng/ml SDF-1. mCh 9 on the PM translocate to IMs resulting in a transient mCh 9 loss on the PM (yellow arrow) and a gain in IMs (white arrow) indicating efficient G protein activation. (B) Plot shows the normalized mCh γ9 fluorescence intensity changes on the front and back PM and IMs. Scale bar, 10 μM.

2.4.2. Can Gαs and Gαq coupled GPCR activation result in result in a similar free

G generation response?

G protein subunits, G are associated with the PM and act as switches that transmit information from cell surface receptors to intracellular effectors93. It was also shown that upon receptor activation the  subunit types  translocate to Golgi complex and

ER94. Here, we examined if Gαs and Gαq coupled GPCRs show differential translocation rates of G subunits when compared to Gαi coupled GPCRs. We imaged the free G generation ability of1-AR (Gs) and GRPR (Gq) with that of the α2-AR (Gi). Cells expressing GFP γ9 were observed for translocation after activation of endogenous α2-AR with 10 µM NE. Similarly, cells expressing 1-AR and GRPR were treated with 10 µM

25 isoproterenol and 1 µM bombesin respectively (Fig 2-4 A, C, E). Plots show the reduction of fluorescence intensity on the PM (black) and increase in IM’s (red) after the agonist addition. (Fig 2-4 B, D, F).

Figure 2-4. (A) Images showing translocation of GFP in HeLa cells before and after activation of Gαi coupled endogenous α2-AR with NE (10 μM) (experiment was performed by Kanishka Senarath). (C) Images showing translocation of GFP in HeLa cells also expressing Gαs coupled 1-AR before and after activation with (10 μM) isoproterenol. (E) HeLa cell expressing Gαq coupled gastrin releasing peptide receptor (GRPR) and GFP show no translocation after GRPR activation with 1 μM bombesin (experiment was performed by Kasun Ratnayake). (B), (D), (F) Plot shows the normalized GFP γ9 fluorescence intensity changes on the PM and in IMs. Scale bar, 10 μM.

Comparative G data show that Gαs coupled GPCRs exhibit similar and robust Gγ9 translocation compared to that of Gαi coupled GPCRs (Fig 2-4). However, Gαq pathway 26 activation only resulted in no translocation response. In order to confirm if GRPR expression is adequate and sufficient activation is induced during this experiment, we examined bombesin induced PIP2 hydrolysis in cells additionally expressing PH-mCh

(PIP2 sensor). A significant expression of GRPR was observed and resulted in a strong

PIP2 hydrolysis upon bombesin addition (Fig 2-5A). Activation of GRPR also resulted in a cell shrinkage (Fig 2-5B), a typical Gq pathway response, suggesting an adequate

GRPR activation. Nevertheless, it failed to generate a detectable amount of free G



Figure 2-5 (A) Gq coupled GPCR activation induces PIP2 hydrolysis. Image sequence shows HeLa cells transfected with Gq coupled GRPR-GFP, overlay of GRPR-GFP – PH- mCh and the change in PH-mCh before and after addition of the bombesin. Cells shows significant PIP2 hydrolysis upon activation with 1 µM bombesin. Yellow arrow shows the loss of PH-mCh on the PM and white arrow shows the accumulation of PH-mCh in the IMs. Plot shows the increase in the fluorescence intensity of PH-mCh in the IM’s upon addition of the ligand. (B) Gq coupled GPCR activation induces cell shape change. HeLa cells transfected with GFP γ9 and Gq coupled GRPR-mCh shows a significant cell shape change upon activation with 1 µM bombesin perturbing the detection of γ9 translocation. The red outline on the image shows the membrane periphery before receptor activation illustrating the extent of cell shape change. (Experiment performed by Kasun Ratnayake). Scale bar, 10 μM.

2.4.3. Can Gαs coupled GPCR activation result in cell migration similar to Gαi?

When compared with Gαi coupled GPCRs, Gαs coupled GPCRs showed similar translocation of Gγ9 subunits upon receptor activation. Evidence shows that Gαi-GPCRs

28 are known to induce cell migration95. So, can we assume that Gαs-GPCRs can also induce cell migration as they show similar G protein activation and translocation to Gαi-GPCRs?

To test this hypothesis we employed an optogenetic chimeric opsin, CrBlue, a Gαs coupled

GPCR. CrBlue was engineered by replacing cytosolic loops of blue opsin with that of the

Gαs coupled box jellyfish (Carybdea rastoni-Cr)78. With retinal, during the initial OA period, RAW cells showed minor migration towards the optical input. The kymograph of small cross section of the PM shows the movement of the LE towards the optical input while at the TE disappears from the fixed virtual plane (orange arrows) (Fig 2-6). However, continuous OA of CrBlue did not result in a continuous directional movement along the axis of GPCR activation (Fig 2-6). This suggests that Gαs-GPCR induced free  generation is insufficient of inducing cell migration (Fig 2-4C, D). Evidence supports this theory as Gαs induced cAMP induces cytoskeleton contractility and inhibits cell migration96.

Figure 2-6. Gαs coupled GPCR activation can induce cell migration. Images on the top shows RAW cell transiently expressing CrBlue mCh. White box indicates the region of the OA, using 445 nm laser light. Blue and green lines indicate the LE and TE of the cell. To the right are the orthogonal slice view of a cross section of these regions. The arrows (orange) shows intensity changes of this cross section over time. Below is the plot showing the change in the mean fluorescence at the front and back of the cell upon OA. Scale bar, 10 μM. (Experiment performed by Kasun Ratnayake)

2.4.4. Cell migration is governed by Gαi-G and calcium.

As localized OA at any position on the surface of a cell can induce G protein activation, we examined whether the directional migration can be controlled entirely by the localized

OA of blue opsin. To examine this, Local OA was applied to a RAW cell expressing blue opsin-mCh. Confined OA of blue opsin in RAW cell resulted in a complete cell migration with lamellepodia formation at the LE and retraction at the TE (Fig 2-7A). To further investigate the Gαi coupled GPCR mediated cell migration we used the inhibitor of Gi/o proteins, PTx. RAW cells were treated overnight with PTx (0.05 μg/mL). PTx-catalyzes the attachment of ADP-ribose to a cysteine residue at the 347 position of the C-terminus and prevents the Gαi heterotrimer dissociation97, 98. A complete inhibition of blue opsin induced cell migration was observed; suggesting that Gαi coupled heterotrimer activation

30 mediates this process (Fig 2-7B). Gis known to interact with the downstream effectors including PI3K to induce lamellepodia formation at the LE93. Therefore, to examine the contribution of Gas an important entity that controls cell migration RAW cells were treated with a small molecule Ginhibitor, gallein (10 μM). Gallein interacts with a 13 amino acid region SIGKAFKILGYPD, which has been identified as a preferred effector interaction surface on Gβ in its propeller region99. In the presence of gallein, cells completely failed to migrate (Fig 2-7C). Histogram shows the comparison of distance migrated by the LE and TE in control cells and cells treated with PTx and gallein (Fig 2-

7G, H). Furthermore, the migration velocities also differed when compared to control cells

(Fig 2-7I).

Evidence shows that activation of Gαi coupled endogenous α2-AR with NE induces an increase in cytosolic calcium100. We hypothesized that, being an important secondary messenger in cells, calcium is also involved in regulating the cell migration, especially in the retraction of TE. To determine if changes calcium regulate cell migration, we examined extracellular and intracellular calcium dynamics during RAW cell migration induced by

OA of blue opsin. First, the cells were incubated in calcium free medium and experiments were performed under the same conditions. When RAW cells were incubated in calcium free cell culture medium for 30 minss by reconstituting the cell culture medium (RPMI- dialyzed fetal bovine serum) with 5µM BAPTA, a chelator of extracellular calcium, cells showed lamellepodia formation but, no TE retraction (Fig 2-7D).

Next, cytosolic calcium chelator, BAPTA-AM was used to examine if it perturbs the directional migration by inhibiting the formation of Ca-CaM complex, which mediates the

TE retraction. RAW cells were incubated with a cell permeable analogue of BAPTA,

BAPTA-AM for 30 minss before OA of blue opsin to induce cell migration. RAW cells treated with BAPTA-AM showed lamellepodia formation at the LE but, little to no retraction at the TE, there by inhibiting the cell migration (Fig 2-7E). To confirm that if the retardation of cell migration is due to chelation of cytosolic calcium by BAPTA-AM, similar experiment was performed using thapsigargin (a SERCA pump inhibitor101) in place of BAPTA-AM. Cells were incubated with 0.5 µM thapsigargin, for 30 mins to substantially reduce sarcoplasmic stored calcium. Thapsigargin treated cells showed no migration while LE activities, including lamellepodia formation, remained unperturbed.

(Fig 2-7F). Histogram shows that both reduction of cytosolic and sarcoplasmic calcium has a greater impact on the TE retraction while LE lamellepodia formation and the overall cell migration velocities are reduced when compared to the control cells (Fig 2-7G, H, I).

Figure 2-7. Gαi coupled receptors can induce complete cell migration in RAW cells. RAW cells transiently expressing Gαi coupled blue opsin-mCh was optically stimulated through localized photo activation to induce asymmetrical directional migration. (A) The image sequence shows subcellular OA of blue opsin-mCh induced directional cell migration in Wt RAW cell. (B) The image sequence shows subcellular OA of blue opsin-mCh does not induce directional cell migration in RAW cell treated with PTx (0.05 μg/mL). (C) The image sequence shows subcellular OA of blue opsin-mCh does not induce directional cell migration in RAW cell treated with gallein (10 μM), a G inhibitor. (D) RAW cell incubated with calcium free HBSS for 1 hour at 37 °C prior to the experiment showed no TE retraction while the LE activities remained unchanged. Similarly, (E) Chelation of cytosolic calcium with 5 μM BAPTA-AM and, (F) Removal sarcoplasmic stored calcium using the sarco/endoplasmic reticulum calcium-ATPase blocker 500 nM thapsigargin (30 mins incubation at 37 °C) resulted in cells responding only with LE lamellepodia formation, however with no effective cell migration. Specially, the cells showed a complete lack of TE retractions. (G), (H), (I) Histogram shows the extent of migration at the LE, retraction at the TE (in µm) and velocity in control type cells, cells treated with PTx and gallein. Cells treated with PTx, gallein, calcium free buffer, BAPTA AM and thapsigargin showed substantial difference in the LE formation and little or no retraction at the TE, endorsing the point that asymmetric directional cell migration is induced by Gαi proteins. The white box marks the region photo activated with 445-nm light. **, P <0.0001 was considered statistically significant value. Scale bar, 10 μm. Time in minutes (0 to 20min).

2.4.5. Will gallein, the G inhibitor interfere with the heterotrimer dissociation and activity of G?

To demonstrate that Gallein binds to the G hotspot and only alters the G-effector interaction, however, does not interfere with heterotrimer dissociation, re-formation as well as the activities of GGTP we performed the following experiments. The translocation of

GFP γ9 upon 1-AR activation was examined in cells pre-treated with 10 M gallein (Fig

2-8A). Next, the Gs induced adenylyl cyclase activation was tested upon 1-AR activation in the presence and absence of 10 M gallein. Here, PKA-RII:PKA-C-YFP, a translocation based probe was used to detect cAMP generation when Gs–GPCRs are activated. Both, cells treated with and without gallein, showed a similar and robust cAMP response, indicating that gallein does not interfere with G signaling (Fig 2-8B, C). Data provide us with the evidence that the Gα signaling is not inhibited upon treatment with

34 gallein and therefore, it can be inferred that free Gβγ is a major player in the chemokine mediated cell migration.

Figure 2-8. Gallein does not interfere with heterotrimer dissociation and GGTP activity. (A) Image sequence shows HeLa cells transfected withAR CFP and GFP 9, before and after addition of the 10 µM isoproterenol. Cells were pre-incubated with gallein (10 µM) for 30 mins at 37 °C. Cells showed significant 9 translocation upon addition of 10 µM isoproterenol. Yellow arrow shows the loss of 9 on the PM and white arrow shows the accumulation of 9 in IM’s. Plot shows the mean fluorescence intensity of GFP 9 in the IM’s and loss on the PM upon addition of the ligand. (B) G activity is independent of gallein: HeLa cells transfected with AR CFP and fluorescent translocation probe of cAMP (PKA-RII:PKA-C-YFP). Image sequence shows the translocation of PKA Cα- YFP after addition of 10 µM isoproterenol to activate 1-AR. Cells were pre-incubated with gallein (10 µM) for 30 mins at 37 °C. Yellow arrow shows the loss of PKA Cα-YFP on the PM and white arrow shows the accumulation of PKA Cα-YFP in the cytosol. Plot showing the mean fluorescence intensity of PKA Cα-YFP in the cytosol and loss on the PM upon addition of isoproterenol. Scale bar, 10 μM.

2.4.6. Is G mediated PIP3 alone sufficient for governing cell migration?

The family of PI3Ks phosphorylate phospholipids bearing the inositol structure at the 3’OH position of the inositol ring. The most common substrate is PIP2, which upon phosphorylation produces PIP3102. PIP3 is an important secondary messenger and play crucial role in lamellepodia formation at the LE103. It was previously shown that, upon switching the optical stimuli, the formation of PIP3 at the new LE and its active reduction at the former LE is synchronized, indicating an active communication and signaling gradient establishment between the LE and TE104. Here, we examined if PIP3 generation at the LE alone can govern the migration as follows. We employed a genetically coded light activatable PI3K, iSH2 fused to fused to the CRY domain with a fluorescent tag, mCh.

A working principle of approach system is shown in Fig 2-9.

Figure 2-9. Schematic representation of mCh-CRY-iSH2 construct and its activation. OA of a spatially confined region on the PM triggers the recruitment of mCh-CRY-iSH2 from the cytosol to the PM. iSH2 is a part of catalytic domain of PI3K and catalyzes the conversion of PIP2 to PIP3 at the LE.

Here, the CRY domain help target the iSH2, inter-Src Homology 2 domain of a p85b regulatory subunit of PI3K (iSH2) which carries endogenous p110 catalytic subunit of

PI3K, to the PM by optical activation. Targeting of PI3K subunits to the PM resulted in a robust PIP3 production at the region of OA but, only a minor cell migration was observed when compared to blue opsin activation induced cell migration. This suggested that PI3K activation induced PIP3 production is insufficient to elicit complete cell migration. PI3K localization on the PM can be observed by the accumulation of mCh-CRY-iSH2 at the region of OA (Fig 2-10A) and a robust PIP3 generation was indicated by the accumulation of AKT-PH-Venus at the region of OA (Fig 2-11B). The plot shows an increase in PIP3 at the LE (Fig 2-10C).

Figure 2-10. (A) A RAW cell expressing mCh-CRY-iSH2, CIBN-CaaX and AKT-PH- Venus was optically activated to recruit mCh-CRY-iSH2 to a confined region. (B)The cell shows a localized PIP3 production, similar to that of blue opsin activation, however, only resulted in a minor migration response. (C) Plot shows the PIP3 production at the LE upon OA. Scale bar, 10 μM (Experiment was performed by Dinesh Kankanmge).

Next, to examine if cell migration, especially the TE retraction is governed by Gβγ induced

PI3K activation, we performed a series of experiments where blue opsin was activated to induce cell migration in RAW cells while inhibiting several signaling proteins involved in the migration pathway as follows. First, Ginteraction with the downstream effectors was inhibited by treating the cells with Gβγ inhibitor, gallein for 30 mins at 37 °C. RAW cells

37 expressing blue opsin-mTurquoise and AKT-PH-mCh were optically activated.

Pretreatment of cells with Gallein made them completely non-responsive to optical activation and failed to migrate (Fig 2-11-I). Second, to inhibit PI3K prior to the migration initiation, cells were incubated with 50 nM wortmannin, a PI3K inhibitor for 30 minss at

37 °C. Interestingly, upon OA of cells treated with wortmannin, cells completely abolished the lamellepodia formation at the LE but managed to show the cytoskeleton retraction globally, reducing the cell size (yellow arrows) (Fig 2-11-II). Furthermore, cytoskeleton shrinkage was not observed in cells treated with gallein, providing us with the evidence for existence of other Gβγ mediated and PI3K independent pathways to control the TE retraction.

Conventional TE retraction pathway includes ROCK mediated actin cytoskeletal reorganization. We checked if inhibition of ROCK activity would still facilitate the G mediated cell migration. RAW cells expressing blue opsin-mTurquoise and AKT-PH-mCh were incubated with 12.5 M GSK 269962 (ROCK inhibitor) for 30 mins at 37 °C. Upon

OA, cells showed lamellepodia formation at the LE but retraction of the TE was abolished

(Fig 2-11-III). Next, RAW cells were treated with both wortmannin and GSK 269962 to examine the effect on G mediated cell migration. Inhibition of both PI3K and ROCK completely abolished the cell migration where no LE or TE activities (Fig 2-11-IV).

Histogram shows the % cell size change which shows the contractility (positive) in wortmannin treated cells and extension (negative) in ROCK inhibited cells (Fig 2-11B).

ROCK inhibition resulted in the inhibition of contraction in cells while opsin activation induced lamellepodia formation resulted in the expansion of cell surface area. While

ROCK is essential for contractility-force generations, it further confirms that the PI3K

38 activation induced PIP3 production at the LE is insufficient to induce complete cell migration. Based on the observation we investigated if G directly activates Rho A and subsequently activating ROCK to facilitate the TE retraction. Data from RNA-seq analysis of RAW cells showed the types of Rho-GEFs that can be potentially involved in activation of RhoA and subsequently ROCK (Fig 2-12). Among them, Rho114-GEF has been identified as a Gβγ activatable RhoGEF and therefore it is likely that in RAW cells, Gβγ controls RhoA pathway signaling during cell migration.

Figure 2-11. (A) The images show of RAW cells transfected with blue opsin-mCh and their responses to localized OA, in the presence of the following inhibitors - 10 μM gallein for Gβγ, 50 nM wortmannin for PI3K and 12.5 μM GSK 269962 for ROCK. Cells were incubated with the appropriate inhibitor/s for 30 mins at 37 °C prior to the imaging and OA. (I) Inhibition of Gβγ showed a complete termination of migration with complete cessation of LE and TE activities, suggesting that Gβγ is likely to control both LE and TE edge functions, and has the complete control over cell migration. (II) While the inhibition of PI3K terminated the migration, it resulted in the reduction of cell size indicating the existence of the contraction forces on the cytoskeleton. (III) While ROCK inhibition alone prevented complete cell migration with no TE retraction, but the LE activities remained unperturbed. (IV) Dual inhibition of PI3K and ROCK resulted in neither cell migration nor cell size change. (B) The histogram shows the corresponding percent cell surface changes. **, P <0.0001. (n=10). Scale bar, 10 μM.

Figure 2-12. RNA-seq data showing the relative expression of Geffectors in RAW cells.

2.4.7. Does intracellular calcium regulate the G mediated PIP3 generation and lamellepodia formation.

We understood that extracellular and intracellular calcium play a crucial role in promoting cell migration specifically in the TE retraction from cell migration data. To examine the extent of the effect of intracellular calcium at the LE, we examined the PIP3 generation in cells treated with BAPTA-AM. RAW cells expressing blue opsin-mTurquoise and AKT-

PH-mCh were incubated with BAPTA–AM for 30 mins at 37 °C. Both global and localized activation of blue opsin in RAW cells were performed to check for the accumulation of

AKT-PH-mCh on the PM. Upon OA, cytosolic AKT -PH-mCh sensor translocates to the

PM, indicating PIP3 generation. An orthogonal view shows AKT-PH-mCh dynamics of a cross section of the cell clearly demonstrating this translocation (Fig 2-13A). Plot shows the accumulation of AKT-PH-mCh on the PM in control and BAPTA-AM treated cells

(Fig 2-13B). Similarly, control and BAPTA-AM treated cells were subjected to localized

OA of blue opsin (Fig 2-13C). Plot shows the accumulation of AKT-PH-mCh on the PM near the region of OA in control and BAPTA-AM treated cells (Fig 2-13D). While both

41 conditions showed robust PIP3 production at the LE, complete cell migration was only observed in control cells. This indicates that altering the intracellular calcium concentration might affect the retraction process in migrating cells. Therefore, calcium signaling pathway seems to be particularly important for TE retraction.

Figure 2-13. (A) RAW cells expressing blue opsin and AKT-PH-mCh, untreated and incubated with cell permeable calcium chelator BAPTA-AM (5 μM, 30 mins incubation at 37 °C) respectively, show a robust PIP3 production on global OA of blue opsin. The kymographs of the cellular cross sections show the reduction of cytosolic and increase in PM bound AKT-PH-mCh. (B) The plot generated using the kymograph shows the corresponding increase in mCh intensity on the PM. (C) RAW cells expressing blue opsin and AKT-PH-mCh, untreated and incubated with cell permeable calcium chelator BAPTA- AM (5 μM, 30 mins incubation at 37 °C) respectively shows a robust localized PIP3 production accompanied with lamellepodia formation upon confined OA of blue opsin. Interestingly, cell incubated for 30 mins with 5 μM BAPTA AM at 37 °C, failed to migrate indicating that cytosolic calcium is required for the directional migration. (D) The plot shows the corresponding AKT-PH-mCh dynamics at the LE (n = 3). Scale bar, 10 μm.

2.4.8. Regulation of Ca-CaM mediated phosphorylation of MLC by G.

Evidence shows that calcium generated at the ER binds to calmodulin and the association of the Ca-CaM complex with the catalytic subunit of MLCK activates the kinase resulting in the phosphorylation of MLC105. Phosphorylation of MLC allows the myosin ATPase to be activated and accounts for the actomyosin contractility (Fig 2-14). Here, we examined if Gβγ actively regulates the MLC phosphorylation and TE retraction. RAW cells expressing blue opsin-mCh were imaged in the presence of a CaM antagonist A7-hydrochloride to check for cell migration. RAW Figure 2-14 cells were incubated with A7-hydrochloride for 30 mins at 37 °C. Calcium mediated actomyosin Upon OA of blue opsin, the inhibition of CaM resulted in the contractility inhibition of cell migration. However, the cells produced lamellepodia at the LE as observed in control cells but the retraction at the TE was inhibited (Fig 2-15A). Next, the effect of the inhibition of MLCK on cell migration was examined using its inhibitor ML-7 hydrochloride. RAW cells were incubated with ML-7 hydrochloride for 30 mins at 37 °C.

Upon OA, cells showed lamellepodia formation at the LE, little or no TE retraction was

43 observed (Fig 2-15B). The Histogram shows the distance travelled by the LE and TE, respectively when cells are treated with A-7 and ML-7 compared to that of the control cells

(Fig 2-15C). Inhibition of both CaM and MLCK did not have an effect on LE activities, lamellepodia formation while the cells showed no TE retraction or directional migrations.

Figure 2-15. (A) The images show a RAW cell transfected with blue opsin-mCh, incubated with 10 μM A-7 hydrochloride, a CaM antagonist for 30 mins at 37 °C, inhibiting the blue opsin induced cell migration, however with intact LE activities. (B) The images show RAW cell transfected with blue opsin-mCh, incubated with 10 μM ML-7 hydrochloride, a MLCK antagonist for 30 mins at 37 °C, inhibiting the retraction at the TE and cell migration, while showing unperturbed lamellepodia formation. (C) Plot shows the distance migrated by LE and TE in control, both CAM and MLCK inhibited cells. **, P <0.0001(n=10). Scale bar, 10 μm.

2.4.9. G mediated PIP2 Hydrolysis

Hydrolysis of PIP2 by specific PLC isoforms regulates a number of cellular processes including calcium release and DAG mediated PKC signaling106, 107. In mammalian cells two isoforms of the PLC (PLC  and) have been identified to mediate PIP2 hydrolysis107.

During PIP2 hydrolysis, PLC isoforms are activated by Gq subunit generated as a results of GPCR activation108. RNA-seq data show that RAW cells have an elevated expression of

PLCβ2, a Gβγ activatable PLC. Data also shows a considerable expression of CaM isoforms, MLCK and MLCP (MYTP1), suggesting the possibilities of Gβγ induced MLCK activation in RAW cells (Fig 2-16).

Figure 2-16. RT-PCR mapping of genes required for Gβγ to induce MLCK activation in RAW cells. The data is presented as the mean ± standard deviation. (n=3)

Next, we examined if Gαi coupled GPCR activation induces PIP2 hydrolysis through free

Gβγ. Here, HeLa cells expressing mCh-PH (PIP2 sensor) was examined by activating the endogenous α2-AR (Gi) using NE as the agonist. PIP2 hydrolysis is measured using

45 previously described translocation based mCh-PH sensor109. The working principle of the sensor is given in the Fig. 2-17.

Figure 2-17 Schematic representation of mCh-PH sensor activity. (1) GPCR in its inactive state and PIP2 attached to mCh-PH on the PM. (2) Activation of GPCR upon ligand addition triggers the G protein activation followed by dissociation. The free βγ activates PLC and catalyzes the hydrolysis of PIP2. mCh-PH now binds to IP3 with higher affinity and translocates to the cytosol. mCh-PH translocation is a direct measure of the extent of PIP2 hydrolysis (3) Molecular representation of PIP2 hydrolysis.

Activation of endogenous α2-AR resulted in PIP2 hydrolysis indicated by the translocation of mCh from the PM to cytosol (Fig 2-18A). Additionally, we performed similar experiments in RAW cells expressing blue opsin-mTurquoise and mCh-PH. Upon OA of blue opsin, a considerable amount of PIP2 hydrolysis was observed (Fig 2-18B). This observation demonstrates that, in addition to PIP2 hydrolysis mediated by Gq, Gi coupled GPCRs can also induce PIP2 hydrolysis upon Gαi-coupled GPCR activation. To validate this hypothesis further, we examined for the PIP2 hydrolysis in cells treated with

PTx to inhibit Gαi heterotrimer activation and gallein to prevent Gβγ effector interaction, respectively. PTx as well as gallein treated RAW cells showed no PIP2 response upon receptor activation (Fig 2-18C, D).

Figure 2-18. (A) The images show PIP2 dynamics in HeLa cells expressing mCh-PH. Upon activation of endogenous α2-AR with 400 µM NE, cells show PIP2 hydrolysis. This is observed by the translocation of the mCh-PH sensor into the cytosol. Plot shows the increase in the intensity of mCh-PH in the cytosol upon receptor activation by agonist addition. (B) Images show PIP2 dynamics in RAW expressing blue opsin-mTurquoise and mCh-PH. Upon OA of blue opsin, cells showed PIP2 hydrolysis. Plot shows the increase in the intensity of mCh-PH in the cytosol upon OA. (C), (D) images show RAW cells expressing blue opsin and mCh-PH, treated with PTx (0.05 μg/mL, overnight incubation at 37 °C) and gallein (10 µM, 30 mins incubation at 37 °C) and checked for the PIP2 hydrolysis by OA of blue opsin. Cells failed to show any PIP2 response. The plots show the corresponding mCh intensity in the cytosol. Scale bar, 10 μm.

2.4.10. Calcium mobilization in RAW cells is steered by activation of Gi coupled

GPCRs

Gq-coupled receptors activate PLC, which hydrolyzes membrane PIP2 into the second messenger’s IP3 and DAG. IP3 binds to its receptor, IP3R, on the ER to stimulate calcium release. Simultaneously DAG, together with mobilized calcium stimulates conventional

PKC110. Previously we have shown that activation of Gαi coupled receptors induces PIP2 hydrolysis (Fig 2-18). The results further show that Gi coupled GPCRs activation also evokes calcium mobilization in RAW cells. We have previously reported that c5aR1 receptors which belongs to the family of Gi coupled GPCR are endogenously expressed in RAW cells91. Cell permeable fluorescence calcium indicator, fluo4-AM can be used to measure cytosolic calcium responses with the excitation wavelength of 488 nm and emission at 515 nm. Activation of c5aR1 upon addition of 25 µM c5a resulted in a robust calcium increase in the cytosol (Fig 2-19A). Plot shows the increase in mean fluorescent intensity of fluo4 upon c5a addition. This proves that with the activation of Gαi pathway, an increase of cytosolic calcium can be observed. To further validate the Gαi pathway involvement and exclude the possibility of c5aR promiscuously activating Gαq heterotrimer, calcium mobilization was examined in RAW cells in the presence of PTx.

Upon addition of c5a, RAW cells failed to show Gαi induced calcium mobilization. The same cells upon addition of thapsigargin showed an increase in the cytosolic calcium (Fig

2-19B). Plot shows no calcium response upon c5a addition (black lines) and an increase in the calcium response upon thapsigargin addition (red lines). Ion channels in the PM are known to facilitate the movement of ions in and out of the cells111. In order to examine if the movement of extracellular calcium into the cytosol is resulting in the calcium response

49 in the cells, we examined c5aR ability to mobilize calcium in cells submerged in a calcium free buffer. While imaging cells pre-incubated with fluo4-AM, we used BAPTA at 50 s to chelate extracellular. Cells were then stimulated with c5a at 150 s which resulted in robust calcium response (Fig 2-19C). This data suggests that the calcium response produced by the cells is indeed due to the activation of c5aR1 receptors. Plot shows no calcium increase in the presence of BAPTA. Nevertheless, c5a addition showed a robust calcium response.

To examine if the cytosolic calcium mobilization is certainly induced by the Gαi medicated free Gβγ, RAW cells were incubated with gallein for 30 mins at 37 °C to inhibit Gβγ. Upon c5a addition cells failed to show calcium response (Fig 2-19D). This data suggest that the calcium mobilization in RAW cells is induced by free Gβγ interacting with PLC. When cells were treated with 2APB (IP3R inhibitor), cells also failed to mobilize calcium upon c5a addition (Fig 2-19E). This further ascertains that Gβγ induced PIP2 hydrolysis pathway is responsible for the calcium response. Furthermore, when cells were incubated in calcium free buffer and BAPTA, cells failed to produce calcium (Fig 2-19F). To demonstrate that the calcium mobilization observed is independent on Gβγ mediated activation of PI3K, cells were incubated with wortmannin, a PI3K inhibitor, for 30 mins at

37 °C and examined c5aR induced calcium mobilization. Inhibition of PI3K did not alter the c5a induced calcium mobilization (Fig 2-20A). We have shown that localized blue opsin (Gi) activation can result in directional migration due to its ability to spatiotemporally control GPCR signaling (Fig 2-7A). We have also shown that perturbing the extracellular and intracellular calcium resulted in curtailed cell migration (Fig 2-7E, F).

Here, activation of Gαi coupled blue opsin was examined to see if it induces calcium mobilization as well. Blue opsin expressing RAW cells were incubated with fluo4-AM

50 were imaged with 488 nm light and upon retinal addition, showed calcium response in a similar manner to that of c5a addition (Fig 2-20B). Blue opsin absorption spectra has a small overlap with the excitation wavelength used for EGFP (488 nm) while that of the

YFP (515 nm) and mCh (594 nm) excitations wavelengths showed no overlap (Fig 2-20).

Due to this overlap, imaging fluo-4, which has similar spectral properties to that of GFP, can result in unintended activation of blue opsin when bound 11-cis retinal is present. This limitation of blue opsin allows us to use retinal as a chemical ligand here in the presence of 488 nm excitation light.

C5a

Before c5a After c5a

PTx

With PTx After c5a ligand After Thapsigargin

free buffer free

Basal After BAPTA After c5a

D E

Gallein 2APB

Before c5a After c5a Before c5a After c5a

BAPTA

Before c5a After c5a

Figure 2-19. (A) Representative images of RAW cells showing calcium mobilization after addition of agonist (c5a, dose= 25 μM). Agonist was added at 20 s. Time course of calcium response after addition of c5a agonist. (B) Representative images of RAW cells treated with PTx showing calcium mobilization before and after thapsigargin (500 nM) addition. Time course of calcium response in the presence of PTx before (black) and after thapsigargin addition (red). (C) Representative images of RAW cells showing calcium mobilization when imaged in calcium free buffer. Time course of calcium response in the presence of BAPTA and after c5a agonist addition. (D), (E), (F) Representative images of RAW cells showing no calcium mobilization when treated with gallein, BAPTA and 2APB. Time course of calcium response in the presence of inhibitors.

Wortmannin Before c5a After c5a

Retinal

Blue Opsin Fluo4 before Fluo4 after mCh retinal addition retinal addition

Figure 2-20. (A) Representative images of RAW cells showing calcium mobilization in the presence of wortmannin (inhibitor of PI3K). Time course of calcium response before and after addition of c5a agonist. (B) Representative images of RAW cells transiently expressing blue opsin-mCh showing calcium mobilization upon OA. Time course of calcium response before (black) and after retinal (agonist binding to blue opsin which induces the OA upon photo activation) addition (red). Scale bar, 10 μm.

Figure 2-21. Plot shows the overlap between the absorption spectrum of blue opsin (blue) and the excitation spectrum of the EGFP. The wavelength sensitivity of blue opsin is such that, EGFP excitation at 488 nm is sufficient to activate blue opsin.

2.5. Discussion

Cell migration is a result from a coordinated regulation of multiple cellular events, involving extension at the LE and contraction of its cytoskeleton at the TE. However, the

53 mechanisms used by cells to transduce the signals from the LE to the TE to induce its retraction facilitating directional migration are yet to be discovered. To study these mechanisms, confined GPCR activation at the LE is required and therefore, we employed an optogenetic approach to limit GPCR activity to a selected area of the cell. For this, we employed light activatable human blue opsin as the trigger to turn on and off G protein activity inducing Gi pathway in single cells. Using the G9 translocation assay, we showed that blue opsin is capable of activating G proteins only in the LE. Subsequently, using blue opsin, we accomplished the directional migration in RAW cells. Spatially confined activation of blue opsin with 445 nm blue light induced similar translocation results when compared with the ligand activated CXCR4 receptors. This indicates that blue opsin can be used as a substitute to mimic the CXCR4 activity in cells to induce cell migration. Inhibition of cell migration in the presence of PTx show that cell migration is indeed mediated by Gi coupled receptors. Furthermore, cessation of cell migration in the presence of gallein proves that free G plays a major role in cell migration. From the translocation data pertaining to Gi, Gs and Gq presented here, it can be argued that

Gs coupled receptors can also mediate cell migration, because their activation also can generate free G. Data shows the inability of cells to migrate upon OA of Gs coupled

CrBlue. When compared to the cell migration induced by the activation of Gi coupled blue opsin, CrBlue expressing cells neither showed lamellepodia formation at the LE nor retraction at the TE. Evidence shows that cAMP generated upon activation of Gs coupled

GPCR is known to inhibit cell migration through cAMP-PKA pathway. PKA can directly phosphorylate RhoA at Ser188 near its C-terminus and this can result in inhibition of interaction of RhoA with its effector ROCK leading to negative effects on actomyosin

54 contractility112. RhoA governs contractility-dependent processes in retraction during migration113.

Free Gcontrols many cellular processes by their interaction with downstream effector molecules. One of the important Gmediated cellular process is the lamellepodia formation at the LE during cell migration. RT-PCR data of RAW cells showed good expression of a Gactivatable RhoGEF (FLJ00018) which is known to induce lamellepodia through Rac1 and Cdc42. To validate the hypothesis of Gbeing a primary mediator at the LE and also for the TE signaling during cell migration, we checked for cell migration in cells independent of free Gby employing an optically activatable domain of PI3K, mCh-CRY-iSH2. OA of mCh-CRY-iSH2 only resulted in PI3K accumulation at the LE but little or no cell migration, especially lacks the TE retraction indicating that there is an alternate Gmediated TE retraction pathway. This is further confirmed when OA of blue opsin in cells treated with wortmannin showed no lamellepodia formation but whole cell contraction. Furthermore, OA of blue opsin in cells treated with GSK 269962 showed lamellepodia formation, but failed to show TE retraction. This suggested that Ggoverns

TE retraction by activating RhoGEFs responsible for TE retraction. Data from RT-PCR of

RAW cells show considerable expression of a Gactivatable RhoGEF, p114RhoGEF which activates RhoA and subsequently ROCK. Inhibition of ROCK inhibits the MLCP and thereby promotes the phosphorylation of MLC. This resulted in MLCK activation, finally resulting in the actomyosin contraction and TE retraction. Even though the pathway

GβγRhoGEFRhoAROCK is essential for the deactivation of the MLCP, the chief signaling pathway required for TE retraction is through the phosphorylation of MLC by

MLCK.

Evidence suggested that with the increase in intracellular concentration of calcium, calcium binds to calmodulin forming Ca-CaM complex. Ca-CaM binds with high affinity to the

M13 catalytic subunit domain of MLCK, activates MLCK. Activated MLCK induce phosphorylation at Ser19 on the regulatory light chain of MLC114. By making use of the commercially available inhibitors for proteins involved in the GβγCa-

CaMMLCKMLC pathway, we checked for the role of calcium in the retraction of the

TE. RT-PCR data of RAW cells show considerable expression of downstream proteins like

CaM, MLCK and MYPT1 (MLCP targeting protein). Data from cell migration studies performed on cells treated with CaM inhibitor, A-7 hydrochloride and MLCK inhibitor,

ML-7 hydrochloride showed good lamellepodia formation at the LE but no retraction at the TE which showed that CaM, MLCK and MLCP play a crucial role in Gmediated

TE retraction.

Furthermore, RT-PCR data also showed expression of Gβγ activatable PLCβ2Data from

α2AR mediated PIP2 hydrolysis point out to the prospect of Gi mediated PIP2 hydrolysis in RAW cells. To examine if Gi coupled GPCRs can mobilize intracellular calcium upon activation, endogenous c5aR1 receptors are activated with c5a resulting in robust calcium mobilization. This illustrates the existence of Gi-βγPLCβIP3IP3RCalcium pathway in RAW cells. This was further proved when cells treated with PTx failed to show calcium response upon c5a addition. Gallein treated cells also failed to show calcium response upon c5a addition showing that free Gβγ interaction with PLCβ is required for calcium mobilization. Furthermore, use of 2APB which bocks the IP3 receptors on the ER showed no calcium mobilization upon c5a addition further strengthening the hypothesis for existence of Gβγ mediated calcium signaling pathway in RAW cells. We employed

56 extracellular calcium chelator, BAPTA to eliminate the possibility of the flow of extracellular calcium into the cells and result in the calcium response. Experiments performed on cells cultured in BAPTA added medium showed robust calcium response upon c5a addition.

To further ascertain the role of Gβγ mediated calcium signaling in the TE retraction, cell migration was observed in cells treated with intracellular calcium chelators and SERCA pump blocker, thapsigargin. Use of BAPTA-AM resulted in complete chelation of the intracellular calcium and upon OA of blue opsin, cells showed unperturbed lamellepodia formation at the LE while displaying minimum or no TE retraction. Also, use of thapsigargin to block calcium release from ER into the cytosol yielded similar results to that of BAPTA-AM. These experiments demonstrate directly that intracellular calcium is essential for TE retraction.

Based on the data, we propose a new signaling pathway for complete cell migration solely governed by Gβγ and calcium (Fig 2-21). At the LE, Gβγ pathway involves the activation of PI3K and lamellepodia formation. At the TE, two systems for actomyosin contraction mediated Gβγ for the retraction involving the activation of RhoGEFs and PLC.

Simultaneous activity of both the systems are vital as

Gβγp114RhoGEFRhoAROCK pathway leads to an increase in calcium sensitivity due to the inhibition of the activity of MLCP, thereby, facilitating the phosphorylation of

MLC by MLCK. GβγPLCβCa-CaMMLCK pathway is essential for the MLC phosphorylation induced actomyosin contractility and required for TE retraction.

Figure 2-22. Proposed signaling pathway for complete cell migration mediated by G

2.6. Conclusion

From the data presented in this thesis, we conclude that reduction in cytosolic calcium activity inhibits cell migration by decreasing the rate of MLC phosphorylation. These observations suggest a novel calcium-dependent mechanism mediated by free G for regulating the TE retraction during migration. Interestingly, we have also shown that

Gcan solely govern complete cell migration by mediating the lamellepodia formation at the LE by activating PI3K and retraction at the TE by activation of RhoGEF and

PLCcalcium. Therefore, data presented here not only can help to unravel the intricacies

58 of cell migration with Gand calcium as mediators, but also uncover new molecular targets for therapy development for pathological conditions including metastasis.

References

1. Bockaert, J., and Pin, J. P. (1999) Molecular tinkering of G protein‐coupled receptors: an evolutionary success, The EMBO J. 18, 1723-1729.

2. Fredriksson, R., Lagerström, M. C., Lundin, L.-G., and Schiöth, H. B. (2003) The G-protein-coupled receptors in the human genome form five main families. Phylogenetic analysis, paralogon groups, and fingerprints, Mol. Pharmacol. 63, 1256-1272.

3. Vassilatis, D. K., Hohmann, J. G., Zeng, H., Li, F., Ranchalis, J. E., Mortrud, M. T., Brown, A., Rodriguez, S. S., Weller, J. R., and Wright, A. C. (2003) The G protein-coupled receptor repertoires of human and mouse, Proc. Natl. Acad. Sci. 100, 4903-4908.

4. Palczewski, K., Kumasaka, T., Hori, T., Behnke, C. A., Motoshima, H., Fox, B. A., Le Trong, I., Teller, D. C., Okada, T., and Stenkamp, R. E. (2000) Crystal structure of rhodopsin: AG protein-coupled receptor, Science 289, 739-745.

5. Pierce, K. L., Premont, R. T., and Lefkowitz, R. J. (2002) Seven-transmembrane receptors, Nat. Rev. Mol. Cell Biol. 3, 639-650.

6. Schöneberg, T., Schulz, A., and Gudermann, T. (2002) The structural basis of G- protein-coupled receptor function and dysfunction in human diseases, In Rev. Physiol. Bioch. P., pp 144-227.

7. Park, J. H., Scheerer, P., Hofmann, K. P., Choe, H.-W., and Ernst, O. P. (2008) Crystal structure of the ligand-free G-protein-coupled receptor opsin, Nature 454, 183-187.

8. Cherezov, V., Rosenbaum, D. M., Hanson, M. A., Rasmussen, S. G., Thian, F. S., Kobilka, T. S., Choi, H.-J., Kuhn, P., Weis, W. I., and Kobilka, B. K. (2007) High- resolution crystal structure of an engineered human β2-adrenergic G protein– coupled receptor, Science 318, 1258-1265.

9. Hollmann, M. W., Strumper, D., Herroeder, S., and Durieux, M. E. (2005) Receptors, G proteins, and their interactions, Anesthesiology 103, 1066-1078.

10. Cabrera-Vera, T. M., Vanhauwe, J., Thomas, T. O., Medkova, M., Preininger, A., Mazzoni, M. R., and Hamm, H. E. (2003) Insights into G protein structure, function, and regulation, Endocr. Rev. 24, 765-781.

11. Birnbaumer, L. (1990) Transduction of receptor signal into modulation of effector activity by G proteins: the first 20 years or so, The FASEB J. 4, 3178-3188.

12. Oldham, W. M., and Hamm, H. E. (2007) How do receptors activate G proteins?, Adv. Protein Chem. 74, 67-93.

13. Simon, M. I., Strathmann, M. P., and Gautam, N. (1991) Diversity of G proteins in signal transduction, Science 252, 802.

14. Kjeldgaard, M., Nyborg, J., and Clark, B. (1996) The GTP binding motif: variations on a theme, The FASEB J. 10, 1347-1368.

15. Hurowitz, E. H., Melnyk, J. M., Chen, Y.-J., Kouros-Mehr, H., Simon, M. I., and Shizuya, H. (2000) Genomic characterization of the human heterotrimeric G protein α, β, and γ subunit genes, DNA Res. 7, 111-120.

16. Smrcka, A. (2008) G protein βγ subunits: central mediators of G protein-coupled receptor signaling, Cell. Mol. Life Sci. 65, 2191-2214.

17. Pronin, A. N., and Gautam, N. (1992) Interaction between G-protein beta and gamma subunit types is selective, Proc. Natl. Acad. Sci. 89, 6220-6224.

18. Lambright, D. G., Sondek, J., Bohm, A., and Skiba, N. P. (1996) The 2.0 angstrom crystal structure of a heterotrimeric G protein, Nature 379, 311.

19. Wall, M. A., Coleman, D. E., Lee, E., Iñiguez-Lluhi, J. A., Posner, B. A., Gilman, A. G., and Sprang, S. R. (1995) The structure of the G protein heterotrimer Giα1β1γ2, Cell 83, 1047-1058.

20. Sondek, J., Bohm, A., Lambright, D. G., Hamm, H. E., and Sigler, P. B. (1996) Crystal structure of a GA protein (Beta gamma) dimer at 2.1 angstrom resolution, Nature 379, 369.

21. Zhang, F. L., and Casey, P. J. (1996) Protein prenylation: molecular mechanisms and functional consequences, Annu. Rev. Biochem 65, 241-269.

22. Oldham, W. M., and Hamm, H. E. (2006) Structural basis of function in heterotrimeric G proteins, Q. Rev. Biophys. 39, 117-166.

23. Higgins, J. B., and Casey, P. J. (1996) The role of prenylation in G-protein assembly and function, Cell. Signal. 8, 433-437.

24. Lambright, D. G., Noel, J. P., Hamm, H. E., and Sigler, P. B. (1994) Structural determinants for activation of the alpha-subunit of a heterotrimeric G protein, Nature 369, 621.

25. Logothetis, D. E., Kurachi, Y., Galper, J., Neer, E. J., and Clapham, D. E. (1987) The ßy subunits of GTP-binding proteins activate the muscarinic K+ channel in heart, Nature 325, 321-326.

26. Pitcher, J. A., Inglese, J., Higgins, J. B., Arriza, J. L., Casey, P. J., Kim, C., Benovic, J. L., Kwatra, M. M., Caron, M. G., and Lefkowitz, R. J. (1992) Role of beta- gamma subunits of G proteins in targeting the beta-adrenergic receptor kinase to membrane-bound receptors, Science 257, 1264-1268.

27. Camps, M., Carozzi, A., Schnabel, P., Scheer, A., Parker, P. J., and Gierschik, P. (1992) Isozyme-selective stimulation of phospholipase C-beta 2 by G protein beta gamma-subunits, Nature 360, 684.

28. Smrcka, A. V., and Sternweis, P. C. (1993) Regulation of purified subtypes of phosphatidylinositol-specific phospholipase C beta by G protein alpha and beta gamma subunits, J. Biol. Chem. 268, 9667-9674.

29. Park, D., Jhon, D.-Y., Lee, C., Lee, K.-H., and Rhee, S. G. (1993) Activation of phospholipase C isozymes by G protein beta gamma subunits, J. Biol. Chem. 268, 4573-4576.

30. Wei-Jen, T., and Gilman, A. G. (1991) Type-specific regulation of adenylyl cyclase by G protein (beta gamma) subunits, Science 254, 1500.

31. Taussig, R., Tang, W.-J., Hepler, J. R., and Gilman, A. G. (1994) Distinct patterns of bidirectional regulation of mammalian adenylyl cyclases, J. Biol. Chem. 269, 6093-6100.

32. Sunahara, R. K., and Taussig, R. (2002) Isoforms of mammalian adenylyl cyclase: multiplicities of signaling, Mol. Interventions 2, 168.

33. Ikeda, S. R. (1996) Voltage-dependent modulation of N-type calcium channels by G-protein (Beta gamma) subunits, Nature 380, 255.

34. Herlitze, S., Garcia, D. E., Mackie, K., and Hille, B. (1996) Modulation of Ca2 positive channels by G-protein (Beta gamma) subunits, Nature 380, 258.

35. Stephens, L., Smrcka, A., Cooke, F., Jackson, T., Sternweis, P., and Hawkins, P. (1994) A novel phosphoinositide 3 kinase activity in myeloid-derived cells is activated by G protein βγ subunits, Cell 77, 83-93.

36. Gerachshenko, T., Blackmer, T., Yoon, E.-J., Bartleson, C., Hamm, H. E., and Alford, S. (2005) Gβγ acts at the C terminus of SNAP-25 to mediate presynaptic inhibition, Nat. Neurosci. 8, 597-605.

37. Welch, H. C., Coadwell, W. J., Ellson, C. D., Ferguson, G. J., Andrews, S. R., Erdjument-Bromage, H., Tempst, P., Hawkins, P. T., and Stephens, L. R. (2002) P- Rex1, a PtdIns (3, 4, 5) P 3-and Gβγ-regulated guanine-nucleotide exchange factor for Rac, Cell 108, 809-821.

38. Niu, J., Profirovic, J., Pan, H., Vaiskunaite, R., and Voyno-Yasenetskaya, T. (2003) G Protein βγ Subunits Stimulate p114RhoGEF, a Guanine Nucleotide Exchange Factor for RhoA and Rac1, Circul. Res. 93, 848-856.

39. Ueda, H., Nagae, R., Kozawa, M., Morishita, R., Kimura, S., Nagase, T., Ohara, O., Yoshida, S., and Asano, T. (2008) Heterotrimeric G protein βγ subunits stimulate FLJ00018, a guanine nucleotide exchange factor for Rac1 and Cdc42, J. Biol. Chem. 283, 1946-1953.

40. Hepler, J. R., and Gilman, A. G. (1992) G proteins, Trends Biochem. Sci 17, 383- 387.

41. Clapham, D. E., and Neer, E. J. (1993) New roles for G-protein beta gamma-dimers in transmembrane signalling, Nature 365, 403.

42. Gether, U., Asmar, F., Meinild, A. K., and Rasmussen, S. G. (2002) Structural basis for activation of G‐protein‐coupled receptors, Pharmacol. Toxicol. 91, 304-312.

43. Berman, D. M., Wilkie, T. M., and Gilman, A. G. (1996) GAIP and RGS4 are GTPase-activating proteins for the G i subfamily of G protein α subunits, Cell 86, 445-452.

44. Hirsch, E., Katanaev, V. L., Garlanda, C., Azzolino, O., Pirola, L., Silengo, L., Sozzani, S., Mantovani, A., Altruda, F., and Wymann, M. P. (2000) Central role for G protein-coupled phosphoinositide 3-kinase γ in inflammation, Science 287, 1049-1053.

45. Mantovani, A. (2009) Cancer: inflaming metastasis, Nature 457, 36-37.

46. Fidler, I. J. (2003) The pathogenesis of cancer metastasis: the'seed and soil'hypothesis revisited, Nat. Rev. Cancer 3, 453-458.

47. Hanahan, D., and Weinberg, R. A. (2011) Hallmarks of cancer: the next generation, Cell 144, 646-674.

48. Chambers, A. F., Groom, A. C., and MacDonald, I. C. (2002) Metastasis: dissemination and growth of cancer cells in metastatic sites, Nat. Rev. Cance 2, 563-572.

49. Iaccarino, G., and Koch, W. J. (2003) Transgenic mice targeting the heart unveil G protein-coupled receptor kinases as therapeutic targets, Assay Drug Dev. Technol. 1, 347-355.

50. Lefkowitz, R. J., Rockman, H. A., and Koch, W. J. (2000) Catecholamines, cardiac beta-adrenergic receptors, and heart failure, Circulation 101, 1634-1637.

51. Dhein, S., Van Koppen, C. J., and Brodde, O.-E. (2001) Muscarinic receptors in the mammalian heart, Pharmacol. Res. 44, 161-182.

52. Xiao, R.-P. (2001) Beta-adrenergic signaling in the heart: dual coupling of the beta2-adrenergic receptor to G (s) and G (i) proteins, Sci Stke 104, re15.

53. Koch, W. J., Rockman, H. A., Samama, P., and Hamilton, R. (1995) Cardiac function in mice overexpressing the beta-adrenergic receptor kinase or a BetaARK inhibitor, Science 268, 1350.

54. Rockman, H. A., Chien, K. R., Choi, D.-J., Iaccarino, G., Hunter, J. J., Ross, J., Lefkowitz, R. J., and Koch, W. J. (1998) Expression of a β-adrenergic receptor kinase 1 inhibitor prevents the development of myocardial failure in gene-targeted mice, Proc. Natl. Acad. Sci 95, 7000-7005.

55. Lorenz, K., Schmitt, J. P., Schmitteckert, E. M., and Lohse, M. J. (2009) A new type of ERK1/2 autophosphorylation causes cardiac hypertrophy, Nat. Med. 15, 75- 83.

56. Ridley, A. J., Schwartz, M. A., Burridge, K., Firtel, R. A., Ginsberg, M. H., Borisy, G., Parsons, J. T., and Horwitz, A. R. (2003) Cell migration: integrating signals from front to back, Science 302, 1704-1709.

57. Van Haastert, P. J., and Devreotes, P. N. (2004) Chemotaxis: signalling the way forward, Nat. Rev. Mol. Cell Biol. 5, 626-634.

58. Lauffenburger, D. A., and Horwitz, A. F. (1996) Cell migration: a physically integrated molecular process, Cell 84, 359-369.

59. Tschumperlin, D. J. (2013) Fibroblasts and the ground they walk on, Physiology 28, 380-390.

60. Lamalice, L., Le Boeuf, F., and Huot, J. (2007) Endothelial cell migration during angiogenesis, Circul. Res. 100, 782-794.

61. Etienne-Manneville, S., and Hall, A. (2002) Rho GTPases in cell biology, Nature 420, 629-635.

62. Dianqing, W. (2005) Signaling mechanisms for regulation of chemotaxis, Cell Res. 15, 52-56.

63. Bokoch, G. M. (2000) Regulation of cell function by Rho family GTPases, Immunol. Res. 21, 139-148.

64. Srinivasan, S., Wang, F., Glavas, S., Ott, A., Hofmann, F., Aktories, K., Kalman, D., and Bourne, H. R. (2003) Rac and Cdc42 play distinct roles in regulating PI (3, 4, 5) P3 and polarity during neutrophil chemotaxis, J. Cell Biol. 160, 375-385.

65. Prasad, S. V. N., Esposito, G., Mao, L., Koch, W. J., and Rockman, H. A. (2000) Gβγ-dependent phosphoinositide 3-kinase activation in hearts with in vivo pressure overload hypertrophy, J. Biol. Chem. 275, 4693-4698.

66. Chung, C. Y., Funamoto, S., and Firtel, R. A. (2001) Signaling pathways controlling cell polarity and chemotaxis, Trends Biochem. Sci 26, 557-566.

67. Kölsch, V., Charest, P. G., and Firtel, R. A. (2008) The regulation of cell motility and chemotaxis by phospholipid signaling, J. Cell Sci. 121, 551-559.

68. Dong, X., Mo, Z., Bokoch, G., Guo, C., Li, Z., and Wu, D. (2005) P-Rex1 is a primary Rac2 guanine nucleotide exchange factor in mouse neutrophils, Curr. Biol. 15, 1874-1879.

69. Kranenburg, O., Poland, M., van Horck, F. P., Drechsel, D., Hall, A., and Moolenaar, W. H. (1999) Activation of RhoA by lysophosphatidic acid and Gα12/13 subunits in neuronal cells: induction of neurite retraction, Mol. Biol. Cell 10, 1851-1857.

70. Dutt, P., Nguyen, N., and Toksoz, D. (2004) Role of Lbc RhoGEF in Gα12/13- induced signals to Rho GTPase, Cell. Signal. 16, 201-209.

71. Kamai, T., Tsujii, T., Arai, K., Takagi, K., Asami, H., Ito, Y., and Oshima, H. (2003) Significant association of Rho/ROCK pathway with invasion and metastasis of bladder cancer, Clin. Cancer. Res. 9, 2632-2641.

72. Bhadriraju, K., Yang, M., Ruiz, S. A., Pirone, D., Tan, J., and Chen, C. S. (2007) Activation of ROCK by RhoA is regulated by cell adhesion, shape, and cytoskeletal tension, Exp. Cell Res. 313, 3616-3623.

73. Totsukawa, G., Yamakita, Y., Yamashiro, S., Hartshorne, D. J., Sasaki, Y., and Matsumura, F. (2000) Distinct roles of ROCK (Rho-kinase) and MLCK in spatial regulation of MLC phosphorylation for assembly of stress fibers and focal adhesions in 3T3 fibroblasts, J. Cell Biol. 150, 797-806.

74. Niggli, V. (1999) Rho‐kinase in human neutrophils: a role in signalling for myosin light chain phosphorylation and cell migration, FEBS Lett. 445, 69-72.

75. Maekawa, M., Ishizaki, T., Boku, S., Watanabe, N., Fujita, A., Iwamatsu, A., Obinata, T., Ohashi, K., Mizuno, K., and Narumiya, S. (1999) Signaling from Rho to the actin cytoskeleton through protein kinases ROCK and LIM-kinase, Science 285, 895-898.

76. Carter, M., and Shieh, J. (2010) Visualizing nervous system function, Guide to Research Techniques in Neuroscience, Elsevier Inc., Canada, 169-189.

77. Boyden, E. S., Zhang, F., Bamberg, E., Nagel, G., and Deisseroth, K. (2005) Millisecond-timescale, genetically targeted optical control of neural activity, Nat. Neurosci. 8, 1263-1268.

78. Karunarathne, W. A., Giri, L., Kalyanaraman, V., and Gautam, N. (2013) Optically triggering spatiotemporally confined GPCR activity in a cell and programming neurite initiation and extension, Proc. Natl. Acad. Sci. 110, E1565-E1574.

79. Nichols, C., and Lopatin, A. (1997) Inward rectifier potassium channels, Annu. Rev. Physiol. 59, 171-191.

80. Fogg, V. C., Azpiazu, I., Linder, M. E., Smrcka, A., Scarlata, S., and Gautam, N. (2001) Role of the γ subunit prenyl moiety in G protein βγ complex interaction with phospholipase Cβ, J. Biol. Chem. 276, 41797-41802.

81. He, C., Yan, X., Zhang, H., Mirshahi, T., Jin, T., Huang, A., and Logothetis, D. E. (2002) Identification of critical residues controlling G protein-gated inwardly rectifying K+ channel activity through interactions with the βγ subunits of G proteins, J. Biol. Chem. 277, 6088-6096.

82. van Blesen, T., Hawes, B. E., Luttrell, D. K., Krueger, K. M., Touhara, K., Porfflri, E., Sakaue, M., Luttrell, L. M., and Lefkowitz, R. J. (1995) Receptor-tyrosine- kinase-and G [beta][gamma]-mediated MAP kinase activation by a common signalling pathway, Nature 376, 781.

83. Baldassarre, M., Pompeo, A., Beznoussenko, G., Castaldi, C., Cortellino, S., McNiven, M. A., Luini, A., and Buccione, R. (2003) Dynamin participates in focal extracellular matrix degradation by invasive cells, Mol. Biol. Cell 14, 1074-1084.

84. Yamaguchi, H., Lorenz, M., Kempiak, S., Sarmiento, C., Coniglio, S., Symons, M., Segall, J., Eddy, R., Miki, H., and Takenawa, T. (2005) Molecular mechanisms of invadopodium formation, J. Cell Biol. 168, 441-452.

85. Yamaguchi, H., Wyckoff, J., and Condeelis, J. (2005) Cell migration in tumors, Curr. Opin. Cell Biol. 17, 559-564.

86. Xie, Y., Wolff, D. W., Wei, T., Wang, B., Deng, C., Kirui, J. K., Jiang, H., Qin, J., Abel, P. W., and Tu, Y. (2009) Breast cancer migration and invasion depend on proteasome degradation of regulator of G-protein signaling 4, Cancer Res. 69, 5743-5751.

87. Phillips, R. J., Mestas, J., Gharaee-Kermani, M., Burdick, M. D., Sica, A., Belperio, J. A., Keane, M. P., and Strieter, R. M. (2005) Epidermal growth factor and hypoxia-induced expression of CXC chemokine receptor 4 on non-small cell lung cancer cells is regulated by the phosphatidylinositol 3- kinase/PTEN/AKT/mammalian target of rapamycin signaling pathway and activation of hypoxia inducible factor-1α, J. Biol. Chem. 280, 22473-22481.

88. Murakami, T., Maki, W., Cardones, A. R., Fang, H., Kyi, A. T., Nestle, F. O., and Hwang, S. T. (2002) Expression of CXC chemokine receptor-4 enhances the pulmonary metastatic potential of murine B16 melanoma cells, Cancer Res. 62, 7328-7334.

89. Ottaiano, A., di Palma, A., Napolitano, M., Pisano, C., Pignata, S., Tatangelo, F., Botti, G., Acquaviva, A. M., Castello, G., and Ascierto, P. A. (2005) Inhibitory effects of anti-CXCR4 antibodies on human colon cancer cells, Cancer Immunol., Immunother. 54, 781-791.

90. Heizmann, C. W., and Hunzlker, W. (1991) Intracellular calcium-binding proteins: more sites than insights, Trends Biochem. Sci 16, 98-103.

91. Senarath, K., Ratnayake, K., Siripurapu, P., Payton, J. L., and Karunarathne, A. (2016) Reversible G Protein βγ9 Distribution-Based Assay Reveals Molecular Underpinnings in Subcellular, Single-Cell, and Multicellular GPCR and G Protein Activity, Anal. Chem. 88, 11450-11459.

92. Karunarathne, W. A., O'Neill, P. R., and Gautam, N. (2015) Subcellular optogenetics–controlling signaling and single-cell behavior, J. Cell Sci. 128, 15-25.

93. Wettschureck, N., and Offermanns, S. (2005) Mammalian G proteins and their cell type specific functions, Physiol. Rev. 85, 1159-1204.

94. Karunarathne, W. A., O’Neill, P. R., Martinez-Espinosa, P. L., Kalyanaraman, V., and Gautam, N. (2012) All G protein βγ complexes are capable of translocation on receptor activation, Biochem. Biophys. Res. Commun. 421, 605-611.

95. Arai, H., Tsou, C.-L., and Charo, I. F. (1997) Chemotaxis in a lymphocyte cell line transfected with CC chemokine receptor 2B: evidence that directed migration is mediated by βγ dimers released by activation of Gαi-coupled receptors, Proc. Natl. Acad. Sci. 94, 14495-14499.

96. Chen, L., Zhang, J. J., and Huang, X.-Y. (2008) cAMP inhibits cell migration by interfering with Rac-induced lamellipodium formation, J. Biol. Chem. 283, 13799- 13805.

97. Mangmool, S., and Kurose, H. (2011) Gi/o protein-dependent and-independent actions of pertussis toxin (PTX), Toxins 3, 884-899.

98. West, R., Moss, J., Vaughan, M., Liu, T., and Liu, T.-Y. (1985) Pertussis toxin- catalyzed ADP-ribosylation of transducin. Cysteine 347 is the ADP-ribose acceptor site, J. Biol. Chem. 260, 14428-14430.

99. Davis, T. L., Bonacci, T. M., Sprang, S. R., and Smrcka, A. V. (2005) Structural and molecular characterization of a preferred protein interaction surface on G protein βγ subunits, Biochemistry 44, 10593-10604.

100. Giri, L., Patel, A. K., Karunarathne, W. A., Kalyanaraman, V., Venkatesh, K., and Gautam, N. (2014) A G-protein subunit translocation embedded network motif underlies GPCR regulation of calcium oscillations, Biophys. J. 107, 242-254.

101. Liang, F., and Sze, H. (1998) A high-affinity Ca2+ pump, ECA1, from the endoplasmic reticulum is inhibited by cyclopiazonic acid but not by thapsigargin, Plant Physiol. 118, 817-825.

102. Manning, B. D., and Cantley, L. C. (2007) AKT/PKB signaling: navigating downstream, Cell 129, 1261-1274.

103. Wong, K.-K., Engelman, J. A., and Cantley, L. C. (2010) Targeting the PI3K signaling pathway in cancer, Curr. Opin. Genet. Dev. 20, 87-90.

104. Karunarathne, W. A., Giri, L., Patel, A. K., Venkatesh, K. V., and Gautam, N. (2013) Optical control demonstrates switch-like PIP3 dynamics underlying the initiation of immune cell migration, Proc. Natl. Acad. Sci.110, E1575-E1583.

105. Hori, M., and Karaki, H. (1998) Regulatory mechanisms of calcium sensitization of contractile elements in smooth muscle, Life Sci. 62, 1629-1633.

106. Tao, F., Dag, S., Wang, L.-W., Liu, Z., Butcher, D. R., Bluhm, H., Salmeron, M., and Somorjai, G. A. (2010) Break-up of stepped platinum catalyst surfaces by high CO coverage, Science 327, 850-853.

107. Wong, R., Hadjiyanni, I., Wei, H.-C., Polevoy, G., McBride, R., Sem, K.-P., and Brill, J. A. (2005) PIP2 hydrolysis and calcium release are required for cytokinesis in Drosophila spermatocytes, Curr. Biol. 15, 1401-1406.

108. Lee, C. H., Park, D., Wu, D., Rhee, S., and Simon, M. I. (1992) Members of the Gq alpha subunit gene family activate phospholipase C beta isozymes, J. Biol. Chem. 267, 16044-16047.

109. Chisari, M., Saini, D. K., Cho, J.-H., Kalyanaraman, V., and Gautam, N. (2009) G protein subunit dissociation and translocation regulate cellular response to receptor stimulation, PLoS One 4, e7797.

110. LaMorte, V., Harootunian, A., Spiegel, A., Tsien, R., and Feramisco, J. (1993) Mediation of growth factor induced DNA synthesis and calcium mobilization by Gq and Gi2, J. Cell Biol.121, 91-99.

111. Adams, D. J., Barakeh, J., Laskey, R., and Van Breemen, C. (1989) Ion channels and regulation of intracellular calcium in vascular endothelial cells, The FASEB J.3, 2389-2400.

112. Dong, J.-M., Leung, T., Manser, E., and Lim, L. (1998) cAMP-induced morphological changes are counteracted by the activated RhoA small GTPase and the Rho kinase ROKα, J. Biol. Chem. 273, 22554-22562.

113. Worthylake, R. A., Lemoine, S., Watson, J. M., and Burridge, K. (2001) RhoA is required for monocyte tail retraction during transendothelial migration, J. Cell Biol. 154, 147-160.

114. Somlyo, A. P., and Somlyo, A. V. (1994) Signal transduction and regulation in smooth muscle, Nature 372, 231.

Appendix: RT-PCR primers

Table 1: RAW 264.7 gene specific primers for RT-PCR