A Dissertation Entitled Interrogation of GPCR-G Protein Signaling Using

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A Dissertation

entitled

Interrogation of GPCR-G Protein Signaling using Novel Optogenetic Tools

Kanishka D Senarath

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

Doctor of Philosophy Degree in Chemistry

______Dr. Ajith Karunarathne, Committee Chair

______Dr. Donald Ronning, Committee Member

______Dr. Dragan Isailovic, Committee Member

______Dr. Song-Tao Liu, Committee Member

______Dr. Cyndee Gruden, Dean College of Graduate Studies

The University of Toledo

May 2019

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

Interrogation of GPCR-G Protein Signaling using Novel Optogenetic Tools

Kanishka D Senarath

Submitted to the Graduate Faculty as partial fulfillment of the requirements for the Doctor of Philosophy Degree in Chemistry

The University of Toledo May 2019

G protein coupled receptors (GPCRs) control a vast majority of signaling responses in the body. Therefore, they are at the center of a large number of diseases, ranging from heart diseases to cancer, and have become the primary target of more than one third of pharmaceutical agents currently available in the market. Due to the limitations of current assays to measure activation of GPCRs and G proteins – such as being time-consuming, indirect, and expensive – an eﬃcient method directly measuring the GPCR-G protein activation was in demand. Therefore, a confocal microscopy imaging-based assay was developed to directly measure ligand-induced GPCR and G protein activity in real time in live cells. The number of active GPCRs governs G protein heterotrimer (Gαβγ) dissociation

(into Gα and Gβγ), and thereby controls the concentration of free Gβγ subunits. The developed Gγ9 assay measures GPCR activation-induced reversible Gβγ9 subunit exchange between the plasma membrane (PM) and internal membranes (IMs). Since G protein activation and dissociation is the immediate next step of GPCR activation, the live cell imaging-based γ9 assay can be used to directly measure ligand-induced GPCR and G protein activity in real time. The Gγ9 assay quantitatively determines the concentration dependency of ligands on GPCR activation, deactivation, and inhibition. Also, the assay

iii demonstrates the high-throughput screening (HTS) adaptability. This assay works eﬀectively for light-sensing GPCRs as well, which enables experimental determination of spatially restricted activation of GPCRs and G proteins not only in single cells but also in subcellular regions of single cells. Overall, the Gγ9 assay provides a robust strategy for quantitative as well as qualitative assessment of GPCR-G protein activation in a single- cell, multicell, and subcellular level, which would be useful in drug discovery eﬀorts.

Gβγ is a major signal transducer and controls multiple cellular processes ranging from cell migration to gene transcription. Despite the significant subtype heterogeneity and diverse cell and tissue specific expressions, Gβγ is often treated as a single signaling entity.

The Gγ subunit bears the only PM anchoring motif in the Gβγ dimer. Our discoveries demonstrate that the differential PM-affinities differentially modulate Gβγ-effector signaling at the PM and subsequent cellular responses (i.e., cell migration) in a Gγ type specific manner. Also, we show that the overall PM-affinity of the Gβγ-pool of a cell type is a strong predictor of its Gβγ signaling activation efficacy at the PM. Overall, our data discloses crucial aspects of Gβγ signaling and cell behavior regulation by Gγ-type specific differential PM-affinities of Gβγ.

Gβγ interacts with the PM through a prenyl group at the carboxy terminus (CT) of

Gγ, which strengthens the PM localization of Gαβγ heterotrimer as well. However, it was not clear how Gβγ possesses this unique and G-type dependent range of PM affinities by using only two types of lipid anchors (either farnesyl or geranylgeranyl) on Gγ. Therefore, we explored the sequence properties of the carboxy terminal region Gγ. Results identified a key existence of hydrophobic residues in the pre-prenylation region of several Gγ types that provided the highest PM-affinity for G. Further, we recognized the crucial

iv contribution of pre-prenylation residues acting as “on-off’’ switches for Gβγ signaling, regardless of the type of prenylation on Gγ. Also, we show that these pre-prenylation regions of Gγ are evolved to be significantly shorter than other prenylated proteins while still maintaining the G protein heterotrimer formation, GPCR-G protein interaction, and G protein signaling ability. Overall, we showed a simple and unique design of pre-prenylation regions of G that allow Gβγ to act as a regulator of GPCR pathways and as a molecular switch for Gβγ signaling.

To optogenetically control GPCR-G protein pathways in vivo, light sensing GPCRs should be able to activate G protein heterotrimers with a comparable efficiency to ligand inducible GPCRs of interest. Over the last two decades, rhodopsin-based chimeric GPCRs have been employed in cultured cells and in vivo to control multiple G protein pathways.

However, these chimeric GPCRs in our studies showed severely defective trafficking to the PM as well as extremely poor signaling activation, both of which could go unnoticed in studies in vitro with purified proteins or in in vivo animal studies. Therefore, we examined the feasibility of chimeric receptor engineering by replacing intracellular loops

(ILs) of human color opsin and found that even a minor change to IL1 and IL2 either drastically reduces or completely abolishes receptor trafficking to the PM. IL3 exhibited more tolerance for mutagenesis, and together with the CT, it allowed switching opsin signaling towards the signaling of the chemokine GPCR of interest. Compared to rhodopsin-based chimeras, these cone opsin-based chimeric GPCRs exhibited several fold higher G protein signaling that can be measured using fluorescence based assays. They also exhibit uninterrupted trafficking. However, in order to reach near similar signaling efficacies in chemical sensing GPCRs, our chimeric receptors require further engineering.

Overall, our work not only resulted in superior light-activatable GPCRs for in vivo optogenetics, but it also serves as a guide for future chimeric GPCR engineering.

Overall, the studies presented establish the significance of the CT of Gγ in determining the PM-affinity of Gβγ and Gβγ mediated signaling activation at the PM. This

Gγ-type dependent PM-affinity is achieved by fine-tuning the residues in the relatively short PM-interacting pre-CaaX regions (compared to other prenylated proteins). For instance, hydrophobic residues (i.e., phe) in the pre-CaaX adjacent to the prenylated-cys in the CaaX motif in Gγ provided the highest PM-affinity for Gβγ. Gγ9 which lacks such hydrophobic character shows the lowest PM-affinity, signified by its ability to elicit a faster

Gβγ translocation rate. This fast and reversible Gβγ9 distribution between the PM and IMs was exploited to develop a universal assay for real-time detection of GPCR-G protein activation, both qualitatively, quantitatively, and in subcellular regions of living cells.

Since GPCR-G protein signaling is universally conserved and Gβγ signaling pathways are major drug targets, mechanisms we identified and described here are likely to have a wide influence in studying and controlling cellular signaling. Also, our newly engineered light activatable chimeric receptors which can signal like chemically sensitive GPCRs with better signaling efficacy than currently available ones will allow to facilitate in vivo optogenetics to study and control numerous GPCR-controlled processes including immune system function, pain, addiction, and many other physiological and behavioral responses with much higher spatial and temporal control.

Acknowledgements

I would like to express my highest gratitude to my advisor, Dr. Ajith Karunarathne, for his support, guidance, and encouragement during the past four years of research.

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

Donald R. Ronning, Dr. Dragan Isailovic, and Dr. Song-Tao Liu. Also, I want to thank Dr.

John L. Payton and Dr. Deborah N. Chadee for their contribution and assistance in computational modeling and Western blotting, respectively, in my projects.

I would like to thank all my lab members for all their support, advice, words of encouragement, and for all the insightful scientific discussions throughout the past few years. I want to thank Elise Harmon for proof reading the abstract of the thesis for me.

Lastly, I would like to thank the Department of Chemistry and Biochemistry, the

University of Toledo for providing me the opportunity to pursue my studies.

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Table of Contents

Abstract ...... iii

Acknowledgements ...... vii

Table of Contents ...... viii

List of Tables ...... xv

List of Figures ...... xvi

List of Abbreviations ...... xxi

List of Symbols ...... xxiv

1 Introduction

1.1 G protein coupled receptors and G proteins ...... 1

1.2 GPCR-G protein activation cycle ...... 2

1.3 GPCR classification and G protein mediated signaling ...... 3

1.4 G protein-plasma membrane interaction ...... 4

1.5 G protein βγ diversity ...... 6

2 Development of a Novel Assay to Detect GPCR-G Protein Activation ...... 7

2.1 Introduction ...... 7

2.2 Results and discussion ...... 10

2.2.1 Characterization of ligand-induced GPCR-G protein activation and

Reversible distribution of Gβγ9 between the PM and IMs ...... 10

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2.2.2 Quantiﬁcation of ligand-concentration-dependent activation of

GPCRs using the Gγ9 assay ...... 14

2.2.3 Dynamic sensing of the environment by the receptor-bound ligand...

...... 16

2.2.4 Assessment of inhibitor-concentration-dependent GPCR inhibition

using the Gγ9 assay ...... 18

2.2.5 Reversible γ9 Distribution as a Universal Assay for Gαi/o- and Gαs-

Coupled GPCR Activity ...... 20

2.2.6 Ligand potency-dependent GPCR activation detection with Gγ9

assay ...... 24

2.2.7 Qualitative Multicellular Screening of Gαi/o- and Gαs- Coupled

GPCR Activation ...... 25

2.2.8 Gγ9 assay is high throughput screening capable ...... 28

2.2.9 Gγ9 acts as a spatiotemporal sensor for subcellular GPCR and G

protein activation ...... 29

2.3 Conclusion ...... 31

2.4 Materials and Methods ...... 32

2.4.1 Constructs, Cell culture, and Transfections ...... 32

2.4.2 Time-lapse imaging ...... 32

2.4.3 Optogenetic control of GPCR signaling and imaging of G protein

dynamics ...... 33

2.4.4 Image Analysis, Data Processing, and Simulation of Laser Power

Distribution ...... 33

3 Gγ Subtype Dependent Plasma Membrane Affinity of Gβγ Controls the Efficacy

of Downstream Signaling Activation and Macrophage Migration ...... 34

3.1 Introduction ...... 34

3.2 Results …. …...... 37

3.2.1 Gγ subtype identity-specific control of PI3Kγ activation by Gβγ .... 37

3.2.2 Optogenetic determination of PM affinities of 12 Gγ subunits using

Tt1/2 of Gγ ...... 41

3.2.3 Gγ-dependent control of chemokine pathway mediated RAW cell

migration ...... 44

3.2.4 Carboxy terminus of Gγ governs rates of Gβγ translocation ...... 48

3.2.5 Control of RAW cell migration by CaaX and pre-CaaX residues in

CT of Gγ ...... 49

3.2.6 Modulation of RAW cell migration potential by Gγ subtype-

dependent activation of PI3Kγ ...... 49

3.2.7 Gγ subtype - dependent control of Gβγ mediated PLCβ activation . 53

3.2.8 Tt1/2 of Gγ is a strong predictor of Gβγ effector activation ability .... 55

3.2.9 Contribution of Gβγ PM-retention on its effector activation ability 59

3.3 Discussion ...... 62

3.4 Conclusion ...... 65

3.5 Materials and Methods ...... 65

3.5.1 Reagents ...... 65

3.5.2 DNA constructs and cell lines ...... 66

3.5.3 Cell culture and Transfections ...... 66

3.5.4 Knockdown of Gγ3 in RAW 264.7 cells ...... 67

3.5.5 Live cell imaging to monitor Gβγ translocation, PIP3 generation, and

optogenetic control of cell migration ...... 68

3.5.6 Cytosolic Ca2+ measurements...... 69

3.5.7 Real-time PCR, transcriptome, and RNAseq data analysis ...... 69

3.5.8 Statistics and reproducibility ...... 70

4 Short Pre-CaaX Regions in Gγ are Uniquely Evolved Switches of Heterotrimeric

G Protein Signaling ...... 71

4.1 Introduction ...... 71

4.2 Results …. …...... 74

4.2.1 Decoding selection criteria of residues in Gγ-CT that tunes Gβγ-PM

interactions ...... 74

4.2.2 Prenylated-cys adjacent phe residues in the CT of Gγ3 are essential

for Gβγ effector activation ...... 76

4.2.3 Location of the phe-duo in Gγ3 pre-CaaX motif is crucial for the

PM-affinity of G ...... 80

4.2.4 The last Gβ-interacting conserved phe in Gγ is not a crucial

determinant of the PM-affinity and activity of Gβγ ...... 82

4.2.5 Positively charged residues in Gγ CT do not significantly contribute

to the PM-affinity and signaling of Gβγ ...... 82

4.2.6 Terminal cys-attached phe-duo is crucial for activation of key

kinases in the chemokine pathway and invasion of cancer cells ...... 84

4.2.6.1 Screening MDA-MB-231 cells for an endogenous

chemokine receptor ...... 85

4.2.6.2 CT of Gγ regulates phosphorylation of kinase; Akt, in

cancer cells ...... 86

4.2.7 Chemokine pathway directed cancer cell invasion is CT of Gγ

dependent ...... 89

4.2.8 G is uniquely evolved to have a short pre-CaaX to support

heterotrimer formation and interaction with GPCRs ...... 91

4.2.9 Short pre-CaaX regions of Gγ are evolutionarily conserved ...... 97

4.3 Discussion …...... 99

4.4 Materials and Methods ...... 103

4.4.1 Reagents ...... 103

4.4.2 DNA constructs and cell lines ...... 103

4.4.3 Cell culture and transfections ...... 104

4.4.4 Live cell imaging to monitor Gβγ translocation and PIP3 generation

… ...... 105

4.4.5 Western blot analysis ...... 105

4.4.6 Cell invasion assay...... 106

4.4.7 Statistical data analysis ...... 106

5 Engineering of Cone Opsin-based Chimeric Receptors with Improved Trafficking

and Signaling ……...... 107

5.1 Introduction ...... 107

5.2 Results …. …...... 110

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5.2.1 Screening rhodopsin-based chimeras and their limitations ...... 110

5.2.2 Blue opsin is an efficient signaling activator ...... 113

5.2.3 IL3 is the most crucial region of the receptor for G protein

interactions ...... 115

5.2.3.1 IL1 and Il2 replacements cause defective receptor

trafficking to the PM ...... 116

5.2.3.2 Size of the loop replacements determine proper folding and

expression of receptor ...... 118

5.2.4 Engineering of β2AR chimeras ...... 121

5.2.4.1 Light-dependent β2AR-specific signaling by chimeric

receptors ...... 121

5.2.4.2 Carboxy terminus of the receptor further strengthens the

β2AR specific signaling of chimeric receptors ...... 127

5.3 Discussion …...... 130

5.4 Materials and Methods ...... 133

5.4.1 Reagents ...... 133

5.4.2 DNA constructs and cell lines ...... 133

5.4.3 Cell culture and transfections ...... 134

5.4.4 Live cell imaging to monitor Gβγ translocation, mini Gs

translocation, cell migration, and PIP3 generation … ...... 135

5.4.5 Statistical data analysis ...... 136

References ...... 137

A Gβ forms Gβγ dimers with overexpressed Gγ types ...... 161

xiii

B Translocation and PIP3 generation properties of Gγ mutants ...... 162

C Sequence alignment between receptors ...... 163

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List of Tables

2.1 Reactions describing the GPCR-G protein activation...... 11

3.1 Translocation properties of WT Gγ types ...... 43

3.2 Translocation properties of Gγ mutants ...... 48

5.1 Excitation and emission wavelengths of common fluorescent proteins ...... 114

B.1 Translocation and PIP3 generation properties of Gγ mutants ...... 162

List of Figures

1 – 1 Steps in GPCR activation ...... 2

1 – 2 G protein mediated signaling ...... 4

1 – 3 Lipid modifications on heterotrimeric G proteins ...... 5

1 – 4 Prenylation process of the Gγ subunits ...... 5

1 – 5 G protein βγ diversity ...... 6

2 – 1 Steps in GPCR activation ...... 9

2 – 2 Continuous heterotrimer shuttling between the PM and IMs ...... 11

2 – 3 Receptor activation-deactivation induced reversible distribution of βγ9 as a

reporter of the GPCR-G protein activity...... 12

2 – 4 Receptor activation-deactivation induced reversible distribution of βγ9 as a

reporter of the GPCR-G protein activity...... 13

2 – 5 Gq coupled GPCR activation induces cell shape change ...... 14

2 – 6 Quantification of concentration dependent endogenous α2-adrenergic receptor

activation in real time in living cells using γ9 assay...... 15

2 – 7 Dynamic sensing of the environment by the α2-adrenergic receptor-bound NE .. 17

2 – 8 Quantification of yohimbine dosage-dependent inhibition of NE-activated α2-AR

...... 18

2 – 9 Receptor co-localization with the FP-γ9 subunit ...... 19

xvi

2 – 10 Gγ9 assay for quantification of ligand carbachol concentration dependent M4-

muscarinic receptor activation ...... 20

2 – 11 Gγ9 assay for quantification of ligand isoproterenol concentration dependent Gs

coupled β1-adrenergic receptor activation ...... 22

2 – 12 GPCR activation universally results in G9 redistribution ...... 23

2 – 13 Differential Gβγ redistribution between plasma and internal membranes with

agonists with different levels of potency for the same receptor ...... 24

2 – 14 Comparison of single and multi-cell GPCR activation by detecting changes in the

cumulative FP-γ9 intensity ...... 25

2 – 15 Qualitative analysis of GPCR activation using low magnification epi-fluorescence

...... 26

2 – 16 Qualitative analysis of GPCR activation under low magnification epi-fluorescence

microscopy ...... 28

2 – 17 High throughput screening adaptability of Gγ9 assay ...... 29

2 – 18 G9 redistribution shows a spatially confined activation of GPCRs and G proteins

in subcellular regions ...... 30

3 – 1 PIP3 generation measurement using the PIP3 sensor ...... 37

3 – 2 Differential PIP3 generation in different cell lines ...... 38

3 – 3 PIP3 generation upon PI3Kγ expression ...... 39

3 – 4 Gγ identity-controlled PIP3 generation...... 40

3 – 5 Gγ-identity driven differential translocation of Gβγ ...... 42

3 – 6 Gγ-identity driven differential PM-affinity of Gβγ ...... 43

3 – 7 Comparison of optical and chemical induced GPCR activation ...... 44

xvii

3 – 8 Subtype specific control of macrophage migration by Gγ ...... 45

3 – 9 Universal nature of HiAf Gγ subunit requirement in cell migration ...... 46

3 – 10 Carboxy terminus of Gγ governs rates of Gβγ translocation ...... 47

3 – 11 Carboxy terminus of Gγ governs the extent of cell migration ...... 49

3 – 12 Gβγ mediated PI3Kγ activation induced PIP3 generation and cell migration ...... 50

3 – 13 Gγ type dependent activation of PI3Kγ during macrophage migration ...... 51

3 – 14 Carboxy terminus of Gγ dependent activation of PI3Kγ activation and

macrophage migration ...... 52

3 – 15 PLCβ activation induced differential Ca2+ response with different Gγs ...... 55

3 – 16 Tt1/2 of Gγ as a predictor of a cell’s ability to control Gβγ effectors ...... 56

3 – 17 Tt1/2 of Gγ as a predictor of a cell’s ability to control Gβγ effectors ...... 58

3 – 18 Predicted model of signal transduction from GPCRs to the cell interior in multi-

Gγ systems ...... 60

3 – 19 Gβγ translocation with Gγ12 mutants ...... 61

4 – 1 Molecular rationale for Gβγ PM-affinity control by pre-prenylation residues of Gγ

...... 75

4 – 2 Pre-CaaX regions of prenylated G proteins ...... 77

4 – 3 Role of prenylated-cys adjacent phe residues in Gγ3 CT in Gβγ effector activation

...... 79

4 – 4 Hydrophobic character in Gγ pre-CaaX motif is crucial for the PM-affinity of G

...... 81

4 – 5 Non-significant influence of positively charged residues in Gγ CT for the PM-

affinity and signaling of Gβγ ...... 83

xviii

4 – 6 Gβγ-driven cell invasion ...... 85

4 – 7 Screening for the endogenous receptors in MDA-MB-231 cells ...... 86

4 – 8 Hydrophobic residues in the pre-CaaX of Gγ control Gβγ-driven PIP3 generation

in MDA-MB-231 cells ...... 87

4 – 9 Hydrophobic residues in the pre-CaaX of Gγ control Gβγ-driven kinases-

activation in MDA-MB-231 cells ...... 88

4 – 10 Gβγ-governed MDA-MB-231 cell-invasion ...... 89

4 – 11 Pre-CaaX residues of Gγ significantly influence Gβγ-governed MDA-MB-231

cell-invasion...... 91

4 – 12 Short pre-CaaX motifs in Gγ compared to Ras family pre-CaaX ...... 92

4 – 13 Activity of Gγ3-KRas mutants ...... 93

4 – 14 Gγ3-KRas mutants are capable of heterotrimer formation...... 95

4 – 15 Inability of Gβγ-Gγ3-KRas mutants to transduce GPCR signaling ...... 96

4 – 16 PM-interacting short pre-CaaX regions of Gγ are evolutionary conserved ...... 98

5 – 1 Limitations of rhodopsin-based chimeras ...... 112

5 – 2 Enhanced blue opsin (BO-E) is as efficient as ligand activatable GPCRs ...... 115

5 – 3 Engineering of blue opsin-CXCR4 chimeras ...... 117

5 – 4 Defective PM trafficking of BO-CXCR4 chimeras ...... 119

5 – 5 Improved BO-CXCR4-IL3(13 AA) chimera induces PIP3 generation and

macrophage migration ...... 120

5 – 6 Loop length dependent receptor activation ...... 123

5 – 7 Gαs activity of BO-β2AR-IL3 chimeras ...... 125

5 – 8 Reduced activation of BO-β2AR-IL3 chimeras upon PtX treatment ...... 126

xix

5 – 9 Gαs signaling reinforcement by the introduction of the CT ...... 128

5 – 10 Reversibility of mini Gs translocation ...... 129

A – 1 Gβ forms Gβγ dimers with overexpressed Gγ types ...... 161

C – 1 Sequence alignment showing the homology between rhodopsin, blue opsin, and

CXCR4...... 163

List of Abbreviations

AC ...... Adenylyl cyclase avg-Tt1/2 ...... Average translocation half time

BO ...... Blue opsin BSA ...... Bovine serum albumin

C5aR ...... Compliment component 5a receptor Ca2+ ...... Calcium cAMP ...... cyclic adenosine monophosphate (cyclic AMP) Cdc42 ...... Cell division control protein 42 CFP ...... Cyan fluorescent protein CT ...... Carboxy terminus CXCR4...... Chemokine receptor 4

D2R ...... D2-Dopamine receptor DAG ...... Diacylglycerol DFBS ...... Dialyzed fetal bovine serum DI ...... Deionized DMEM ...... Dulbecco’s modified eagle medium DMSO ...... Dimethylsulfoxide DOF ...... Depth of field

EC50 ...... Concentration half response |EF|calc ...... Calculated effector response/activity |EF|exp ...... Experimental effector response/activity ER ...... Endoplasmic reticulum

FP ...... Fluorescent protein FRAP-PA ...... Fluorescence recovery after photobleaching and photoexcitation FRET...... Fluorescence/Foster recovery energy transfer

GAP ...... GTPase activating/accelerating proteins GDI ...... Guanine dissociation inhibitor GDP ...... Guanosine diphosphate GEF ...... Guanine nucleotide exchange factor GFP ...... Green fluorescent protein

xxi

GIRK...... G protein-gated inwardly rectifying potassium channels GPCR ...... G protein coupled receptor GPI ...... Glycosylphosphatidylinositol GRK ...... GPCR kinase GTP ...... Guanine triphosphate Gα ...... G protein alpha subunit Gβγ ...... G protein beta gamma Gβγ*...... Transient active conformation of Gβγ

HBSS ...... Hank’s balanced salt solution HiAf ...... High plasma membrane affinity HRP ...... Horseradish peroxidase

IC50...... Half maximum inhibitory concentration Icmt ...... Isoprenyl-cysteine carboxyl methyl transferase IMs ...... Internal membranes IP3 ...... Inositol triphosphate Iso ...... Isoproterenol

K+ ...... Potassium

LoAf ...... Low plasma membrane affinity mCh ...... mCherry MEM ...... Minimum essential medium MoAf...... Moderate plasma membrane affinity MoR1R...... Mu opioid receptor

NE ...... Norepinephrine NT ...... Amino terminus

OA ...... Optical activation

PDE ...... Phosphodiesterase PFS...... Perfect focus system PI3K ...... Phosphatidylinositol-4,5-bisphosphate 3-kinase PIP2 ...... Phosphatidylinositol 4, 5-bisphosphate PIP3 ...... Phosphatidylinositol 3, 4, 5-trisphosphate PKA ...... Protein kinase A PLC ...... Phospholipase C PM ...... Plasma membrane PS ...... Penicillin-Streptomycin PtX ...... Pertussis toxin

Rac1 ...... Ras-related C3 botulinum toxin substrate 1 RCE1...... Ras converting CaaX endopeptidase

xxii

RGS ...... Regulators of G protein signaling ROI ...... Region of interest RPMI...... Roswell Park Memorial Institute

S-1-P ...... Spingosine-1-phosphate SD ...... Standard deviation SDS ...... Sodium dodecyl sulfate SEM ...... Standard error of the mean STR ...... Short tandem repeat

TIRF ...... Total internal reflection TM ...... Transmembrane Tt1/2 ...... Translocation half time

UV ...... Ultraviolet

LE ...... Leading-edge velocity TE ...... Trailing-edge velocity

WB ...... Western blot WT ...... Wild type

YFP ...... Yellow fluorescent protein

α2-AR ...... Alpha 2 adrenergic receptor β2-AR ...... Beta 2 adrenergic receptor

xxiii

List of Symbols

α ...... Alpha β ...... Beta γ ...... Gamma

0C ...... Degrees Celsius n ...... Number of cells or individual experiments

xxiv

Chapter 1

Introduction

This chapter contains materials which were originally published in the International Review of Cell and Molecular Biology. Kanishka Senarath, Dinesh Kankanamge, Saroopa Samaradivakara, Kasun Ratnayake, Mithila Tennakoon, Ajith Karunarathne. Regulation of G protein βγ signaling. Reprinted with permission from the Journal Int. Rev. Cell. Mol. Biol. 2018; Vol 339:133-191. Copyright © 2018 Elsevier.

1.1. G protein coupled receptors and G proteins:

GTP binding protein (G protein) coupled receptors (GPCRs) represent the largest family of cell surface receptors found in all eukaryotic genomes which control the majority of cellular signaling in the body. GPCRs sense a wide range of external stimuli including hormones, neurotransmitters, small peptides, chemical ligands, lipids as well as light (1,

2). All GPCRs share a common structure with seven transmembrane (TM) domains, an extracellular amino terminus (NT), and an intracellular carboxyl terminus (CT).

Transmembrane regions are connected through three extracellular and three intracellular/cytosolic loops. Light sensitive GPCRs in the vision system; rhodopsin and opsins, also have the same seven TM structure, and covalently bind to their ligand/inverse agonist; 11-cis retinal, by forming a Schiff base with a Lys residue (Lys296) in the seventh

TM helix of the receptor (3, 4). Upon photon absorption, receptor bound 11-cis-retinal gets

1 isomerized to all-trans-retinal and activates the receptor by triggering a receptor conformational change (5, 6).

1.2. GPCR-G protein activation cycle:

GPCRs interact with heterotrimeric G proteins, consisting of three subunits; Gα, and Gβγ (1, 7). In the G protein inactive state, Gα is bound to a guanosine diphosphate

(GDP) nucleotide and is in association with Gβγ which prevents the spontaneous

Fig. 1-1. Steps in GPCR activation. In the GPCR inactive basal sate, G protein exists as a heterotrimer with GγGDP in association with Gβγ (GαGDPβγ). This association assists in Gα-GPCR coupling and acts as a guanine nucleotide dissociation inhibitor (GDI) to prevent GDP release from Gα. Upon GPCR activation by the ligand binding, GPCR undergoes conformational change, and exchange bound GDP to a GTP on Gα subunit by acting as a GEF. GTP binding leads to heterotrimer dissociation to GαGTP and Gβγ and both active subunits independently interact with downstream effectors to initiate downstream signaling. Intrinsic GTPase activity of GαGTP hydrolyzes GTP to GDP resulting the system to reach the inactive state, which is enhanced by RGS proteins acting like GAPs for Gα subunits in vitro. dissociation of GDP from Gα by acting as a guanine dissociation inhibitor (GDI) (8, 9).

Upon GPCR activation, the receptor undergoes conformational changes, which acts as a guanine nucleotide exchange factor (GEF) and results in a GDP to guanine triphosphate

(GTP) nucleotide exchange in Gα making the G protein active. This guanosine nucleotide exchange, which is driven by a conformational change in three flexible switch regions

2 surrounding the nucleotide-binding pocket in Gα (8, 10), results in a Gα conformational change, leading to active G protein heterotrimer dissociation in to GTP bound active Gα and Gβγ (11-14) (Fig. 1-1). Activated G proteins; both Gα (GTP bound) and free Gβγ, interact with effectors on or near the plasma membrane (PM) and regulate the majority of eukaryotic signaling to govern a cohort of biological processes including tissue homeostasis, development, and immunity (15-17).

With the hydrolysis of GTP into GDP on the active Gα subunit by the inherent guanosine triphosphatase (GTPase) activity of Gα, the GDP bound inactive Gα re-associate with free Gβγ, reforming the Gα(GDP)βγ heterotrimers. This GTP hydrolysis is controlled by a family of regulators of G protein signaling (RGS) proteins by acting as GTPase accelerating proteins (GAPs) (Fig. 1-1) (8).

1.3. GPCR classification and G protein mediated signaling:

GPCRs can be classified based on the type of the Gα subunit type in the G protein heterotrimers they interact with (Gαs, Gαi/o, Gαq and Gα12/13 coupled). Stimulatory

(Gαs) pathway activates adenylyl cyclase (AC), which catalyzes the formation of the second messenger cyclic AMP (cAMP). Generated cAMP then activates cAMP stimulated protein kinase A (PKA), which phosphorylates target proteins in the cell. These interactions are attenuated by phosphodiesterases (PDE), which break down cAMP and dampens the signal (18). Activation of inhibitory G protein (Gαi/o) pathway inhibits AC and decreases cAMP, inactivates PKA, and induces opposite effects elicited by the Gαs

(18, 19). Gαq pathway stimulates phospholipase C (PLC), which hydrolyzes phosphatidylinositol 4, 5-bisphosphate (PIP2), a PM phospholipid, and generates two

3 second messengers; inositol trisphosphate (IP3) and diacylglycerol (DAG). IP3 diffuses through the cytosol, binds IP3 receptors in the endoplasmic reticulum (ER), and releases stored Ca2+ increasing the Ca2+ level in the cell. DAG, which stays on the PM, stimulates

Fig. 1-2. G protein mediated signaling. protein kinase C (PKC) activation, thereby phosphorylates target proteins (18).

Once dissociated from the heterotrimer, active free Gβγ also initiates multiple signaling pathways by activating an array of effectors such as Ca2+ and K+ channels (20), certain isoforms of AC (AC2, AC4, and AC7 are activated by Gβγ, while AC1, AC5, and

AC6 are inhibited by Gβγ) (21, 22), and phosphoinositide-specific PLCβ (23). Also, free

Gβγ can detach from the PM and translocate to internal membranes (IMs) (Golgi, endoplasmic reticulum) as well (24, 25).

1.4. G protein-plasma membrane interaction:

Heterotrimeric G proteins are interacting with the PM through lipid modifications on Gα and Gγ. Gα subunits are post-translationally modified at the NT with either 14-C myristoyl and/or 16-C palmitoyl group attachment (26), while Gγ subtypes are modified at the CT with a prenyl group attachment, either 15-C farnesyl or 20-C geranylgeranyl (8, 27, 28) (Fig. 1-3). The prenyl group is attached at the cys residue of the C terminal CaaX motif in Gγ, where ‘aa’ Fig. 1-3. Lipid modifications on heterotrimeric G represents any two aliphatic amino acids and proteins, which assist their plasma membrane ‘X’ is any amino acid. Depending on the last interaction. residue of the CaaX motif (X), either one of two prenyl groups are attached to the cys residue in CaaX through a thioether bond by a specific prenyl transferase; either farnesyl transferase or geranylgeranyl transferase (28). After the prenylation, the last three residues

Fig. 1-4. Prenylation process of the Gγ subunits.

(aaX) are cleaved off by an endoprotease; Ras converting CaaX endopeptidase (RCE1), and the new CT prenylcysteine is methylated by a methyltransferase; isoprenyl-cysteine carboxyl methyl transferase (Icmt) (29) (Fig. 1-4).

All twelve Gγ types possess either one of the two prenyl groups (either farnesyl or

5 geranylgeranyl) at the cys motif in their CaaX motif. Except Gγ1, Gγ9, and Gγ11which are farnesylated, all other Gγs are known to be geranylgeranylated (30, 31). This lipid modification assists in membrane anchorage of Gγ subunits, promoting Gβγ interaction with Gα subunit, thereby forming the heterotrimer.

1.5. G protein βγ diversity

In human and mouse genome, five Gβ (32, 33) and twelve Gγ (34, 35) isoforms have been identified. From the five Gβ isoforms, only Gβ1–4 subtypes interact with a Gγ subunit to make a functional Gβγ dimer, and share greater than 80% amino acid sequence similarity

(Fig. 1-5), while Gβ5 subunit exhibits only less than 50% identity with the other Gβ subtypes (36). In contrast, the 12 Gγ subtypes show a significant amino acid sequence variation, ranging from 20% to 80% (37) (Fig. 1-5). These 12 Gγ types dimerize with any

Fig. 1-5. G protein βγ diversity. Gβ1-4 shows greater than ~80% sequence similarity compared to the huge diversity between Gγ isoforms ranging from ~20-80%.

Gβ type which can dimerize (Gβ1-

Gβ4) resulting 48 possible Gβγ

combinations. The sequence

diversity of Gγ has resulted

differential Gβγ translocation rates

depending on the Gγ type the Gβγ

possess (38, 39). Similarly, Gβγ exhibit Gγ subtype-specific interactions with Gβγ effectors leading to the occurrence of Gγ subtype-specific G protein signaling activation as well (38).

Chapter 2

Development of a Novel Assay to Detect GPCR-G

Protein Activation

This chapter contains materials which were originally published in the Journal of Analytical Chemistry. Kanishka Senarath, Kasun Ratnayake, Praneeth Siripurapu, John L. Payton, and Ajith Karunarathne. Reversible G Protein βγ9 Distribution-Based Assay Reveals Molecular Underpinnings in Subcellular, Single-Cell, and Multicellular GPCR and G Protein Activity. Reprinted with permission from the Journal of Analytical Chemistry 2016, 88 (23), 11450-11459. Copyright © 2016 American Chemical Society.

2.1. INTRODUCTION:

G protein coupled receptors (GPCRs) and their interacting partners, heterotrimeric

G proteins, are universal controllers of cellular signaling. Other than controlling physiological conditions such as immune system function, neuronal development, vision, heart muscle contraction, etc., GPCRs and G proteins are implicated in a majority of pathological conditions, ranging from heart diseases, immune system malfunctions/failures to cancer as well. Although ∼30% of all drugs on the market act on GPCRs, the molecular mechanisms of their activation and signal transduction from the extracellular environment to the cellular interior are just coming to light (40).

Measuring a ligand’s ability to induce conformational changes in a GPCR-inducing heterotrimeric G protein activation is important in pharmacology, and this action is required to identify potent and selective modulators of signaling. In early GPCR assays, radioisotope-labeled ligand binding to isolated cell membranes has been used, and in addition to the risks of radiation exposure, the lack of dynamic signaling information has been an impediment (41, 42). As a direct measure of GPCR activation and heterotrimer dissociation, the GTPγS binding assay was developed. Although it measures G protein activation in multiple types of GPCRs and has been developed into a high-throughput assay, extensive protein puriﬁcation requirements and radioactive material usage have diminished its wide applicability (43, 44). Fluorescence- and bioluminescence-based methods that have been developed later possess the ability to acquire information on ligand-GPCR interactions and second messenger activities in living cells in a high- throughput fashion. Among these, the fluorescence resonance energy transfer (FRET)- based ePAC sensor has been frequently used to measure the activities of both Gαs and

Gαi/o (activation and inhibition of cAMP production) (45, 46). Furthermore, several assays have been developed to measure FRET between G proteins - G proteins and G proteins -

GPCRs to measure GPCR activation and heterotrimer dissociation (14, 47, 48). However, these assays have certain limitations, including inadequate sensitivity, the requirement of multiple genetically encoded protein expression, and the design of speciﬁc assay components for every ligand or GPCR. They are also susceptible to interference from other signaling entities, especially when detecting downstream signaling molecules (43, 49, 50).

In recent years, there has been a substantial interest in developing label-free live cell assays, in which GPCR activity-induced morphological, as well as electrical changes

8 of cells have been measured (51). However, label-free assays are hampered by the lack of speciﬁcity, matrix interference, and their limited sensitivity. In contrast, the γ9 assay described below has overcome these limitations through its higher sensitivity, selectivity, and high-throughput screening adaptability gained by measuring the common signaling element for all GPCR pathways: the heterotrimer dissociation (40).

Gα subunits and Gγ subunits are post-translationally modiﬁed with a diverse group of lipids at the N-terminus and C- terminus, allowing them to interact with and remain bound to the PM lipid bilayer when they are in the heterotrimeric form (inactive) (26,

40). Initially, free Gβγ has been thought to be restricted to the PM, Fig. 2-1. Steps in GPCR activation. although it transiently interacts with the PM through C-terminal prenyl moiety of the Gγ subunit (27). Depending on the CaaX motif in the CT of Gγ, the lipid modification (prenyl moiety) can either be farnesyl or geranylgeranyl (28). However, this prenylation is not sufficient for its membrane targeting and, therefore, support from the C-terminal polybasic peptide region is required (52). Mammalian cells express 12 Gγ subunits and they possess different C-terminal polybasic regions, allowing Gβγ subunits to have different PM affinities (53, 54).

Interestingly, all of the Gγ subunits are capable of reversibly moving between the

PM and IMs upon activation of Gαi/o, Gαs, and Gαq coupled receptors with distinctly diﬀerent rates (24, 53), while activated Gα subunits remain bound to the PM (24, 55).

Among the 12 Gγ subunits, Gγ3 shows the slowest translocation rate (translocation half- time: t1/2 ≳ 250 s), while Gβγ subunits with Gγ9 possess the fastest shuttling between the

PM and IMs with the forward translocation of t1/2 ≈ 10 s (24). Therefore, the ability of free

Gβγ9 subunits to reversibly distribute between the PM and IMs was employed and developed as an assay (γ9 assay) to detect ligand concentration-dependent GPCR-G protein activation and deactivation in living cells (40).

2.2. RESULTS AND DISCUSSION:

2.2.1. Characterization of ligand-induced GPCR-G protein activation and Reversible distribution of Gβγ9 between the PM and IMs:

Previous work has shown that all Gγ subunits are capable of translocating as Gβγ dimers from the PM to the IMs with distinctly diﬀerent rates upon activation of GPCRs

(24). A group of reactions describing this process is given in Table 2.1 which shows:

(a) ligand (L) binding to the receptor (R) and RL complex formation,

(b) RL-Gα(GDP)βγ ternary complex formation,

heterotrimer dissociation resulting in generation of Gα(GTP) and free Gβγ at

the inner leaﬂet of the PM,

(d) free Gβγ redistribution between the PM and IMs,

(e) hydrolysis of GTP in α due to its intrinsic GTPase activity accelerated by GAPs,

(f) heterotrimer regeneration due to the higher aﬃnity of Gα(GDP) for Gβγ. 10

Table 2-1. Reactions describing GPCR-G protein activation (a)

(b, c)

(d)

(e)

(f)

Basal Bleach Recovery

erlay

CFP

αo

mCherry

γ9

- YFP

Fig. 2-2: Continuous heterotrimer shuttling between the PM and IMs: Images of a HeLa cell expressing αo-CFP, mCherry- β1 and YFP- γ9 before, after photobleaching of fluorescence in IMs and after fluorescence recovery after photobleaching (FRAP). Images and the plot show the recovery of fluorescence, due to shuttling of G protein subunits from the PM to IMs. All three subunit types show a similar recovery (yellow arrows) indicating that they shuttle as the heterotrimer (scale bar: 5µm).

IM photobleaching and ﬂuorescence recovery data showed that G proteins

continually shuttle between the PM and IMs, even as the Gα(GDP)βγ heterotrimer (Fig. 2-

2). This transient interacting nature of G proteins prevents the detection of their subcellular

locations using immuno-ﬂuorescence.

HeLa cells expressing GFP-Gγ9 show a distinct PM distribution and, after

activation of endogenous CXCR4 receptors, Gγ9 subunits translocated to IMs (Golgi and

endoplasmic reticulum-ER), until equilibrium was reached (Fig. 2-3). The addition of the

CXCR4 antagonist, AMD3100, completely reversed this process, suggesting that the free

A. B. AMD3100 SDF

Before SDF

After SDF

Fig. 2-3: Receptor activation-deactivation induced reversible distribution of βγ9 as a reporter of the GPCR-G protein activity. A. 4 dimensional (4D) confocal live cell imaging of HeLa cells expressing YFP-γ9 before and after activation of endogenous CXCR4 receptors with 100 ng/ mL SDF-1α. Note the robust accumulation of YFP fluorescence in IMs (yellow arrow). B. Single confocal plane images of HeLa cells (on the plot) showing CXCR4 activation induced translocation of YFP-γ9 from the PM to IMs while the PM distributed CFP-αo remained unchanged. The Gβγ accumulated in IMs was completely reversed to the PM when CXCR4 was inhibited with its antagonist, AMD3100 (10 µM). Curves show the intensity of YFP-9 on the PM (black), in IMs (red) and the IM/PM intensity ratio (blue).

Gβγ subunit continuously shuttles between the PM and IMs. (Fig. 2-3B). This allows free

Gβγ to continuously sense the PM and reverse upon Gα(GDP) formation. The plot shows

GFP dynamics on the PM (black), in IMs (red), and the IM/PM ratio (blue) before and after

the ligand (SDF-1α) addition and after the antagonist addition (Fig. 2-3B). Since GPCR

activation results in GFP-Gγ9 ﬂuorescence reduction on the PM and an increase in IMs,

either the ﬂuorescence in IMs (FIM) or the ratio (FIM/PM) can be considered as the extent of

heterotrimer dissociation as well as GPCR activation in living cells (Fig. 2-3B).

Other than ligand inducible GPCRs, G9 senses the activation of light activatable

GPCRs; opsins, as well. When a cell expressing light sensing GPCR; blue-opsin and

mCherry-9 was pre-incubated with 11-cis-retinal and optically activated (OA) blue opsin

at 20 Hz by exposing cells globally to 445 nm laser light (500 nW/μm2), mCherry-9

A. B.

Before OA Recovery

Fig. 2-4: Receptor activation-deactivation induced reversible distribution of βγ9 as a reporter of the GPCR-G protein activity. A. A HeLa cell expressing blue- opsin-GFP and mCherry-9 was pre-incubated with 11-cis-retinal and globally exposed to 445 nm laser to activate opsin. Observed translocation of mCherry-9 from PM to IMs with opsin activation was reversed after termination of the activation, scale bar = 5 µm. B. Plot shows the fluorescence intensity of mCherry- 9 in IMs upon OA of blue-opsins. IM fluorescence is decreased over time after termination of OA at t = 100 s. (n = 7 cells) (Mean  Standard error-(SEM)).

translocation from the PM to IMs was observed, which reached a steady state with t1/2 < 10

s which was reversed after termination of OA (Fig. 2-4).

2.2.2. Quantiﬁcation of ligand-concentration-dependent activation of GPCRs using the

Gγ9 assay:

Both Gαi/o and Gαs coupled receptors can induce a profound Gβγ redistribution

upon activation, while Gαq-coupled receptors produce only a marginally detectable signal

(Fig. 2.5). In addition, Gαq-coupled receptor activation induces a signiﬁcant change in cell

morphology, introducing artifacts in live cell assays (Fig. 2.5.A).

A. B. GFP-γ9 GRPR-mCh Overlay

Activated Basal Activated

Fig. 2-5: Gq coupled GPCR activation induces cell shape change. A. HeLa cells transfected with GFP-γ9 and Gq coupled gastrin releasing peptide receptor- mCherry shows a significant cell shape change upon activation with 1 µM Bombesin perturbing the detection of γ9 translocation. The white outline on the overlay image shows the membrane periphery before receptor activation illustrating the extent of cell shape change. The plot shows the fluorescence intensity of IMs over time.

To examine the ligand-concentration-dependent activation of endogenous

α2 adrenergic receptors (α2-ARs) (56), HeLa cells expressing GFP-Gγ9 were activated with

norepinephrine (NE) (Fig. 2-6). As the concentrations of these ligands increase, a gradual

increase in the ﬂuorescence in IMs and a decrease in the PM, compared to the basal level, was

observed (Fig. 2-6A). The plots show FIM/PM is sensitive to a broad range of concentrations,

ranging from the nanomolar level to the micromolar level of NE concentrations. 14

Despite the transient expression of Gγ9 and subsequent heterogeneous GFP expression among cells, once normalized to a basal level ﬂuorescence (before NE addition), all of the cells showed a uniform dose-response relationship. Regardless of having a slightly diﬀerent expression of

GFP-Gγ9 in the two cells in Fig. 2-6A, a clear GFP intensity change on the PM and in IMs can be observed in both cells. The FIM/PM data, at steady state with no further accumulation of Gβγ in IMs, were used to construct the dose-response curve (Fig. 2-6B). Fitting the experimental

A. B. Basal 30 nM 200 nM 800 nM

7 μM 15 μM 25 μM 100 μM

Fig 2.6: Quantification of concentration dependent endogenous α2-adrenergic receptor activation in real time in living cells using γ9 assay. A. A HeLa cell transiently expressing GFP-γ9 was exposed sequentially to 0.5 nM to 100 μM NE while capturing time-lapse images at every one second interval. After each concentration of NE exposure, cells were given 200 seconds to reach the equilibrium before the addition of the next concentration. Images show the GFP-γ9 distribution in two cells. Note the gradual reduction of GFP intensity on the PM and simultaneous accumulation in IMs with the step-up increase of NE. B. Plot shows the change in fluorescent intensity in IMs and the PM calculated as their ratio (FIM/PM) which is normalized to the baseline (n=20 cells) (Mean  SEM). Red line shows the DoseResp 푦 = 퐴1 + 퐴2−퐴1 function fitted to the experimental data to calculate the EC50 1+10((퐿표푔 푥0−푥ሻ∗푝ሻሻ value (A1 = initial Y value (initial response), A2 = final Y value (final response), x = logarithm of agonist concentration/dose, x0 = center of the curve (EC50/concentration for half response) and p = slope factor/Hill coefficient).

data with the DoseResp function (in OriginLab) resulted in a submaximal concentration of 2.13

± 0.88 μM, which is equivalent to the EC50 value of NE in this single cell assay. In vitro ligand

15 response studies with mesenteric vascular smooth muscle contraction in rats have shown that these tissues respond to NE with an EC50 value of 400 nM (57). Given that the Gγ9 assay is performed in cultured single cells, there are several reasons that can cause this discrepancy in the EC50 values, such as (i) the limits of detection of the imaging sensor for ﬂuorescently tagged free Gγ9, (ii) receptor concentration on the PM, and (iii) molecular considerations inherent to

Gβγ generation, such as its limited lifetime governed by the lifetime of Gα(GTP), and iv) the diffusion time of NE in the media. After each experiment, the NE-containing media was replaced with fresh imaging buffer and a complete reversal of Gγ9 back to the PM was observed, indicating that no significant receptor desensitization occurred during the experiment.

2.2.3. Dynamic sensing of the environment by the receptor-bound ligand:

Equation (a) in Table 2-1 shows the equilibrium in RL complex formation, indicating the reversible exchange of the ligand in the bulk media with the binding pocket of the receptor. This environment sensing mechanism was examined using the γ9 assay to determine if the gradual reduction of ligand concentration allows the reaction to reach a steady state with a new equilibrium. HeLa cells expressing GFP-Gγ9 were exposed to a saturating concentration of the ligand and allowed 200 s to reach steady state before stepping down the concentration. Images of the subcellular regions show the return of IM- bound GFP-γ9 to the PM, once the [NE] reaches a subnanomolar value (∼0.5 nM) (Fig. 2-

7). The kymograph of a one-pixel-wide horizontal cross section (orange line) of the cell shows the change in GFP intensity during the gradient dilution of NE (Fig. 2-7A. bottom).

The scale below the image shows the concentration of NE at diﬀerent time points. The plot shows the corresponding changes in FIM/PM during the gradual dilution of 10 μM NE (Fig. 16

2-7B). The gradual dilution of NE resulted in a lower EC50 value (859 ± 79 nM), compared

to that of the gradient addition, suggesting a bistable receptor activation-deactivation

behavior. A similar observation has been reported in previous studies, which explained

using the ability of ligands to move among receptors, as a result of the dilution (58, 59).

A. B. IM PM

IM PM

Fig 2-7: Dynamic sensing of the environment by the α2-adrenergic receptor-bound NE. A. GFP-γ9 expressing HeLa cells were exposed to 10 μM NE and GFP-γ9 distribution was captured using a confocal time-lapse experiment. Once the maximum FIM/PM was observed, concentration of NE in the cell culture medium was systematically reduced by doubly diluting with the fresh imaging medium. Images (top) show a subcellular region of a cell from this experiment exposed to 10 μM NE (left) and after diluting down to 8 nM. The green line shows the cross section used to construct the orthogonal slice view below. The orthogonal slice view of the cross section shows the minimum PM and maximum IM fluorescence at 10 µM NE. Note the return of GFP-γ9 in IMs to the PM as the [NE] reduces. B. The plot shows the baseline normalized FIM/PM ratio (n=5 cells). This shows the intensity changes of this cross section over time. Concentrations of NE (in μM) at specific time points are shown in the scale below the images. Note that, upon dilution, IM bound γ9 returns to the PM. Red line on the plot shows the DoseResp fit.

2.2.4. Assessment of inhibitor-concentration-dependent GPCR inhibition using the Gγ9 assay:

The sensitivity of the γ9 assay to detect the gradual inhibition of activated α2-AR was tested in HeLa cells expressing GFP-Gγ9 treated with 10 μM NE. Cells with fully activated α2-AR were exposed to 0.5 nM to 20 μM yohimbine at intervals of 200 s (Fig. 2-

8). In order to visualize the cumulative ﬂuorescence changes on the PM and in IMs (orange and blue regions of interest (ROIs), respectively), time stacks of the cropped ROIs were

A. B. Before NE After NE Yohimbine

Fig 2-8: Quantification of yohimbine dosage-dependent inhibition of NE-activated α2- AR using the γ9 assay. A. Images (top) show the GFP-γ9 distribution before and after 10 µM NE and after 20 µM yohimbine. Orange and blue ROIs in the left image were used to generate a 3D time stack to capture the multi-pixel thick PM and IMs through spatial rendering. The image strips show the time stacks of the PM and IMs indicating the loss and the gain of fluorescence respectively as the yohimbine concentration increases. B. The plot shows the experimental dose response curve (black) and the DoseResp fit (red) from origin (n=20 cells, error bars: mean  SEM). created (Fig. 2-8A). The time stack image of IMs shows a gradual reduction, while that of the PM shows a gradual increase in GFP as yohimbine concentration increases. Subcellular confocal images show the distribution of GFP ﬂuorescence before and after NE and after the addition of 20 μM yohimbine. Average dose response curve from multiple cells shows

18 an eﬀective inhibitory action of yohimbine, ranging from 1 nM to 1 μM with the IC50 value of 59 ± 4 nM (Fig. 2-8B). In the presence of an agonist, using an in vitro assay, it was reported that yohimbine is eﬀective in the concentration range from 10 pM to 100 nM, with an IC50 value of ∼5 nM (60). While the higher IC50 value of yohimbine observed in single cells can be explained by the same reasons discussed above, the sigmoidal dose-response curve has the same 100-fold range as reported in the in vitro assay (60). Therefore, the higher range for signaling inhibition appears to be physiological and not due to the lack of sensitivity in the γ9 assay. A. B. BO-GFP mCh-γ9 Overlay β2AR-CFP GFP-γ9 Overlay

2D view

3D view

C. M4 - MTQ GFP - γ9 Overlay D. β1AR-CFP GFP-γ9 Overlay

2D view

3D view

Fig. 2-9: Receptor co-localization with the FP-γ9 subunit. HeLa cells transfected with fluorescently tagged receptors and γ9 subunits were imaged under 60X objective to capture their co-localization to examine if FP tagged γ9 subunits universally interact with GPCRs. Two- and three-dimensional images generated using confocal optical sectioning show the distribution of fluorescently tagged GPCRs and γ subunits. A. blue opsin-GFP and mCherry-γ9. B. α2-adrenergic receptor-CFP and GFP-γ9. C. M4-muscarinic receptor-mTurquoise and GFP- γ9. D. β1-adrenergic receptor-CFP and GFP-γ9 (Scale = 5 µm).

2.2.5. Reversible γ9 Distribution as a Universal Assay for Gαi/o- and Gαs-Coupled

GPCR Activity:

Even though it is controversial if the GPCRs and G proteins are pre-coupled in the receptor inactive state or G proteins interact with GPCRs upon ligand binding (14, 61-63),

A. Basal 50nm 250nm 500 nM

5 μM 20 μM 50 μM 100 μM

B. Before After 1000X dilution

IM PM

5 2.5 1.25 .6 .3 .15 .08 .04 .02 μM

Fig 2-10: Gγ9 assay for quantification of ligand carbachol concentration dependent M4-muscarinic receptor activation in living cells. A. Images show GFP-γ9 distribution during carbachol dosage dependent activation of M4-muscarinic receptors in HeLa cells. Plot shows the corresponding dose-response (black) and DoseResp fit (red) curves (n=10 cells, error bars: SEM). B. Gradient dilution of 5 µM carbachol results in dosage dependent GFP-γ9 reversal from IMs to the PM. Images show the GFP-γ9 distribution before and after 5 µM carbachol and after 1000-fold dilution of carbachol. The orthogonal slice view below generated using green ROI clearly shows the re-appearance of the PM as the carbachol concentration decreases. Plot shows the corresponding dose- response (black) and DoseResp fit (red) curves (n=8 cells, error bars: SEM).

20 with the overexpression of different types of GPCRs and Gγ subunits, we could observe that they form overlapping microdomains on the PM, indicating that they either are pre- coupled or exist in close proximity, which is common for a wide variety of Gαi/o-, Gαs-, or Gαq-coupled receptors (Fig. 2-9). In order to examine if γ9 assay can be used to measure the activation and deactivation of Gαi/o- or Gαs-coupled GPCRs in general, a series of experiments were conducted with heterologous expression of M4-muscarinic receptors and

β1-adrenergic receptors.

HeLa cells expressing M4-muscarinic-mTurquoise and GFP-Gγ9 showed a dose- dependent redistribution of Gγ9 between the PM and IMs upon gradient-addition (Fig. 2-

10A), as well as dilution of carbachol (Fig. 2-10B). Similar to a typical dose response curve, plots show that the Gγ9 response is sensitive to carbachol concentrations ranging from 10 nM to 10 μM with a non-linear relationship. The plots were ﬁtted with the

DoseResp equation with a nonlinear Hill slope, as described in Fig. 2-6. During the activation and deactivation of M4-muscarinic receptors, the dose-response behavior observed was similar to that of NE (Fig. 2-10). The curve ﬁtting resulted in an EC50 value of 378 ± 65 nM for carbachol gradient additions, which was shifted to the left with an EC50 value of 71 ± 12 nM during dilution experiments. An in vitro study conducted on tissues extracted from guniea-pig ileum has shown an EC50 value of 177 nM for carbachol, indicating the comparable sensitivity of the γ9 assay to that of the in vitro assay (64).

Similarly, the activation of β1-AR in HeLa cells expressing β1-AR-CFP and GFP-

Gγ9 with Iso concentrations from 0.1 nM to 16 μM produced a dose-dependent FIM/PM response with an EC50 value of 158 ± 60 nM (Fig. 2-11A). A chemiluminescence assay to examine the ability of Iso to induce the luciferase expression conducted in cardiomyocytes 21 has resulted in an EC50 value of ∼400 nM, indicating that, for certain ligand-receptor

Basal 10 nM 50 nM 250 nM

500 nM 1 μM 4 μM 16 μM B. IM

PM PM IM PM

16 2 .5 .25 .1 .05 .01 .001 .0005 μM

Fig 2-11: Gγ9 assay for quantification of ligand isoproterenol concentration dependent Gs coupled β1-adrenergic receptor activation in living cells. A. GFP time-lapse images of a HeLa cell expressing GFP-γ9 and β1AR-CFP were captured at 1 Hz. After each dose of agonist addition, cells were given 200 s to reach the equilibrium before the addition of the next concentration. Images show the GFP-γ9 distribution in a single cell. Note the gradual reduction of GFP intensity on the PM and simultaneous accumulation in IMs as the ligand concentration increases. Plot shows the dose-response curve (n = 5 cells, error bars: SEM) D. Examination of dosage dependent reversal of GFP-γ9 to the PM. HeLa cells were exposed to 16 μM Iso and GFP-γ9 distribution was monitored while doubly diluting the ligand gradually. Images show the distribution before and after the addition of 16 µM Iso and diluting it. The generated orthogonal slice view (with respect to the cross section shown in green) shows that, IM bound GFP-γ9 return to the PM upon dilution. Plot shows the corresponding dose-response curve (black) and the DoseResp fit (red) (n = 18 cells, error bars: SEM). systems, Gγ9 assay can outperform the conventional methods (65). Serial dilution of Iso from 10 μM to 0.5 nM and examination of GFP-γ9 return from IMs to the PM resulted in a slightly left-shifted dose-response curve with an EC50 value of 120 ± 5 nM (Fig. 2-11B). 22

Every Gαi/o- or Gαs-coupled receptor tested so far with the Gγ9 assay, including the μ and κ opioid, c5a, and D2 dopamine showed a robust Gγ9 translocation upon receptor

A. B. Kappa opioid receptor Mu opioid receptor (MOR1R) (KOR)

Before After Before After

C. D. Compliment component 5a D2-Dopamine receptor (D2R) receptor (c5aR)

Before After Before After

Fig 2-12: GPCR activation universally results in G9 redistribution. HeLa cells transiently transfected with GFP-Gγ9 show translocation upon activation of, A. KOR with 1 µM U50488, B. MOR1 4 µM DAMGO, C. D2R with 10 µM Dopamine. D. endogenous C5aR in RAW 264.7 cells with 10 µM C5a. Yellow arrows on images point the GFP accumulation in IMs. Here, translocation was measured only using FIM (scale = 5 µm).

activation (Fig. 2-12). The crystal structure of beta 2 adrenergic receptor (β2-AR) (PDB

number: 3SN6) with Gαs heterotrimer shows that only Gα interacts with the receptor, while

neither Gβ nor Gγ show physical interactions with the GPCR (66). Collectively, these data

suggest that the heterotrimers that contain Gγ9 universally interact with all GPCRs, and

this Gγ9 assay is universally sensitive to ligand-concentration-dependent Gαi/o- and Gαs-

coupled GPCR activation.

2.2.6. Ligand potency-dependent GPCR activation detection with Gγ9 assay:

A. B.

t=0 After Tizanidine After NE t=0 After NE After Tizanidine

Fig 2-13: Differential Gβγ redistribution between plasma and internal membranes with agonists with different levels of potency for the same receptor. Endogenous α2-ARs in HeLa cells A. activated with Tizanidine first and then Norepinephrine, B. activated with Norepinephrine first and Tizanidine second. Note the change in the PM and IMs fluorescence intensity with each agonist addition.

The ability of γ9 assay to distinguish diﬀerences in the ligand strength was

tested using agonists for α2-ARs; Tizanidine, and NE. A study conducted using rabbit

aortic strips showed that NE is more eﬀective than Tizanidine (67). The γ9 assay shows

that 10 μM NE can increase the FIM/PM response in cells that are already activated with 10

μM Tizanidine, while 10 μM Tizanidine reduces the FIM/PM response of cells that are pre-

activated with 10 μM NE (Fig. 2-13). These data demonstrate that the γ9 assay can

distinguish between strong and weak agonists for the same receptor.

2.2.7. Qualitative Multicellular Screening of Gαi/o- and Gαs- Coupled GPCR

Activation:

High-magniﬁcation ﬂuorescence imaging of FP-γ9 and monitoring of the entire cell

ﬂuorescence change allowed determination of GPCR activation in single cells (Fig. 2-

14A). Using the same images, calculation of the cumulative ﬂuorescence change in

A. B. C. Cumulative multi-cellular Single cells

5 µm Before After Before After

Fig 2-14: Comparison of single and multi-cell GPCR activation by detecting changes in the cumulative FP-γ9 intensity. Confocal Optical cross sections of HeLa cells expressing GFP-γ9, before and after receptor activation are shown. A. Single cell. B. Whole field. C. The plot shows the entire single cell (dim- colored curves) and the entire field (black) fluorescence intensity after M4- muscarinic receptor activation with carbachol. Note the cumulative fluorescence intensity change observed without specifically analyzing IMs or the PM (without ROI selection).

multiple cells also showed a robust increase in ﬂuorescence upon GPCR activation (Fig.

2-14B, C). Using these results as the basis, the feasibility of adopting γ9 assay for high

throughput screening to detect GPCR activation using epiﬂuorescence microscopy was

tested.

Time-lapse images of HeLa cells expressing GFP-Gγ9 and appropriate GPCRs

(when applicable) were captured using an epiﬂuorescence microscope with a 10×

objective, before and after the agonist addition. Images were digitally magniﬁed and IMs

of single cells were selected as ROIs to examine the Gβγ9 redistribution (Fig. 2-15A). A

A. B. Before After

NE 5 µm

Fig 2-15: Qualitative analysis of GPCR activation using low magnification epi- fluorescence. HeLa cells transfected with GFP-γ9 were imaged under 10X objective. A. Digitally magnified images before and after ligand addition show that, even under 10X, the endogenous α2-AR activation induced GFP- γ9 redistribution can be clearly seen in subcellular regions (yellow arrow). The heat map images show the increase in IM fluorescence after NE addition. B. The plot shows no change in the cumulative fluorescence of the entire cell (red). However, ROI (yellow region) based analysis shows that receptor activation induces fluorescence increase in IMs (black).

signiﬁcant increase in ﬂuorescence in IMs was observed upon addition of NE, indicating

the corresponding GPCR activation, as well as heterotrimer dissociation (black trace in

Fig. 2-15B). Although the heat map images clearly showed an increase in IM ﬂuorescence,

26 no change was observed in the whole-cell ﬂuorescence without ROI selection (red trace in

Fig. 2-15B).

Similarly, cumulative time averages of the entire-field fluorescence showed no intensity change upon agonist addition (Fig. 2-16). The large depth of focus that covers the entire height of cells in epifluorescence imaging may have prevented detection of the changes in cumulative fluorescence in GFP-Gγ9 in cells. Therefore, confocal imaging was used to observe an optical cross section across the girth of the cell, only to capture the GFP accumulation in IMs. Under 60× magnification, the cumulative GFP-Gγ9 fluorescence of the entire cell or even the entire field of vision results in an overall fluorescence increase upon GPCR activation (Fig. 2-14). Therefore, confocal imaging is advantageous as the loss of FP-γ9 fluorescence from the limited-detectable PM regions is smaller, compared to the gain of fluorescence signal in relatively abundant IMs. However, even confocal imaging was unable to capture a change in whole-cell fluorescence associated with the ligand- induced GPCR activation under low-magnification imaging.

To understand the reason behind this, the depth of ﬁeld (DOF) was calculated using

Berek’s formula, and the DOF value was determined to be ∼14 μm for the 10×, 0.3 NA objective used in this experiment. Considering this DOF and the average ∼10−20 μm height of a HeLa cell, even with confocal imaging, detection of cumulative entire-cell

ﬂuorescence is unavoidable. In order to eliminate ﬂuorescence signals accumulated in camera pixels from molecules scattered on the PM, the intensity of each pixel (I(x,y)) was readjusted to Ig = {1, where the global threshold was P ≤ I(x,y). The operation was automated by saving time-lapse images as multi- TIFF image stacks, importing them to FIJI, and using its auto-thresholding algorithms. Surprisingly, elimination of I(x,y) from each pixel of the

27 images in the stack resulted in a substantial increase in cumulative ﬂuorescence in the images acquired after GPCR activation. Analysis of 10× time-lapse images using this approach showed an increase in the cumulative ﬂuorescence intensities capturing the activation of receptors (Fig. 2-16).

A. 100X-Raw

25 µm

Before After

B. 100Xthresholded NE

Before After

Fig 2-16: Qualitative analysis of GPCR activation under low magnification epi- fluorescence microscopy. HeLa cells transfected with GFP-γ9 were imaged under 10X objective. A. Raw (unprocessed) images of the entire field of vision under 10X before and after NE. The ligand addition only induces a barely visible fluorescence intensity change while the plot shows that the NE addition does not change the cumulative fluorescence. B. The intensity of thresholded-images in the entire field of vision before and after exposure to 10 µM NE. Yellow arrows show a visible increase in single cell fluorescence after NE. The corresponding plots show an increase in the signal after ligand addition after thresholding, which filters out the noise.

2.2.8. Gγ9 assay is high throughput screening capable:

Pre- and post-agonist GFP-Gγ9 images captured using a Cytation 5 Cell Imaging plate reader show a clear accumulation of GFP in IMs after activation of endogenous α2-

AR with 10 μM NE (Fig. 2-17, top row). Upon thresholding, the post-agonist images show a signiﬁcant intensity increase (Fig. 2-17, bottom row and the plot). This demonstrates that the γ9 assay can be used for high throughput screening of compounds for their ability to activate GPCRs.

Fig 2-17: High throughput screening

adaptability of Gγ9 assay. Plate reader

experiments show high throughput

Raw

screening capabilities of the γ9 assay. A 12 5 µm well glass bottom plate, each well seeded with 0.1 million HeLa cells, were transfected with GFP-γ9. Cells were imaged using a Cytation™ 5 Cell Imaging Multi-

Mode Reader with a 20X objective using Thresholded Thresholded DAPI cube before and after activation of endogenous alpha 2-adrenergic receptors (α2-ARs) using 10 µM norepinephrine (NE). Top row shows images before and after ligand addition. The bottom row shows thresholded-images used to quantify the GPCR activation induced fluorescence intensity changes. Note the increase in fluorescent intensity in IMs after ligand addition (white arrows). The plot shows the normalized cumulative fluorescence indicating a ~50% increase in intensity after NE addition (n=6, *p=0.001).

2.2.9. Gγ9 acts as a spatiotemporal sensor for subcellular GPCR and G protein activation: (Experiment and image courtesy of Fig. 2-18: Kasun Ratnayake)

Once exposed to a spatially restricted extracellular ligand, it is not clear how

GPCRs in a cell confine the activities of G proteins to govern asymmetric signaling and behaviors such as cell migration and neuron development. A lack of approaches to spatiotemporally control signaling in single cells hinders mapping the information flow from the stimulus onset to the GPCR, G protein, and effector activation, as well as the reversal of the processes after stimulus termination. Therefore, such events have mostly been explored using computational models (68). Using the γ9 assay, the accessibility of 29 activated receptors in a localized region to the heterotrimer pool and the span of the G protein activation envelope has been explored.

A. B.

t = 35 s t = 42 s Recovery C. 8 OA-58 s 6

4 m)

μ 2 0 -2 Distance ( Distance -4 -6 0 30 60 90 120 150 180 210 -8 Time (s)

Fig. 2-18: G9 redistribution shows a spatially confined activation of GPCRs and G proteins in subcellular regions. A. A HeLa cell expressing blue opsin– GFP and mCherry-9. White line indicates the region of the 445 nm stimulus pulsed at 0.5 Hz (scale bar: 5 µm). B. The region within the blue box in A is magnified. Upon optical activation (OA) of blue opsin in the line region, mCherry-9 on the PM translocates to IMs resulting in a transient mCherry loss at the activated PM region. The distribution of mCherry on the PM before (t=42 s), during (t=92 s) OA and the recovery after termination of the activation pulse. C. Kymograph generated in ImageJ using the multiple kymograph tool showing mCherry-9 loss on PM and plotted with distance on the PM versus time. Red lines indicate the start and end of OA (t= 42 s to t= 100 s). Yellow dashed line indicates the location of OA on the PM region over time.

A diﬀraction-limited line optical input of 445 nm described in the Materials and Methods section was used to induce the conﬁned activation of GPCRs in a selected PM region.

Time-lapse images of a HeLa cell expressing mCherry-Gγ9 show that localized opsin activation results in a conﬁned loss of mCherry (at 42 s) (Fig. 2-18A, B. Termination of the opsin activation results in the recovery of mCherry-Gγ9 on the PM (Fig. 2-18B, recovery). To reduce the eﬀect of inherent heterogeneity in mCherry-γ9 expression, the pixel intensities were baseline-normalized prior to the kymograph generation. The kymograph which was generated using the line ROI shows the mCherry-9 loss on PM

(around the active site) over time (Fig. 2-18C).

2.3. CONCLUSION:

Contrary to limitations in current assays to detect GPCR-G protein activities, the

γ9 assay described here provides (i) an enhanced detection ability of G protein activation, even at low ligand concentrations (higher detection limit), (ii) monitoring of native GPCR-

G protein interactions, (iii) direct detection of G protein activation and deactivation by a

GPCR, (iv) detectability of spatially and temporally conﬁned GPCR-G protein activation, and (v) adaptability for high throughput screening. Therefore, a combination of these characteristics in the γ9 assay allows quantitative measurement of receptor-ligand interactions and their dose-response relationships. Data further show that cells can achieve a series of graded G protein activation states by reaching multiple dose-dependent steady states. This may allow cells to control a series of unique dose-dependent signaling events for a single ligand. Measurement of the G protein activation envelope suggests that, during spatially restricted GPCR activation, cells have mechanisms in place to contain the localized active regions, so that cells can continually maintain internal signaling gradients and direct polarized cell behaviors. Overall, the studies presented here conclusively 31 demonstrate that, reversible Gγ9 distribution can be adopted as a versatile assay for qualitative and quantitative assessment, as well as a subcellular sensor for GPCR-G protein interaction in living cells.

2.4. MATERIALS AND METHODS:

2.4.1. Constructs, Cell culture, and Transfections:

Constructs, GFP-γ9, mCherry-γ9, and blue opsin-GFP have been described previously (24, 55, 69, 70). β1-AR-CFP was a gift from Professor N. Gautam, Washington

University, St. Louis, MO. M4 muscarinic-mTurquoise was created by PCR ampliﬁcation of M4 with NotI and XbaI from M4-CFP and sub cloning to corresponding sites of blue opsin-mTurquoise, after restriction digestion. This construct was in pcDNA3.1

(Invitrogen). HeLa cells (ATCC) were cultured in minimum essential medium (CellGro) containing 10% dialyzed fetal bovine serum (Atlanta Biologicals), in the presence of 1% penicillin−streptomycin in 60 mm tissue culture dishes. At 75% conﬂuency, cells were lifted after incubating with versene-EDTA (CellGro) for 3 min at 37 °C, centrifuged at

1000g for 3 min, and versene-EDTA was aspirated before resuspending in the regular culture media at a cell density of 1 × 106/mL. One day before the transfection of DNA into cells, 1 × 105 cells were seeded on 35 mm glass-bottomed dishes (In Vitro Scientiﬁc). The transfection was performed using the transfection reagent PolyJet (SignaGen), according to the manufacturer’s protocol.

2.4.2. Time-lapse imaging:

These experiments were performed with a 60×, 1.4 NA oil objective or a 10×, 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 FRAP-PA (ﬂuorescence recovery after photobleaching and photoactivation) module, a laser combiner with 40−100 mW 445, 488, 515, and 594 nm solid-state lasers and iXon ULTRA 897BV back-illuminated deep-cooled EMCCD camera. Fluorescently tagged-γ9 (FP-γ9) translocation in cells, cultured on glass- bottomed imaging dishes, was examined by imaging either GFP or mCherry, using 488 nm excitation−515 nm emission or 594 nm excitation−630 nm emission settings, respectively. During time lapse imaging at a frequency of 1 Hz, ligands were added at 2× concentration to activate corresponding

GPCRs in appropriate volumes to achieve efficient diffusion and 1×final concentration in the imaging media. Imaging was continued until fluorescence intensities of the PM and

IMs reached the ﬁnal equilibrium.

2.4.3. Optogenetic control of GPCR signaling and imaging of G protein dynamics:

HeLa cells expressing blue opsin-GFP and mCherry-γ9 were cultured on 35 mm glass- bottomed imaging dishes, as described above. Using a 445 nm optical stimulus, opsins in cells were activated globally or locally using a computer-steered galvo device.

2.4.4. Image Analysis, Data Processing, and Simulation of Laser Power Distribution:

Time-lapse images were analyzed using the analytical tools accompanied by Andor iQ 3.1 software (Andor Bioimaging). Additional image analysis was performed using

ImageJ (National Institute of Health) (71), and custom-built Python algorithms (Python

Software Foundation).

Chapter 3

Gγ Subtype Dependent Plasma Membrane Affinity of

Gβγ Controls the Efficacy of Downstream Signaling

Activation and Macrophage Migration

This chapter contains materials which were originally published in the Journal of Biological Chemistry. Kanishka Senarath, John L. Payton, Dinesh Kankanamge, Praneeth Siripurapu, Mithila Tennakoon, and Ajith Karunarathne. Gγ identity dictates efficacy of Gβγ signaling and macrophage migration. Reprinted with permission from the Journal of Biological Chemistry 2018, 293 (8), 2974-2989. Copyright © 2018 American Society for Biochemistry and Molecular Biology, Inc. (ASBMB).

3.1. INTRODUCTION:

G protein coupled receptors (GPCRs) primarily transduce signals by activating G protein heterotrimers consisting of Gα and Gβγ subunits. Active G proteins GαGTP and

Gβγ interact and control a cohort of effectors and regulate the majority of metazoan signaling (16, 72, 73). Although Gα signaling has been the primary focus in the field, recent findings show that Gβγ subunits regulate even more pathways and cellular functions. Some of these Gβγ effectors include phosphatidylinositol-4,5-bisphosphate 3-kinase (PI3Kγ), adenylyl cyclase (AC) isoforms (activation of AC2, 4, 7, and inhibition of AC1, 5), G protein-gated inwardly rectifying potassium (GIRK) channels, PLC isoforms (PLCβ2, β3),

Ca2+ channels (N, P/Q type), GPCR kinases (GRKs), and guanine nucleotide exchange factors (GEFs) such as Ras-related C3 botulinum toxin substrate 1 (Rac1), cell division control protein 42 (Cdc42), guanine nucleotide exchange factor (FLJ00018), and p114-

RhoGEF (23, 74-82). These effectors coordinate a wide range of cellular and physiological functions such as cellular secretion, gene transcription, contractility, and cell migration, and are therefore involved in numerous pathological conditions including cancer and heart disease (16, 72, 73).

Among Gβγ controlled activities, cell migration plays a key role in many physiological functions including embryonic development and immune responses. Hence, altered cell motilities are implicated in pathological processes such as immune deficiencies, lack of wound healing, tissue repair and cancer metastasis (83-86). We have recently shown that Gβγ is the primary regulator of macrophage migration (87). In addition to PI3K-PIP3 signaling that primarily controls leading edge functions, we demonstrated that Gβγ also activates the PLCβ pathway which is essential for macrophage migration (87).

Mammalian cells express twelve Gγ and five Gβ subunits and form stable Gβγ dimers with the exception of Gβ5, giving rise to 48 possible combinations of Gβγ (88, 89).

It has been shown that most Gγ subtypes comparably interact with the two most predominant Gβ types in cells, Gβ1 and Gβ2, with the exception of Gγ11 for Gβ2 (90).

Similar affinities of Gαi1 for Gβγ types have also been demonstrated (89). Studies have suggested the possibility of specific Gβγ subtypes possessing higher affinities towards certain GPCRs or effectors. Using in vitro reconstituted heterotrimers and activated

GPCRs, heterotrimers with certain Gγ subtypes exhibited higher affinities for specific

GPCRs (89). In addition, specific structural motifs in GPCRs preferring certain Gβγ

35 isoforms have been reported for adenosine family receptors (91, 92). Assigned cellular functions to the availability of specific Gβ or Gγ subtypes have also been shown (93, 94).

For instance, modulation of Golgi vesiculation and cellular secretions by Gγ11 and differential ion channel control by Gγ9 and Gγ3 subunits have also been demonstrated (93,

94). Gγ3 and Gγ5 were shown to control predisposition of mice to seizures (95).

While these investigations have primarily assigned subunit identity of either Gβ or

Gγ subtype to specific signaling activities and cellular functions, molecular and mechanistic basis of such signaling specificity has not been provided. Gβ subunits have a conserved structure with a >80% identity among their isoforms. However, Gγ isoforms show a significant sequence diversity ranging from ~ 20-80% (37, 88). Therefore, if the

Gβγ diversity is a crucial modulator of its signaling and associated cell behaviors, the Gγ identity in these dimers is likely to be the primary regulator of Gβγ signaling. Although

Gβγ is considered PM bound, recent work has shown that, upon GPCR activation, Gβγ translocates from the PM to IMs until an equilibrium is reached (94). Interestingly, translocation half time (Tt1/2) and the extent are governed by the type of accompanying Gγ subunit (24, 55). These results further suggest that the PM-affinity of a Gβγ is Gγ subtype dependent. Since, Gγ provides the only PM contact point for Gβγ, the accompanying Gγ subtype-dependent differential translocation ability of G suggests that, Gγ type controls the PM-affinity of Gβγ. Since, the majority of Gβγ-effector interactions take place at the

PM, Gγ-governed PM-affinity is likely to be crucial for Gβγ signaling. This study was focused on examining how cells employ a selected group of Gγ subunits to tune signal propagation from activated GPCRs to Gβγ effectors, controlling signaling and macrophage migration.

3.2. RESULTS:

3.2.1. Gγ subtype identity-specific control of PI3Kγ activation by Gβγ:

PIP3 is a major regulator of lamellipodia formation in the leading edge of migratory cells (96). Class IB PI3Kγ, which is composed of cytosolic non-catalytic p101 and catalytic p110 subunits, has been identified to be activated by Gβγ (97, 98). Since Gβγ-

PI3K interaction leads to the PIP3 generation (Fig. 3-1), we examined whether PIP3 production is controlled in a Gγ subtype dependent manner. To interact with Gβγ and

Ligand

   p p PI3K p p p PIP2 PIP3 p p Akt-PH mCh

Fig. 3-1: PIP3 generation measurement using the PIP3 sensor. Fluorescently tagged PIP3 sensor; Akt-PH-mCherry, translocates from the cytosol to the PM to interact with the PIP3 generated on the PM. Accumulation of the PIP3 sensor fluorescence is proportional to the Gβγ mediated PI3Kγ activation induced PIP3 generation. catalyze PIP3 production, PI3K subunit p110 should translocate to the PM upon activation

(97, 98). The signaling circuit that drives PIP3 production is composed of GPCRs, Gβγ, and PI3Kγ subunits. PIP3 generation was measured using translocation of a fluorescently tagged PIP3 sensor (Akt-PH-mCherry) from cytosol to the PM (Fig. 3-1).

A. B. Akt - PH - mCh C.

GPCR

Gβγ

PI3Kγ

PIP3 RAW 264.7 HeLa Before After

D. GFP - Gγ 9 GFP - Gγ3 E.

HeLa

RAW RAW

Before After Before After

Fig. 3-2: Differential PIP3 generation in different cell lines. A. Pathway for Gβγ mediated PI3K activation. B. WT HeLa and RAW 264.7 cells expressing the PIP3 sensor; Akt-PH-mCherry and blue opsin-mTurquoise. Cells supplemented with 50 µM 11-cis- retinal were imaged every 5 s for mCherry (with 594 nm). Blue opsin activation with 445 nm blue light induced translocation of cytosolic PIP3 sensor to the PM only in RAW cells but not in HeLa cells. The plot shows the accumulation Akt-PH-mCherry on the PM. Blue arrow represents the initiation of optical activation (at 30 s). Intensities are baseline normalized (Scale bar: 5 µm, error bars: SEM). Comparison of GPCR activation induced Gβγ translocation in HeLa and RAW cells. D. HeLa and RAW cells transiently transfected with GFP-Gγ9 and GFP-Gγ3 show similar translocation in terms of rate and the extent with receptor blue opsin activation. HeLa cells were imaged at 60X magnification and RAW cells were at 100X. E. Graph shows the GFP fluorescence intensity increase in IMs due to Gβγ translocation. Note the similarity of Gγ9 and Gγ3 translocation in both cell types (Error bars: SEM, n=8; scale: 5 µm).

Gβγ generated upon activation of Gi-coupled GPCR, blue opsin, induced a robust

PIP3 production in RAW cells (Fig. 3-2B-top, 3-2C), while HeLa cells failed to produce an observable PIP3 response (Fig. 3-2B-bottom, 3-2C). Interestingly, the translocation of

YFP-Gγ9 and YFP-Gγ3 upon blue opsin activation was comparable in HeLa cells and

RAW cells, indicating equivalent G protein activations (Fig. 3-2D, E).

Similar to HeLa cells, PC12 cells also failed to show PIP3 production upon GPCR activation. However, unlike HeLa cells, PC12 cells exhibited augmented PIP3 generation upon expression of PI3Kγ (Fig. 3-3).

PI3K-CFP Akt-PH-mCh

PC12 HeLa HeLa PC12

Before After Fig. 3-3: PIP3 generation upon PI3Kγ expression. PI3K expression in a HeLa cell failed to induce PIP3 generation on blue opsin activation (black trace). PC12 cells that showed no PIP3 response, elicited a robust repose upon expression of PI3Kγ (green trace). Blue arrow indicates optical activation. (Scale bar: 5 µm, Error bars: SEM).

Real-time PCR data from RAW and HeLa cells revealed that they express substantially different Gγ subunit profiles (Fig. 3-4A). Compared to RAW cells,

HeLa cells show a ~6-fold lower expression of Gγ3, and expression of Gγ4 is also significantly lower in HeLa cells. In order to examine if Gγ3 or Gγ4 is the missing signaling component in HeLa cells required to induce PIP3 production, we expressed them and

39 studied opsin activation-induced PIP3 production. Expression of Gγ3 resulted in an elevated basal PIP3, even without GPCR activation (Fig. 3-4C).

A. B.

C. Gγ4 Gγ3

mCh

- Akt Before After Before After

Gγ9

mCh

- Akt Before After

Fig. 3-4: Gγ identity-controlled PIP3 generation. A. Comparison of real time PCR Gγ profiles of HeLa and RAW cells. HeLa cells express mRNA for Gγ12 and Gγ5 in abundance, while Gγ4 and Gγ3 are prominent in RAW cells. B. Gγ9 (red) and Gγ3 (green) overexpression induced changes to the Gγ profile in HeLa cells. The overexpressed Gγ type appears to dominate native Gγ. C. HeLa cells expressing Gγ3, blue opsin-mTurquoise, and Akt-PH-mCherry showed an intense PIP3 generation compared to the WT cells upon blue opsin activation. Images and the plot show Gγ4 expression showed a minor (blue trace), while Gγ9 showed no PIP3 generation (green trace), compared to WT (black trace) and Gγ3 (red trace) on the PM. The plot shows the corresponding PIP3. Intensity values are baseline normalized, blue arrow indicates optical activation (Scale bar: 5 µm, n= 8, Error bars: SEM).

Blue opsin activation subsequently resulted in a robust PIP3 production in these cells.

Although Gγ4 expression did not promote elevation of basal PIP3, opsin activation exhibited a minor increase in PIP3 (Fig. 3-4C-yellow arrows). Nevertheless, Gγ9 expressing cells failed to induce PIP3 production either at the basal state or upon opsin activation (Fig. 3-4C). Real-time PCR data indicated that overexpression of a Gγ subunit results in the reduction of the fractional contribution of endogenous Gγ subunits to the pool, making the introduced Gγ subunit dominant, creating nearly a mono-Gγ system (Fig. 3-

4B).

3.2.2. Optogenetic determination of PM affinities of 12 Gγ subunits using Tt1/2 of Gγ:

GPCR activation by ligand addition is prone to experimental artifacts which obstructs precise measurement of kinetics in G protein activation, while OA avoids delays and inconsistencies in ligand addition and allows for the calculation of precise Tt1/2 as well as extent of translocation. Thus, using optically controlled activation of GPCR-G protein signaling, by employing blue opsin supplemented with 11-cis retinal, we determined precise Gβγ translocation parameters as follows. HeLa cells expressing YFP tagged 1-13

Gγ types respectively, were examined for translocation of Gβγ (Fig. 3-5A, B). The Gβγ translocation was measured using YFP fluorescence dynamics in IMs (FIM vs time curves)

(Fig. 3-5B). Since GPCR activation results in an approximately sigmoidal increase in Gβγ in IMs, which reaches saturation over time (Fig. 3-5B), the FIM vs time curves were fitted

푇푚푎푥−푇푏푎푠푒 to the logistic function (퐹퐼푀 = 푝 + 푇푏푎푠푒) in OriginPro. Using these fits, Tt1/2 and 1+(푡1/2/푡0ሻ

the extent of translocation |T| = (푇푚푎푥 − 푇푏푎푠푒) of individual Gγ subtypes were calculated

(Table 3-1). The plot of |T| vs Tt1/2 (Fig. 3-5C) exhibits a strong correlation (adjusted

R2=0.94) between the two.

A. Gγ1 Gγ2 Gγ3 Gγ4 Gγ5 Gγ7 Gγ8 Gγ9 Gγ10 Gγ11 Gγ12 Gγ13

Basal s 50 600 s B. C.

Fig. 3-5: Gγ-identity driven differential translocation of Gβγ. A. HeLa cells expressing blue opsin-mTurquoise and each of the 12 Gγ subunit with a YFP fluorescent tag. Cells supplemented with 50 µM retinal were imaged for YFP (515 nm) and activated with 445 nm light at 2 s intervals in a time-lapse series. This process was continued for 10 mins where the YFP fluorescence changes reached the equilibrium. B. Plots show baseline normalized YFP fluorescence increase in IMs over time (error bars: SEM, n= 10; Scale bar: 5 µm). C. Plot of Tt1/2 vs. |T| shows an exponential decay relationship.

Considering the link between half time of Gβγ translocation (Tt1/2) and the extent of translocation (|T|) (Fig. 3-5C), it suggests that they are likely to be controlled by the ability of Gβγ to interact with the PM (Fig. 3- Table 3-1: Translocation properties of WT Gγ types

6). Tt1/2 values of Gγ9 translocation were identical in HeLa and RAW cells (Fig. 3-2D,

E). This demonstrates that translocation properties of Gγ types are conserved among cell types, suggesting conserved PM-affinities of Gγs. Although Gγ only possesses two types of lipid anchors (geranylgeranyl and farnesyl) at their CaaX motif, data shows that, they have a discrete series of Tt1/2 values (Table 3-1).

Therefore, distinct PM-interacting pre-CaaX

Fig. 3-6: Gγ-identity driven differential PM-affinity of Gβγ. Schematics of GPCR activation induced G protein activation and dissociation. LoAf-Gβγ translocate from the PM to IMs faster compared to HiAf-Gβγ, while HiAf- Gβγ interact with effectors to initiate signaling pathways leading to cellular responses efficiently compared to LoAf-Gβγ.

43 motifs of Gγ subunits appear to provide additional control over their PM affinities, resulting in a spectrum of Tt1/2 values.

3.2.3. Gγ-dependent control of chemokine pathway mediated RAW cell migration:

Since the subtype identity of Gγ controls PIP3 formation, we examined whether cell migration is also similarly controlled. We have previously shown that localized blue opsin activation results in a robust PIP3 production at the leading edge and directional motility of RAW264.7 cells (99). Since both blue opsin and chemokine receptor 4

(CXCR4) activate G proteins with nearly similar efficacies, blue opsin was employed to induce macrophage migration (Fig. 3-7). Real-time PCR data showed that ~30% of Gγ in

WT RAW 264.7 cells is Gγ3, a high PM-affinity (HiAf) Gγ type (Tt1/2 = ~270 s) (Fig. 3-

4A).

A. B. C.

with CXCR4 with blue opsin

After Before Before After

Fig. 3-7: Comparison of optical and chemical induced GPCR activation. A. A HeLa cell transiently transfected with A. mCherry-Gγ9 and, B. another cell with blue opsin-GFP + mCherry-Gγ9 expression showed Gγ9 translocation from the PM to IMs A. with endogenous receptor CXCR4 activation with 50 ng/µL SDF-1α and B. blue opsin activation with 50 µM 11-cis retinal addition followed by 5 min incubation in dark and OA with 445 nm blue light. C. Graph showsthe similar Gγ9 translocation with both optical and chemical induced GPCR activation comparatively (error bars: SEM, n=8; scale bar: 5 µm).

In response to localized optical activation of blue opsin, RAW cells migrate efficiently with the leading-edge velocity (LE) of 0.82 µm/min and trailing-edge velocity

A. WT Gγ1 Gγ2 Gγ3

t=0 20 0 20 0 20 0 20 Gγ4 Gγ9 Gγ3 shRNA Gγ3 shRNA+ Gγ2

0 20 0 20 0 20 0 20 mins

B. Fig. 3-8: Subtype specific control of macrophage migration by Gγ. A. RAW 264.7 cells expressing blue opsin-mCherry and a selected Gγ subunit. Blue opsin was activated in confined regions of cells using a 445 nm laser with 0.22 µW/µm2 power in every 2 s interval (white boxes). The images show cells before and after 20 mins of blue opsin activation. Note the difference in cell movement towards the optical input with respect to the Gγ type the cell possesses. Gγ3 expressing cell shows an almost identical cell migration as the WT, and Gγ2 also supports migration. Note the inhibition of cell migration in Gγ3 knocked down cells. This migration loss was rescued by expressing HiAf-Gγ2, but none other. B. Bar graph shows the relative displacement of cells’ leading and trailing edges, with BO activation (error bars: SEM, n= 12, * P= 0.021, ** P< 0.0001, *** P< 0.0001; scale bar: 5 µm).

(TE) of 0.51 µm/min (Fig. 3-8A, B). High PM-affinity Gγ3 and Gγ2 overexpression also

45 showed good cell migration while expression of low PM-affinity (LoAf) Gγ subtypes showed a marked reduction of migration, i.e. Gγ9 (Tt1/2= ~5 s)→ LE:0.20 µm/min, TE:

0.03 µm/min and Gγ1 (Tt1/2= ~13 s)→ LE: 0.24 µm/min, TE: 0.04 µm/min (Fig. 3-8A,

B). Expression of moderate PM- affinity (MoAf) Gγ4 (Tt1/2= ~116 s) reduced the migration ability of WT RAW cells substantially (LE: 0.38 µm/min, TE: 0.18 µm/min). LoAf-Gγ expressing cells occasionally showed lamellipodia formation at the leading edge, while trailing edge retraction was not observed.

Knockdown of endogenous Gγ3 using the most effective shRNA identified by screening five constructs resulted in a complete cessation of cell migration (Fig. 3-8A, B).

Non-specific shRNA did not affect WT RAW cell migration. Expression of HiAf-Gγ2

(Tt1/2 = ~181 s) in Gγ3 knocked down cells, resulted in rescuing the lost migration ability with LE:0.61 µm/min and TE:0.28 µm/min (Fig. 3-8A, B).

To examine the universal nature of HiAf Gγ subunit requirement in chemokine pathways, we examined if the introduction of HiAf-Gγ3 helps non-migratory HeLa cells to migrate. Localized opsin activation in HeLa cells expressing Gγ3 showed a distinct trailing edge retraction, with lamellipodia formation at the leading edge resulting in net

t =0 t = 40 mins Fig. 3-9: Universal nature of HiAf Gγ subunit

requirement in cell migration. Expression of

Gγ3 ov Gγ3

– Gγ3 in HeLa cells induces cytoskeleton changes leading to lamelipodia formation at

HeLa HeLa the leading edge and trailing edge retraction

upon localized optical activation of blue opsin

(white boxes), while Gγ9 expressing cell (or Gγ9 ov ov Gγ9

WT cells-data not shown) showed no such –

motility (scale bar: 5 μm).

HeLa HeLa

46 movement of cells. No responses were observed in WT or Gγ9 expressing HeLa cells for similar signaling activation (Fig. 3-9).

A. B. Gγ3 WT ---- NPFREKKFFCALL Gγ Gβ Gγ3 with Gγ9 CaaX ---- NPFREKKFFCLIS Gγ3 with Gγ9 CT ---- NPFRE-KGGCLIS Gγ9 WT ---- NPFKE-KGGCLIS Gγ9 with Gγ3 CaaX ---- NPFKE-KGGCALL Gγ9 with Gγ3 CT ---- NPFKEKKFFCALL

C. Gγ3 with Gγ9 CaaX Gγ9 with Gγ3 CT

t=0 t=50s t=600 s t=0 t=50s t=600 s

Gγ3 with Gγ9 Tt1/2 (s) CT 5±2 14±2 21±3

t=0 t=50s t=600 s Gγ9 with Gγ3 CaaX

37± 1863 ± 5270 ± t=0 t=50s t=600 s 4

Fig. 3-10: Carboxy terminus of Gγ governs rates of Gβγ translocation. A. RAW Crystal structure of the CT region of Gγ in complex with Gβ (~F59 of Gγ- the last Gβ contact point) exposing the hydrophobic binding pocket in Gβ. B. Sequence alignment of CT mutants of Gγ3 and Gγ9. C. HeLa cells expressing GFP-Gγ mutants and blue opsin-mCherry, supplemented with 11-cis retinal. The cells were imaged for GFP (488 nm) to capture blue opsin activation induced Gβγ translocation. Note the significant difference in mutant translocation compared to WT counterparts. (error bars: SEM, n=10; scale bar: 5 µm).

3.2.4. Carboxy terminus of Gγ governs rates of Gβγ translocation:

CT sequences of Gγ exhibit significant diversity (Fig. 3.6B) (37, 88). Sequence alignment and structural data show that, after the conserved F59 in all Gγ subtypes (except

Gγ13), CT region loops out from a conserved hydrophobic pocket on Gβ, delineating the last contact point with Gβ (Fig. 3.10A) (100). The pre-CaaX region (region between F59 and the CaaX) therefore appears to interact with the PM and partially modulates the PM- affinity of Gβγ. Lack of electron density in Gβγ crystal structures indicates that, CT of Gγ is unstructured and suggests flexible and dynamic interactions. Therefore, next we employed a group of Gγ mutants comprising the body of HiAf-Gγ with substituted CaaX and or pre-CaaX motifs from LoAf-Gγ and vice-versa (Fig. 3.10B). Translocation properties of these mutants depicted characteristics of Gγ in which the introduced motifs originated (Table 3-2). For instance, Gγ9 with pre-CaaX + CaaX of Gγ3 (Gγ9-γ3CT) exhibited similar translocation properties to Gγ3. On the contrary, Gγ3 with pre-CaaX +

CaaX regions of Gγ9 (Gγ3-γ9CT) exhibited similar translocation properties to Gγ9 (Fig.

3.10C).

Table 3-2: Translocation properties of Gγ mutants

3.2.5. Control of RAW cell migration by CaaX and pre-CaaX residues in CT of Gγ:

Since CT of Gγ provides sites for Gβγ to anchor and interact with the PM, the properties of G CT on RAW cell migration was also examined expressing above mentioned mutants. For instance, Gγ9-γ3CT mutant induced cell migration. On the contrary, cells expressing Gγ3-γ9CT mutant failed to migrate, recapitulating migration behavior of RAW cells expressing Gγ9 (Fig. 3-11).

A. Gγ3 with γ9 CaaX Gγ3 with γ9 CT Gγ9 with γ3 CaaX Gγ9-γ3 CT

t=0 20 0 20 0 20 0 20 mins

B. Fig. 3-11: Carboxy terminus of Gγ governs the extent of cell migration. A. RAW 264.7 cells expressing each of the Gγ mutant and blue opsin-mCherry, supplemented 11-cis- retinal. Blue opsin in cells were activated locally (white box) in 2 s intervals for 20 min to induce migration. B. Histogram shows the movement of leading and trailing edges. Permutations to the CT sequences clearly altered the cell migration ability (error bars: SEM, n=12, * P= 0.0009 for the leading edge and 0.5714 for the trailing edge, ** P< 0.0001, *** P< 0.0001, **** P< 0.0001; Scale bar: 5 μm).

3.2.6. Modulation of RAW cell migration potential by Gγ subtype-dependent activation of PI3Kγ:

Since PIP3 is a key regulator of chemokine induced cell migration, we examined if PIP3

production is Gγ-type dependent. RAW cells expressing Akt-PH-mCherry PIP3 sensor

showed significant PIP3 accumulation at the leading edge upon localized optical activation

of blue opsin (Fig. 3.12B). Inhibition of Gβγ with gallein and PI3Kγ with wortmannin,

ceased PIP3 production as well as migration (Fig. 3-12A, B). Gallein-like compound,

fluorescein, was used as a control which did not show any effect.

A. B.

t=0 10 mins

GPCR

WT WT

Gallein G Gallein Gallein

PI3K Wortmannin

PIP3 Wortmannin Wortmannin

Fig. 3-12: Gβγ mediated PI3Kγ activation induced PIP3 generation and cell migration. A. GPCR mediated PIP3 generation pathway and its selected inhibitory points. B. RAW 264.7 cells expressing Akt-PH-mCherry and blue opsin, supplemented with 50 µM 11-cis-retinal. On localized blue opsin activation with 445 nm (white box), WT cells showed the PIP3 production at the activated leading edge. Cells treated with PI3K inhibitor; wortmannin and Gβγ inhibitor gallein inhibited both PIP3 production and cell migration, confirming that PIP3 is required for directional cell migration (n=10; Scale bar: 5 µm).

Cells expressing Gγ3 showed a PIP3 production at the leading edge and directional migration similar to the responses exhibited by WT RAW cells (Fig. 3-13). Plots show that

Gγ9 expressing RAW cells exhibit mild or no PIP3 production. These cells failed to

A. B. WT Gγ3 Gγ3 shRNA

t=0 10 0 10 0 10

Gγ4 Gγ12 Gγ9

0 10 0 10 0 10 mins

Fig. 3-13: Gγ type dependent activation of PI3Kγ during macrophage migration. A. RAW 264.7 cells expressing Akt-PH-mCherry, blue opsin-mTurquoise, a Gγ subunit (Gγ3, Gγ9, Gγ4, Gγ12) and supplemented with 50 µM 11-cis- retinal. On localized blue opsin activation with 445 nm (white box), Gγ3 expressing cells showed PIP3 generation at the leading edge. However, Gγ4, Gγ12 and Gγ9 expressing cells showed minor/no PIP3 accumulation. Gγ3 knocked down cells also showed no PIP3 generation. B. Plots show the PIP3 generation with Gγ3, Gγ9, Gγ4, Gγ12 overexpression and Gγ3 knockdown compared to the WT (error bars: SEM, n=10; scale: 5 µm). migrate as well. Gγ3 knocked down cells neither showed PIP3 production at the leading edge nor cell migration upon opsin activation (Fig. 3-13).

Additionally, RAW cells expressing Gγ3 mutants composed of either pre-CaaX or

CaaX motifs or both from Gγ9 failed to induce PIP3 production or cell migration (Fig. 3-

14). Interestingly, cells expressing Gγ9-γ3CT mutant showed both PIP3 production and

cell migration. However, Gγ9 mutants with either pre-CaaX alone or CaaX alone from Gγ3

failed to show PIP3 production or cell migration. This can be understood by examining PM

affinities (Tt1/2 values) listed in Table 3-2, in which they show the order; Gγ3 > γ9-γ3CT >

γ3-γ9CaaX > γ9-γ3CaaX > γ3-γ9CT > γ9. Interestingly, the data illustrates that cells

A. Gγ9 with γ3 CaaX Gγ9 with γ3CT Gγ3 with γ9 CaaX Gγ3 with γ9 CT

t=0 10 0 10 0 10 0 10 mins

B. Fig. 3 - 14: Carboxy terminus of Gγ dependent activation of PI3Kγ activation and macrophage migration. RAW 264.7 cells expressing Akt-PH-mCherry, blue opsin-mTurquoise, and each of the mutant Gγ types. Upon migration induction, cells expressing the mutant Gγ9-γ3CT showed both PIP3 as well as migration. Failure to exhibit migration in Gγ9-γ3CaaX cells shows the significance of the pre-CaaX motif of Gγ in Gβγ signaling. Gγ3-γ9CT mutant cells exhibited neither PIP3 production nor migration. Plot shows PIP3 generation in RAW cells expressing CT mutants of Gγ (error bars: SEM, n=15; Scale bar: 5 µm).

expressing only HiAf-Gγ subtypes, including Gγ3, Gγ2, and Gγ9-γ3CT mutant elicited a

robust PIP3 generation, which may be essential for cell migration. Also, both WT and

HiAf-Gγ3 expressing RAW cells possess comparable mean rates of PIP3 production;

0.0022 sec-1 and 0.0030 sec-1, respectively. However, the mean rate of PIP3 generation in

MoAf-Gγ4 expressing cells (0.0009 sec-1) was closer to Gγ9 (0.0007 sec-1) and Gγ12

(0.0012 sec-1) than to Gγ3.

3.2.7. Gγ subtype - dependent control of Gβγ mediated PLCβ activation:

We examined if PLCβ activity in RAW cells is also controlled in a Gγ subtype dependent manner, in the same way it controlled PI3Kγ activation. We have demonstrated that Gi pathway-induced RAW cell migration requires an increase in cytosolic Ca2+, which is governed by Gβγ mediated activation of PLCβ to induce trailing edge retraction (87).

Ca2+ mobilization upon endogenous Gi-coupled complement component 5a receptor

(c5aR) activation with 10 µM c5a (101) was measured using a fluorescence probe for calcium (Ca2+), fluo-4-AM. WT and HiAf-Gγ3 expressing cells showed Ca2+ responses to a higher degree, while LoAf-Gγ9 expressing RAW cells showed minor or no Ca2+ response upon c5aR activation (Fig. 3-15A-D). Unexpectedly, MoAf-Gγ4 and Gγ12 expressing cells exhibited only a weak response (Fig. 3-15E, F). Replacement of the entire CT or CaaX motif alone in Gγ3 with those of Gγ9 respectively, resulted in a loss of Ca2+ mobilization ability of Gγ3 (Fig. 3-15G, H). Although, expression of Gγ9-γ3CaaX mutant failed to elicit

Ca2+ mobilization, mutant Gγ9-γ3CT showed a Ca2+ response, which is equivalent to responses exhibited by WT as well as Gγ3 expressing RAW cells (Fig. 3-15G, H). Also, we confirmed that resultant Ca2+ responses are similar for Gγ with different fluorescent tags (Fig. 3-15A-D).

mCherry-GPI Untransfected

Before Response After Before Response After D. B. c5a

mCherry-γ3 GFP-γ3

Before Response After Before Response After

mCherry-γ9 GFP-γ9

Before Response After F. Before Response After

E. YFP-γ4 YFP-γ12 c5a

Before Response After Before Response After

H. G. GFP-γ3 with γ9 CaaX GFP-γ3 with γ9 CT c5a

Before Response After Before Response After GFP-γ9 with γ3 CaaX GFP-γ9 with γ3 CT

Before Response After Before Response After

Fig. 3-15: PLCβ activation induced differential Ca2+ response with different Gγs. RAW 264.7 cells expressing different WT Gγs and Gγ mutants were stimulated with 10 µM c5a addition to activate endogenous c5a receptors (c5aRs) after 30 min fluo-4 incubation. Cells were imaged at 40X magnification to capture the Ca2+ response. A. Control (mCherry-GPI and untransfected) and, B. Gγ3 expressing cells showed greater Ca2+ response compared to, C. Gγ9 expressing cells, which showed almost no Ca2+. Scale bar, 10 µm. D. Plot shows the difference in fluo-4 signal (GFP fluorescence) increase in cells, indicating differential Ca2+ release to the cytoplasm depending on the Gγ subtype they overexpress. Also, it shows that the fluorescent tag of the Gγ subtype is not affecting the Ca2+ response (n=8). E, F. MoAf-Gγ4 and Gγ12 expressing cells showed minor Ca2+ response with c5aR activation (n=8). G, H. Gγ9 mutants with Gγ3 CaaX and Gγ3 CT showed an increased Ca2+ response compared to WT Gγ9, while Gγ3 with Gγ9 CaaX and Gγ9 CT showed a reduced Ca2+ response compared to WT Gγ3, confirming differential Gβγ-effector interactions with respect to the difference in the PM affinity thus different PM residence times of Gβγ (error bars: SEM, n=9).

3.2.8. Tt1/2 of Gγ is a strong predictor of Gβγ effector activation ability:

The purpose was to examine if Gβγ effector responses of a cell can be predicted using averaged Tt1/2 of endogenous Gγ. The experimental process used to test this model is given in Fig. 3-16A. We examined PIP3 production in HeLa cells upon optical activation of blue opsin for each of the 12 Gγ subtypes by imaging the PIP3 sensor; Akt-PH-mCherry

퐺훾 (Fig. 3-16B, C). The extent of effector activation|퐸퐹|푒푥푝, upon Gγ expression was measured using baseline-normalized increase of mCherry fluorescence at the PM due to accumulation of generated PIP3 (Fig. 3-16B, C). Tt1/2 values of each Gγ type translocation were also similarly calculated by measuring YFP-Gγ translocation (Fig. 3-5).

C. D.

Fig. 3-16: Tt1/2 of Gγ as a predictor of a cell’s ability to control Gβγ effectors. A. Experimental process of predicting Gβγ effectors activity using Gγ type dependent PM affinity (Tt1/2). B. Plots showing the extent of blue opsin activation induced PIP3 generation in HeLa cells expressing each of the 12 Gγ types. C. Smoothed and logistic function fitted curves of PIP3 generation described in B. D. Plot of |EF| vs Tt1/2 of all 12 Gγ types. The |EF| was measured using PIP3 production on the PM in HeLa cells expressing each of the 12 Gγ types and Akt-PH-mCherry.

In order to examine whether predictions about Gβγ effector responses of a cell type can be made using averaged translocation properties of its endogenous Gγ, we first plotted

퐺훾 2 |퐸퐹|푒푥푝 vs Tt1/2 (HeLa effector plot) (Fig. 3-16D). The fitted straight line exhibited an R

퐺훾 value of 0.94, indicating that the |퐸퐹|푒푥푝 values were linearly related to the corresponding

Tt1/2 (PM-affinity) values of Gγ type expressed in HeLa cells, in the order of

Gγ3>γ2>γ8>γ4>γ13>γ10>γ12>γ5>γ7>γ11>γ1>γ9 (Fig. 3-16D).

Next, we examined translocation properties of endogenous Gβγ pool in HeLa and

RAW cells by capturing blue opsin activation induced YFP-Gβ1 translocation (Fig. 3-17A,

D). Since Gβ translocates with endogenous Gγ, Tt1/2 of Gβ is an indicator of translocation properties of endogenous Gγ population, thus we termed it average Tt1/2 (avg-Tt1/2).

Interestingly, avg-Tt1/2 value of RAW cells (221±5 s) was greater than avg-Tt1/2 of HeLa cells (93±2 s) (Fig. 3-17A, B, D). This result suggests that, compared to HeLa cells, RAW cells express more HiAf-Gγ types. This data is in agreement with real-time PCR data of

Gγ mRNA (Fig. 3-4A). To ensure that the type of Gβ does not influence endogenous Gγ translocation, similar experiments were performed both in HeLa and RAW cells, however expressing YFP-Gβ2 (Fig. 3-17A, B, D). The observed Tt1/2 of Gβ1 and Gβ2 are comparable, suggesting that the type of Gβ does not alter Tt1/2 of Gγ. The small Tt1/2 of Gβ observed in Gγ9 expressing cells (Tt1/2 of Gβ1=7±2 s and Gβ2=6±1 s) confirms that Gβ represents translocation properties of endogenous or introduced Gγ (Fig. 3-17 A, B, D).

Without expressing any Gγ, PIP3 production in WT HeLa and WT RAW cells

(|퐸퐹|푒푥푝or ∆[PIP3]) was calculated, using the change in Akt-PH sensor intensity on the

A. β1-YFP β2-YFP

9 9 γ -

mCh Before After Before After

B. HeLa RAW

YFP YFP

- β1

YFP - β2 Before After Before After

C. D. Translocation properties of Gβ

Fig. 3-17: Testing Tt1/2 of Gγ as a predictor of a cell’s ability to control Gβγ effectors. A. HeLa cells expressing blue opsin-mTurquoise, mCherry- Gγ9, either YFP-Gβ1 or YFP-Gβ2 respectively, supplemented with 50 µM 11-cis-retinal. On blue opsin activation, both Gβ1 and Gβ2 exhibited Tt1/2 closer to that of Gγ9, further confirming that the translocation properties of Gβ represent the prominent Gγ subtype expressed in the cell. B. HeLa and RAW cells expressing blue opsin-mTurquoise and either YFP-Gβ1 or YFP-Gβ2, were supplemented with 50 µM 11-cis- retinal. Cell was imaged for YFP and BO was activated with 445 nm light every 3 s. Gβ translocation exhibited the average translocation properties of the entire pool of endogenous Gγ. Gβ type does not influence translocation properties of endogenous Gγ in HeLa cells. The plot shows that Tt1/2 of Gβ1 and Gβ2 translocation was closer to the Tt1/2 of the most abundant Gγ of each cell type (error bars: SEM, n=10; scale bar: 5 µm). 58

Fig. 3-17: Testing Tt1/2 of Gγ as a predictor of a cell’s ability to control Gβγ. C. Blue opsin activation induced experimental IEFI (PIP3 response) measured in WT HeLa and RAW cells expressing blue opsin and the PIP3 sensor. D. Translocation properties of Gβ.

PM (Fig. 3-17C). Tt1/2 values of Gβ translocation in HeLa and RAW cells were considered as the averaged Tt1/2 of their endogenous Gγ (Fig. 3-17A, B, D). The values of avg-Tt1/2 obtained using Gβ1 translocation above (were then extrapolated from the effector plot of

HeLa cells (Fig. 3-16D) to obtain predicted effector activities (|EF|calc). Ratio of experimental and calculated effector activities (|퐸퐹|푒푥푝:|퐸퐹|푐푎푙푐) for HeLa and RAW cells were respectively 0.82 and 0.99. The |퐸퐹|푒푥푝: |퐸퐹|푐푎푙푐 values were closer to 1 for both

HeLa and RAW cells indicating that, the avg-Tt1/2 of endogenous Gγ pool is a strong predictor for a cell’s Gβγ effector-activation ability.

3.2.9. Contribution of Gβγ PM-retention on its effector activation ability:

MoAf-Gγ expressing cells maintain considerably high Gβγ concentration on the

PM after GPCR activation. It was unclear why MoAf-Gγ expression does not promote robust PIP3 production, compared to Gγ3 expressing cells. To comprehend this, a model was proposed where Gβγ on the PM stays in a transient active conformation (Gβγ*) that can activate effectors. The lifetime (τ) of this active conformation is assumed to be Gγ type or more specifically the PM-affinity of Gγ type dependent. When GPCRs are activated, because of the prominent translocation of LoAf-Gβγ, PM should only be enriched with

HiAf and MoAf-Gβγ types. Although HiAf and MoAf Gβγ only exhibit a marginal

59 difference in translocation extent, they significantly differ in Tt1/2 and their ability to activate effectors (Fig. 3-5B, C, and Table 3-1).

A. B.

Fig. 3-18: Predicted model of signal transduction from GPCRs to the cell interior in multi-Gγ systems. A. The reactions representing the proposed mechanism of GPCR-G protein activation used in the model. B. Proposed 푛∗ Gβγ fluctuation between active-inactive conformations (퐺훽훾푃푀 ⇆ 푛 퐺훽훾푃푀ሻ, which is assumed in the optimized model.

Using Gγ12 as a model MoAf Gγ, we examined if the observed lack of translocation in Gγ12 is controlled by factors independent of its CT. We substituted CT of Gγ12 with

CT of Gγ3 and Gγ9 respectively (Fig. 3-19). Compared to the moderate translocation observed in Gγ12 (Tt1/2 = ~80 s), Gγ12-γ9CT mutant showed a fast translocation with Tt1/2

= ~8 s, resembling translocation properties of Gγ9. As expected, Gγ12-γ3CT translocated slower than Gγ12 (Tt1/2 = ~232 s) (Fig. 3-19C). Since the CT of Gγ does not interact with the receptor, the fast translocating mutants of MoAf and HiAf Gγ suggest that, their heterotrimers are likely to be equally activated by the GPCR, as seen for heterotrimers with

LoAf Gγ. These observations indicate that, though MoAf-Gβγ are liberated from heterotrimer and present on the PM, a fraction of them are not conformationally active.

These findings are consistent with recent reports which suggest that K-Ras possesses orientation dependent effector binding (102).

A. Gγ3 Gγ12 Gγ12 with Gγ3 CT Gγ12 with Gγ9 CT Gγ9

Basal

50 s s 50

600 s

B. C.

Fig. 3-19: Gβγ translocation with Gγ12 mutants. A. HeLa cells transfected with blue opsin-mTurquoise and YFP tagged Gγ12 mutants were monitored for Gγ translocation with blue opsin activation with 50 µM 11-cis-retinal addition followed by 0.1% 445 nm blue light exposure (error bars: SEM, n=10; scale bar: 5 µm). B. Plot shows the significant difference in Gγ translocation of Gγ12 mutants compared to wild type Gγ12, Gγ9 and Gγ3. C. Translocation properties of Gγ12 mutants.

3.3. DISCUSSION:

Considering diverse and unique tissue and cell type specific Gγ type distribution patterns, the Gγ identity specific regulation of Gβγ signaling can have a broader impact on the current understanding of GPCR-G protein signal transduction. If Gβγ were to be a unitary signaling entity, cells would have intense Gβγ signaling on all occasions of GPCR activation, which can be deleterious. For instance, RAW cells have a Gγ profile with HiAf-

Gγ that supports PI3K activation and PIP3 production. However, for a usually immobile cell type like HeLa, this level of PIP3 production may not serve a purpose and therefore there is no need for HiAf-Gγ expression. Validating this, Gγ3 expression allowed HeLa cells to produce PIP3. The fundamental difference identified between introduced Gγ3 over endogenous Gγ types in HeLa cells was the ability of Gγ3 to make Gβγ more available at the PM, where PIP3 production takes place. To catalyze PIP2 to PIP3, Gβγ recruits and activates PI3K subunits to the PM (103). Out of the 12 Gγ types, only Gγ3 and Gγ2 promoted PI3K activation, likely due to their weak translocation properties and associated increased Gβγ abundance on the PM. Gγ dependent differential PIP3 generation in HeLa cells hints at a plausible mechanism of how Gβγ effectors are recruited to the PM and activated by PM bound HiAf-Gβγ. We have recently shown that Gβγ controlled PLCβ activation induced Ca2+ mobilization is mandatory for trailing edge retraction during RAW cell migration (87). Since similar to PI3Kγ, PLCβ1 and PLCβ2 are also cytosolic, we could explain this response as the Gβγ induced differential PLCβ activation induced differential calcium responses (104, 105). Our data suggest that, PM targeting and-or activation of these are likely to be governed by the PM-affinity of Gβγ. The extent of effector responses suggests that, the stronger the PM-affinity of Gβγ, the greater its potential to control

62 signaling. The distinct rates of Gβγ translocation away from the PM reflect differential abilities of each of the 12 Gγ types to exhibit varying abilities to stay bound to the PM. The slower translocators expend longer time on the PM, therefore possessing a high PM-affinity and concentration on the PM compared to faster translocating Gγs. The kinetics of Gβγ translocation reveals that Tt1/2 is a measure of the PM-affinity of Gβγ.

Gγ subunits interact with the PM through the prenyl group. The type of prenylation is decided by the CaaX motif sequence of Gγ. The prenylation with 20-carbon geranylgeranyl lipid provides a higher PM-affinity to Gβγ versus the 15-carbon farnesyl lipid attachment. Except for farnesylated Gγ9, Gγ1 and Gγ11, all other Gγ types are geranylgeranylated. However, only Gγ3 and Gγ2 supported RAW cell migration, suggesting factors additional to the type of prenylation control the PM affinity of Gγ.

Interestingly, pre-CaaX regions of Gγ3 and Gγ2 are composed of ~80% positively charged and hydrophobic residues, as opposed to ~50% in geranylgeranylated-Gγ subunits.

Extensive mutagenesis to the pre-CaaX region of Gγ suggested that this 5 or 6 -residue region modulates Gβγ-PM interactions, in which positively charged and hydrophobic amino acids strengthen the PM-affinity. Previously reported translocation data of Gγ mutants with altered pre-CaaX residues further validate the role of this motif in controlling the PM-affinity (55). The complete loss of PM localization observed in Gγ9 upon cysteine removal from CaaX motif indicates that, pre-CaaX region only serves as a strong modulator of PM-affinity, but prenylation is essential for primary PM anchoring of Gβγ.

By modulating properties of their pre-CaaX motifs, geranylgeranylated-Gγ subunits managed to possess a discrete series of PM affinities.

Heterotrimers with specific Gγ types have been shown to possess higher affinities towards certain GPCRs (89, 106). However, an exchange of CT of slow translocating Gγ3 and moderate translocating Gγ12 with CT of Gγ9 resulted in fast translocating mutants, comparable to Gγ9. This can suggest that either (a) heterotrimer activation process is controlled by CT of Gγ through modulating Gαβγ-GPCR interactions, or (b) the PM- affinity of generated Gβγ is dependent on CT of Gγ subunit. Regardless, CT of Gγ should hold a crucial control over Gβγ function, although our data strongly support (b). We anticipate that, among the available Gβγ pool, LoAf and MoAf Gβγ types exist primarily to support GαGTP generation, while HiAf-Gβγ subunits activate Gβγ effectors. We also anticipated that the PM bound Gβγ composed of HiAf-Gγ types stay for a fractionally longer time in the active conformation compared to their LoAf and MoAf associates. Lack of migration ability in Gγ3-knocked down RAW cells strongly support this notion.

Nevertheless, we are aware that, in addition to Gγ diversity, there are converging and diverging pathways and signaling components including integrins, secretory proteins (i.e. matrix metalloproteinases), that can influence the migration potential of a cell (107-109).

Therefore, differences in cell migration potentials among cell types with diverse origins can be associated with these possible inherent variables. While we are in concert with these reports, cells cannot migrate at all using the Gi pathway, if they lack appropriate Gγ-types with proper PM-affinity. Supporting this finding, even non-migratory HeLa cells attempted to undergo blue opsin induced directional migration upon HiAf-Gγ3 expression.

Avg-Tt1/2 of endogenous Gγ measured using Gβ translocation accurately predicted the ability of native Gβγ to control its effectors. Predicted effector activity using this method was similar to PIP3 production observed for both RAW and HeLa cells. These

64 observations suggest that Tt1/2 and PM-affinity of a Gγ are strong indicators of the ability of Gβγ to activate effectors. Therefore, our observations collectively indicate that Gγ subunit diversity in a cell is a crucial in determining whether the cell has the ability to activate Gβγ effectors sufficiently to orchestrate intended behaviors including migration.

3.4. CONCLUSION:

In summary, this study demonstrates that distinct translocation abilities of the 12

Gγ types provide Gβγ a diverse range of PM and effector interaction abilities. Since, most

Gβγ-effector activities occur at the PM, data confirms that, the PM affinities of Gγ types expressed in a cell are deterministic to the potency of Gβγ effector as well as downstream signaling activation. Although we only checked Gγ identity dependent control of PI3Kγ and PLCβ, and their regulation of cell migration, it is likely that a plethora of Gβγ mediated functions are similarly regulated. Since GPCR-G protein signaling is universally conserved and Gβγ signaling pathways are major drug targets, mechanisms we describe here can have a wide influence not only on cell migration but also in many areas of signaling.

3.5. MATERIALS AND METHODS:

3.5.1 Reagents:

The reagents; Gallein (TCI AMERICA), Fluo-4 AM (Molecular probes, Eugene,

Oregon), Wortmannin (Cayman Chemical, Ann Arbor, MI), c5a (Eurogentec), U50488 hydrochloride (Tocris) were initially dissolved in DMSO and then diluted in HBSS (Gibco

65 laboratories) before adding to cells. 11-cis retinal (National Eye Institute) was initially resuspended in absolute ethanol and 2 µl aliquots (50 µM) were further diluted (2 µl for each aliquot) with absolute ethanol before introducing (2 µl) to cells in dark. SDF-1α

(PeproTech) was reconstituted in DI water to a concentration 100 µg/mL and further diluted with a buffer containing 0.1% BSA before adding to cells.

3.5.2. DNA constructs and cell lines:

Engineering of DNA constructs used; blue opsin-mCherry, blue opsin-mTurquoise,

Akt-PH-mCherry and YFP tagged Gγ1-Gγ13, have been described previously (99, 110,

111). YFP-β1 and 2, kappa-opioid receptor, PI3K-CA-CFP and mCh-GPI were kind gifts from Professor N. Gautam’s lab, Washington University, St. Louis, MO. Gγ3, Gγ9 and

Gγ12 mutants were generated using Gibson assembly (NEB) (112). Parent constructs; mCherry-Gγ3, mCherry-Gγ9, and YFP- Gγ12 were PCR amplified with overhangs containing expected nucleotide mutations. DpnI (NEB) digestion was performed on the

PCR product to remove the parent construct. DpnI digested PCR product was then mixed with the Gibson assembly master mix (NEB) and incubated at 50 0C for 45 min, which was followed by transformation of competent cells and plating on Ampicillin LB agar plates.

All the constructs used in this study possess the Ampicillin resistant pcDNA 3.1 vector backbone. Cell lines (HeLa, RAW 264.7, PC12, and HEK cells) were originally purchased from the American Tissue Culture Collections (ATCC) and authenticated using a commercial kit to amplify 9 unique STR loci.

3.5.3. Cell culture and Transfections:

RAW 264.7 cells used in migration and PIP3 generation experiments were cultured in RPMI 1640 (10-041-CV; Corning, Manassas, VA) with 10% dialyzed fetal bovine serum (DFBS; from Atlanta Biologicals) and 1% Penicillin−Streptomycin (PS) in 60 mm tissue culture dishes. HeLa cells were maintained in minimum essential medium (MEM; from CellGro) supplemented with 10% DFBS and 1% PS. Around 80% cell confluency, the growth medium was aspirated, 2 mL versene (EDTA) (CellGro) was added, incubated

0 for 3 minutes at 37 C, 5% CO2 incubator, and then cells were lifted and suspended in versene. The cell suspension was centrifuged at 1000 g for 3 minutes, versine (EDTA) was aspirated, and the cell pellet was resuspended in their normal growth medium (RPMI/

DFBS/ PS for RAW, and MEM/ DFBS/ PS for HeLa) at a cell density of 1 × 106 /mL. For imaging experiments, cells were seeded on 35 mm glass-bottomed dishes (8 × 104 cells on each) with 15 mm inner diameter, prepared using #1 German cover glasses. Before cell seeding, dishes were washed with 2 N NaOH for 20 min, ethanol washed, and sterilized for one hour using UV irradiation. A day following cell seeding, cells were transfected with appropriate DNA combinations using the transfection reagent PolyJet (SignaGen),

0 according to the manufacturer’s protocol and then incubated in a 37 C, 5% CO2 incubator.

Cells were imaged after 16 hours of the transfection.

3.5.4. Knockdown of Gγ3 in RAW 264.7 cells:

Five shRNAs (TRCN0000036794-98; Sigma-Aldrich) were screened in RAW cells by co-expressing with GFP- Gγ3. Cells were screened for GFP expression and the shRNA construct that induced the highest reduction in GFP-Gγ3 expression was selected as the most effective shRNA. The identified TRCN0000036795 shRNA was employed to

67 knockdown Gγ3 in the subsequent experiments. A scrambled shRNA was used as the control.

Functional shRNA sequence:

CCGGGCTTAAGATTGAAGCCAGCTTCTCGAGAAGCTGGCTTCAATCTTAAGCT

TTTTG

3.5.5. Live cell imaging to monitor Gβγ translocation, PIP3 generation, and optogenetic control of cell migration:

Imaging system: Spinning-disk XD confocal TIRF (total internal reflection) imaging system composed of a Nikon Ti-R/B inverted microscope, a Yokogawa CSU-X1 spinning disk unit (5000 rpm), an Andor FRAP-PA (fluorescence recovery after photobleaching and photo-activation) module, a laser combiner with 40−100 mW 445, 488, 515, and 594 nm solid-state lasers and iXon ULTRA 897BV back-illuminated deep-cooled

EMCCD camera. Live cell imaging was performed using a 60X, 1.4 NA (numerical aperture) oil objective. In cell migration and PIP3 generation experiments, mCherry tagged receptor blue opsin and the PIP3 sensor Akt-PH were imaged using 594 nm excitation−630 nm emission. To activate blue opsin, 50 µM 11-cis retinal (National Eye Institute) was added and incubated 3-5 min in dark. After incubation, the fluorescent sensor in cells was imaged to capture basal signaling in cell migration and PIP3 generation experiments (i.e.,

Akt-PH-mCherry in PIP3 experiments and blue opsin-mCherry in cell migration experiments), and then receptor blue opsin was activated by shinning 445 nm blue light at

0.1% transmittance and imaging was continued for 20 min. To examine the Gβγ translocation, YFP and GFP fluorescent tags on Gγ subunits were imaged for 10 min using

515 nm excitation 527 nm emission or 488 nm excitation 515 nm emission, respectively.

Regular culture media or HBSS supplemented with 1g/mL glucose preincubated in a 37

0 C, 5% CO2 incubator for 30 minutes were used as the imaging medium. During imaging, cells were maintained at 37 0C. To prevent focal plane drifts, Nikon Perfect Focus System

(PFS) was engaged.

3.5.6. Cytosolic Ca2+ measurements:

For intracellular Ca2+ measurements, RAW cells seeded on glass-bottom dishes and maintained in a 37°C with 5% CO2 were transfected with a Gγ subtype on the following day of cell seeding. After 12 to 16 h of transfection, cells were washed twice with Ca2+ containing HBSS (pH 7.2) and incubated for 30 min at room temperature with the fluorescent Ca2+ indicator, fluo-4 AM in the dark. After incubation, cells were washed twice with HBSS and 500 µl of HBSS was then used as imaging medium. The fluorescence intensity of flow-4 AM was continuously imaged at 1 s intervals using 488 nm excitation-

515 nm emission to capture signal-before activation for 50 s. Endogenous c5aRs in RAW cells were activated with 10 µM c5a. Observed fluo-4 AM fluorescence increase due to

Ca2+ release was base line normalized.

3.5.7. Real-time PCR, transcriptome, and RNAseq data analysis:

In order to obtain the Gγ profile of WT HeLa and WT RAW 264.7 cells, RNA was extracted from cells grown in 100 mm tissue culture dishes after reaching 90%-100% cell confluency. RNA extraction was performed using the GeneJet RNA purification kit following their given protocol. Extracted RNA was used as the template for cDNA synthesis with Radient cDNA synthesis kit. cDNA product was quantified using the

69 nanodrop and used for real-time PCR (BioRad CFX96Real-Time qPCR system) in 96 well plates to obtain the Gγ profile. Radient Green Lo-ROX qPCR kit (Alkali Scientific) was used in real-time PCR experiments and β actin gene was used as the house keeping gene.

To screen the Gγ profile alteration with Gγ3 and Gγ9 overexpression, HeLa cells were seeded in 100 mm tissue culture dishes, transfected with GFP-Gγ3 and GFP-Gγ9 respectively at 70%-80% cell confluency, and RNA was extracted after confirming greater than 70% transfection efficiency by observing under the microscope. This was followed by cDNA preparation and real-time PCR.

3.5.8. Statistics and reproducibility:

Results of all quantitative assays (Gβγ translocation, cell migration, and PIP3 generation) are expressed as standard error of mean (SEM) from n numbers of cells

(indicated in the figure legends) from multiple independent experiments. Statistical analysis of cell migration data of WT and mutant Gγ subtypes was performed using two- tailed unpaired t-test. P value < 0.05 was considered as statistically significant.

Chapter 4

Short Pre-CaaX Regions in Gγ are Uniquely Evolved

Switches of Heterotrimeric G Protein Signaling

4.1. INTRODUCTION:

G protein heterotrimers (Gαβγ) interact with the inner leaflet of the plasma membrane (PM) primarily through their covalent lipid modifications (fatty acylations), and we showed that these modifications provide an additional layer of G protein activity regulation (113, 114). G protein α (Gα) subunits are modified at their N terminus (NT) by covalent attachment of 14-carbon (14-C) myristate or 16-C palmitate fatty acid groups or both, while G protein γ (Gγ) subunits are prenylated at their C terminus (CT) (113, 115).

Prenylation of the twelve Gγ types with either a 20-C isoprenoid geranylgeranyl or a 15-C isoprenoid farnesyl group attachment involves a stable thioether bond formation at the cys residue CaaX motif in the CT of Gγ (113). Although only two types of prenyl attachments exist, we demonstrated a Gγ-type dependent unique and multiple PM affinities of Gβγ that differentially regulate Gβγ signaling in a Gγ-type specific manner (39, 116). For instance,

Gγ3, which has the highest PM-affinity, exhibited the greatest effector activation, while

Gγ9 has the lowest PM-affinity and showed almost no effector activation (39, 116). The

PM-affinity is measured as the inverse of the translocation ability of Gβγ observed after

GPCR activation. Interestingly, several Gγ subtypes with moderate PM-affinities, in which a considerable portion of Gβγ is left on the PM even after GPCR activation, exhibit an unexpectedly low effector activation.

In addition to the prenyl lipid modification on the C-terminal CaaX motif, secondary interactions between amino acid residues in the Gγ CT and the PM also have been shown, which can be explained by the composition and chemical properties of the

PM (117, 118). The PM possesses a characteristic phospholipid bilayer structure with embedded proteins, which forms a fluidic mosaic (119, 120). It contains asymmetrically distributed four major types of phospholipids; phosphatidylcholine, phosphatidylethanolamine, phosphatidylserine, and sphingomyelin, each composed of two acyl lipid anchors linked to a polar phosphate head group through a glycerol molecule

(121). The polar negatively charged and hydrophilic head faces the cytosol and due to the hydrophobicity, the acyl groups of phospholipids are oriented into the core of the bilayer structure. Out of the four-phospholipid types, phosphatidylinositol, phosphatidylethanolamine, and phosphatidylserine are abundant in the inner leaflet of the

PM, defining the properties of the PM (121-123). Since the polar head groups of both phosphatidylserine and phosphatidylinositol are negatively charged, their predominance provides a net negative charge to the cytosolic inner leaflet of the PM (124, 125). Overall, the PM is considered as negatively charged and hydrophobic and changes in PM lipid composition has been shown to alter their association with G proteins (126, 127).

We previously showed that Gγ3 and Gγ2 possess the highest affinity to the PM and cells expressing these subtypes show the best Gβγ effector activities (116). Both of these

Gγ subtypes possess hydrophobic phe as well as positively charged lys and arg residues at

72 their CT; especially in their pre-CaaX region. However, Gγ that exhibited low PM-affinity and low effector activation ability (i.e., Gγ9, Gγ1) possess neutral gly as well as negatively charged glu (39, 116). Thus, we hypothesized that, these differential effector activation abilities of Gβγ are governed by chemical properties of the PM and amino acid residues of the CT regions of Gγ types. Additionally, three-dimensional structure of a protein is crucial for its biological function and is determined by both the primary sequence of the polypeptide, as well as its surrounding chemical environment (128, 129). PM interacting proteins are either embedded or interacting with the PM, and the structures and the function of these proteins are heavily influenced by the properties of the PM. The fluid-mosaic membrane structure permits several modes of movements to the proteins interacting with

PM bilayer (119, 130). The rotational movements allow proteins to sample its immediate neighborhood and interact with effectors. Lateral diffusion allows proteins to migrate from one PM-microdomain to another and reach distant effectors (131, 132). This lateral mobility of proteins in biological membranes is affected by the degree of crowding, membrane domains (i.e., lipid rafts), and interactions with cytoskeletal components (133-

135). The crystal structure of GRK2-Gβγ shows rotation of Gβγ by ~ 900 and while moving over 100 angstroms, (i) allowing Gβγ to interact with the binding site on GRK2 and (ii) exposing both the receptor and GαqGTP to interact with GRK2 (136-138). Therefore, lateral and rotational movements of Gβγ on the PM are likely to affect the activation of other Gβγ effectors as well. Since the CT of Gγ interacts with the PM, differential sequence properties of CT domains among Gγ subtypes can result in differential lateral as well as rotational movements of Gβγ on the PM.

Similar to Gγ, many other G proteins including Ras and Ras-like proteins are prenylated and possess a CaaX and pre-CaaX motifs at their CT (139, 140). These proteins play key roles in many cellular functions including regulation of protein trafficking, cell proliferation, differentiation, and survival (31, 140, 141). However, compared to the 5-6 residue short peptide in Gγ, the Ras proteins possess a several-fold longer and primarily polybasic pre-CaaX region. We question if this significant difference in the pre-CaaX region is related to the unique behavior of Gβγ as a signal transducer when compared to

Ras family proteins. Especially, we quest how crucial the short pre-CaaX motif in Gγ for

Gβγ as well as heterotrimer signaling. Since only Gγ2, γ3 and γ4, however neither rest of the Gγ types nor other prenylated G proteins, possess the unique two phe residues before the prenylated-cys, we ask what role this unique phe-duo plays in Gβγ signaling? We examined these questions below to shed the light on the molecular level interactions of Gγ with the PM. Our results reveal a unique evolution of the pre-CaaX region of Gγ, initially to facilitate heterotrimer formation, to interact with the activated GPCRs, and subsequently to act as an ‘‘on-off’’ switch for Gβγ signaling.

4.2. RESULTS:

4.2.1. Decoding selection criteria of residues in Gγ-CT that tunes Gβγ-PM interactions:

Although prenylated Ras superfamily proteins contain a lipid modification for membrane anchoring, they all contain a 15-20 residue poly basic CT region to reinforce

PM anchoring. How does Gγ accomplish anchoring only with a 5-6 residue CT region?

A. B. Conserved phe pre-CaaX region conserved phe GNG1 --NP F KELKGG CVIS GNG2 --NP F REKKFF CAIL GNG3 --NP F REKKFF CALL prenyl GNG4 --NP F REKKFF CTIL anchor GNG5 --NP F RPQKV- CSFL GNG7 --NP F KDKKP- CIIL Gγ GNG8 --NP F RDKRLF CVLL pre- GNG9 --NP F KE-KGG CLIS CaaX Gβ GNG10 --NP F REPRS- CALL region GNG11 --NP F KE-KGS CVIS GNG12 --NP F KDKKT- CIIL GNG13 --NP W VEKGK- CTIL

C. PM PM PM

Fig. 4-1: Molecular rationale for Gβγ PM-affinity control by pre-prenylation residues of Gγ. A. Sequence alignment of the CT regions of the twelve Gγ subtypes. Conserved phe→ blue box, pre-CaaX region→green box, and terminal cys→brown, which undergoes prenylation and carboxy methylation. B. Gβγ crystal structure (PDB ID: 2BCJ) modified to show its interaction with the PM. The green box shows the pre-CaaX region, blue residue represents the conserved phe residue, which is also the last contact point of Gγ with Gβ. The prenyl lipid anchor that interacts with the PM is shown in black. C. Hypothesized interactions between CT of Gγ9 and Gγ3 with the PM. Polar- charged and hydrophobic groups in pre-CaaX residues of Gγ respectively interacts with polar head groups and hydrophobic core of the PM. Negatively- charged glu (red) residues likely to modulate Gγ – PM interactions. This model provides molecular reasoning for PM-affinity differences among Gγ-members including Gγ9 and Gγ3.

Except Gγ13, all Gγ subtypes possess a conserved phe residue (i.e. phe65 in Gγ3 and phe60 in Gγ9), which functions as the last contact point of Gγ with Gβ (Fig. 4-1A-blue box).

Using site directed mutagenesis, we perturbed this phe and the subsequent 5-6 residues in the pre-prenylation (pre-CaaX) region (Fig. 4-1A-green box). The pre-CaaX region is in the close proximity to the PM, likely interacting with it (Fig. 4-1 B).

Out of the twelve Gγ subunits, Gγ9 imparts the highest translocation rate for Gβγ from the PM to internal membranes (IMs) upon GPCR activation (39, 116). Thus, Gγ9 exhibits the lowest Gβγ concentration at the PM, and therefore elicits the lowermost Gβγ- effector signaling (39, 116). On the contrary, Gγ3 indulges the lowest translocation rate for

Gβγ and thus the highest Gβγ effector activity. Although geranylgeranyl modification in

G3 is a stronger membrane anchor than farnesyl in Gγ9, we have previously shown evidences for pre-CaaX region controlling the PM-affinity and Gβγ-effector activity (116).

Pre-CaaX sequence properties of high PM-affinity Gγ subunits showed the presence of primarily positively charged and hydrophobic amino acids (Fig 4-1A-underlined).

Properties of their side-chains establish electrostatic interactions with negatively charged phospholipid head groups and hydrophobic interactions with the core of the PM, respectively (Fig. 4-1C) (116). Therefore, to examine how just two lipid anchors results in a discrete series of PM-affinities as well as Gβγ effector activations, we systematically perturbed pre-CaaX residues in Gγ3 and Gγ9, since they provide two-extreme PM-affinity characteristics for Gβγ. Our strategy was to examine whether we could generate Gγ3-like mutants from Gγ9 and vice-versa, by only altering residues in the pre-CaaX region.

4.2.2. Prenylated-cys adjacent phe residues in the CT of Gγ3 are essential for Gβγ effector activation:

Gγ sequences display two distinct groups of residues in their pre-CaaX region; non- hydrophobic as in Gγ1, γ5, γ7, γ9, γ10, γ11, γ12, and γ13, and hydrophobic as in Gγ2, γ3,

γ4, and γ8. In particular, we show that, among all prenylated G proteins, including heterotrimeric as well as Ras family, only Gγ3, γ2, and γ4 have a unique two hydrophobic residues next to the prenylated-cys (phe-duo) (Fig. 4-2). Since Gγ3, γ2, and γ4 provide the highest Gβγ signaling activity, we predicted that the phe-duo act as additional PM anchors, strengthening Gγ-PM interactions via the methyl phenyl side group on phe (Fig. 4-1C-left, black circle). We also anticipated that, this additional anchoring requires close proximity

Fig. 4-2. Pre-CaaX regions of prenylated G proteins. CT residue alignment of several classes of prenylated-proteins including heterotrimeric and Ras family G proteins. Note that only Gγ3, γ2, and 4 have the unique two phe residues next to the prenylated-cys (red box).

to the PM bilayer and is provided by the

adjacent prenyl anchor. In contrast, Gγ9,

which shows the lowest Gβγ activity,

possesses two gly residues next to the prenylated-cys, and is expected to loosely interact with the PM (Fig. 4-1C-right).

To explore these hypotheses, we perturbed the two residues before prenylated-cys in Gγ3 and Gγ9 respectively. When the phe-duo in Gγ3 were replaced with two gly residues

(Gγ3-phe-phe→gly-gly: NPFREKKGGCALL), as in Gγ9, the resulted mutant expressing

77 cells exhibited a ~70% increase in Gβγ translocation t1/2 (t1/2= 92±8 s) compared to WT

Gγ3 expressing cells (t1/2= 290±7 s) (Fig. 4-3A-yellow arrow, plot and Table B-1). Since this mutant is still expected to possess geranylgeranyl lipid anchor, the observed significant change in translocation t1/2 indicated that, irrespective of the type of prenylation in the pre-

CaaX, prenylated-cys adjacent hydrophobic residues are crucial for PM interactions of

Gβγ. Since G3 expressing cells exhibit a robust PIP3 production upon Gi/o-coupled

GPCR activation (116), we examined Gi-pathway activation-induced PIP3 production in this G3-phe-phe→gly-gly mutant expressing HeLa cells. Compared to control cells expressing G3, Gγ3-phe-phe→gly-gly mutant (NPFREKKGGCALL) expressing cells showed a ~50% attenuated rate of PIP3 generation (0.00042 s-1) upon activation of endogenous α2-adrenergic receptors (α2-ARs) (Fig. 4-3B, plot and Table B-1).

To further validate the crucial nature of this phe-duo in regulating the PM-affinity as well as the efficacy of effector activation of Gβγ, we replaced two adjacent gly residues next to prenylated-cys in Gγ9 with two phe residues (Gγ9-gly-gly→phe-phe:

NPFKEKFFCLIS). This mutant exhibited over ~80% slower translocation (t1/2= 169±7 s) compared WT Gγ9 (t1/2= 37±4 s) (Fig. 4-3A, Table B-1), indicating that phe residues at this location significantly enhance the PM-affinity of Gγ. Although Gγ9 expression suppress Gβγ induced PIP3 production in HeLa cells, this Gγ9-gly-gly→phe-phe mutant expressing cells exhibited a ~ 50% enhanced PIP3 production rate (0.00038 s-1) upon activation of α2-ARs (Fig. 4-3B, Table B-1).

A. Gβγ9 translocation

Gγ3 WT Gγ3-FF→GG Gγ9 WT Gγ9-GG→FF

Basal Basal NE

After NE NE After

B. PIP3 generation

Gγ3 WT Gγ3-FF→GG Gγ9 WT Gγ9-GG→FF

Basal Basal NE

After NE NE After

Fig. 4-3: Role of prenylated-cys adjacent phe residues in Gγ3 CT in Gβγ effector activation A. HeLa cells transfected with GFP tagged Gγ9, Gγ3, or their mutants. Translocation of Gγ from the PM to IMs (yellow arrows) was captured by imaging cells for GFP (488 nm-ex, 515 nm-em) at 1Hz for 10 mins, before and after endogenous α2-ARs activation with 100 μM norepinephrine. Plot shows GFP fluorescence (n= Gγ3: 13, Gγ3-FF→GG: 18, Gγ9: 14, Gγ9-GG→FF: 16 cells from more than 3 independent experiments) in IM regions over time. B. Time- lapse images of HeLa cells transfected with Gγ3, Gγ9, or their mutants, along with the PIP3 sensor; Akt-PH-mCherry before and after activation of α2-ARs with 100 μM NE. PIP3 production was captured by imaging mCherry (594 nm- ex, 630 nm-em) for 10 mins at 1 Hz. Note distinct levels of PIP3 production with Gγ3 and Gγ9, and the alterations to these WT responses in mutant-Gγ expressing cells. Plot shows mCherry fluorescence (n= Gγ3: 12, Gγ3-FF→GG: 12, Gγ9: 12, Gγ9-GG→FF: 14 cells from more than 3 independent experiments) on the PM over time in cells (scale: 5 μm, error bars: SEM).

4.2.3. Location of the phe-duo in Gγ3 pre-CaaX motif is crucial for the PM-affinity of

G:

Gγ3 and Gγ9 mutants above (Fig. 4-3) established that the phe-duo significantly enhance the PM-affinity as well as signaling activation of Gβγ. Out of twelve Gγ subtypes,

Gγ3, Gγ2, and Gγ4 possess this conserved phe-duo and we have shown that only these Gγ types expression allows Gβγ effectors; PI3K and PLCβ, activation upon Gi-pathway activation (116). Thus, we examined whether this conserved phe-duo reinforce as well as stabilize the PM anchoring of Gβγ as follows.

A Gγ3 mutant was generated in which this conserved phe-duo was shifted to the beginning of the pre-CaaX region (Gγ3-phe-phe shifted: NPFFFREKKCALL) (Fig. 4-4A,

D). Surprisingly, G3-phe-phe shifted mutant expressing cells exhibited a ~ 75% increase in Gβγ translocation (t1/2= 68±5 s), compared to WT G3 cells (t1/2= 290±7 s) upon activation of α2-AR activation in HeLa cells. This mutant expressing cells also exhibited a

50% reduction in the associated PIP3 generation rate (0.00032 s-1) compared to WT G3- cells (0.00069 s-1) (Fig. 4-4E, H), suggesting that the close proximity of this phe-duo to the prenyl-cys is essential for the activity of Gβγ.

To examine if the remaining activity in the Gγ3-phe-phe shifted mutant is due to the poly basicity of the pre-CaaX region, we mutated two lys residues just before the CaaX motif in Gγ3-phe-phe shifted mutant (NPFFFREKKCALL) to two gly residues (Gγ3-lys- lys→gly-gly: NPFFFREGGCALL) (Fig. 4-4B,D). Interestingly, G3-NPFFFREGGCALL mutant expressing cells exhibited only a slightly increased Gβγ translocation (t1/2= 47±5 s) than Gγ3-phe-phe shifted mutant: NPFFFREKKCALL. Further, both mutants exhibited a nearly equal and attenuated PIP3 generation rates (compared to Gγ3-cells) upon α2-AR

80 activation (Fig. 4-4F, H). These observations suggested that the hydrophobic character in the pre-CaaX region and its location next to prenyl-cys is crucial and acts as the primary regulator for switching Gβγ signaling on-off.

A. B. C. E. F. Gγ3-FF shifted Gγ3-KK→GG Gγ3-F65→G Gγ3-FF shifted Gγ3-KK→GG Gγ3-F65→G

Basal Basal

After NE After After NE After D. H.

Fig. 4-4: Hydrophobic character in Gγ pre-CaaX motif is crucial for the PM-affinity of G. A-C. HeLa cells transfected with GFP tagged Gγ9, Gγ3, or their pre- CaaX mutants. Gβγ translocation from the PM to IMs (yellow arrows) captured by imaging GFP at 1Hz for 10 mins, before and after α2-ARs activation with 100 μM norepinephrine. D. Plot shows GFP fluorescence (n= Gγ3-FF shifted: 14, Gγ3-KK→GG in FF shifted: 12, Gγ3-F65→G: 18 cells from more than 3 independent experiments) in IM regions over time. E-G. Images of HeLa cells expressing Gγ3, Gγ9, or their mutants, along with Akt-PH-mCherry before and after activation of α2-ARs with NE. PIP3 generation was captured by imaging mCherry for 10 mins at 1 Hz. H. Plot shows mCherry fluorescence on the PM over time (n= Gγ3-FF shifted: 13, Gγ3-KK→GG in FF shifted: 12, Gγ3- F65→G: 12 cells from > 3 independent experiments) (scale: 5 μm, error bars: SEM).

4.2.4. The last Gβ-interacting conserved phe in Gγ is not a crucial determinant of the

PM-affinity and activity of Gβγ:

This conserved phe interacts with a hydrophobic pocket in Gβ (142). Since we established the significance of phe residues on Gγ-PM interactions from experiment above, we next examined how vital its contribution to the PM-affinity and effector activation ability of Gβγ. A mutant, in which this conserved phe65 in Gγ3 was replaced with gly

(Gγ3-phe65→gly: NPGREKKFFCALL), showed an increased translocation rate (t1/2=

142±9 s) compared to the WT G3, (Fig. 4-4C, D, Table B-1). This Gγ3-phe65→gly mutant cells also exhibited a ~20% reduction in the rate of PIP3 production (0.00056 s-1) (Fig. 4-

4G, H). The increased rate of translocation of this mutant can be due the loss of Gβ orientation in the Gβγ dimer.

4.2.5. Positively charged residues in Gγ CT do not significantly contribute to the PM- affinity and signaling of Gβγ:

It was noted that both Gγ3 and Gγ9 have homologous regions with three residues, arg-glu-lys in Gγ3 and lys-glu-lys in Gγ9, respectively. We hypothesized that the positive charges of arg and lys side chains collectively allow Gβγ to transiently interact with the negatively charged polar head groups of PM phospholipids, where negatively charged glu may likely to cause repulsive forces from the PM (Fig. 4-1C). Considering these opposite charge characteristics next to each other; “+ - +”, we anticipated a “pseudo-spring’’ like behavior for Gγ with wiggling movements showing transient interactions - repulsions with and from the PM (Fig. 4-1C). To examine this, mutations were introduced to arg-glu-lys

(in Gγ3) and lys-glu-lys (in Gγ9). However, a Gγ9 mutant generated replacing the above

A. B. C. E. F. G. Gγ9- Gγ3- Gγ9 with Gγ3 Gγ9- Gγ3- Gγ9 with Gγ3

KEK→GGG RE→GGGG pre-CaaX KEK→GGG RE→GGGG pre-CaaX

Basal Basal

After NE After After NE After D. H.

NE NE

Fig. 4-5: Non-significant influence of positively charged residues in Gγ CT for the PM- affinity and signaling of Gβγ. A-C. HeLa cells expressing GFP tagged Gγ9, Gγ3, or their pre-CaaX mutants. Gβγ translocation (yellow arrows) was captured by imaging GFP fluorescence at 1Hz for 10 mins, before and after α2-ARs activation with 100 μM norepinephrine. D. Plot shows GFP fluorescence change in IM regions over time (n= Gγ9-KEK→GGG: 12, Gγ3-RE→GGGG: 15, Gγ9 with Gγ3 pre-CaaX: 16 cells from more than 3 independent experiments). E-G. Images of HeLa cells expressing Gγ3, Gγ9, or their mutants, along with Akt-PH-mCherry before and after activation of α2-ARs with NE. PIP3 generation was captured by imaging mCherry for 10 mins at 1 Hz. H. Plot shows mCherry fluorescence accumulation on the PM over time (n= Gγ9-KEK→GGG: 12, Gγ3-RE→GGGG: 12, Gγ9 with Gγ3 pre-CaaX: 12 cells from > 3 independent experiments) (scale: 5 μm, error bars: SEM).

mentioned lys-glu-lys residues with three gly residues (Gγ9-lys-glu-lys→gly-gly-gly:

NPFGGGGGCLIS) showed translocation (t1/2= 25±3 s) (Fig. 4-5A, D) and PIP3 generation characteristics (rate= 0.00023 s-1) (Fig. 4-5E, H, Table B-1) similar to Gγ9 WT

(rate= 0.00025 s-1).

Since the mutant Gγ9-lys-glu-lys→gly-gly-gly (NPFGGGGGCLIS) also did not show a significant change on the Gβγ activity, another Gγ3 mutant was designed by

83 replacing arg-glu with two gly s and introducing two more extra gly residues (Gγ3-arg glu→gly-gly-gly-gly: NPFGGGGKKFFCALL), to further test if the above proposed

“pseudo-spring” behavior exists. However, this mutant expressing cells only exhibited a

-1 minor reduction in the PM-affinity (t1/2= 204±5 s) and PIP3 generation (0.00067 s ), compared to WT G3 cells (Fig. 4-5B, D, F, H, Table B-1). Overall, the mutants of the proposed “+ - +” region showed that their contribution to the PM-affinity as well as the

Gβγ signaling activation is minor.

Also, a Gγ9 mutant with the entire pre-CaaX region of Gγ3 (Gγ9-with Gγ3 pre-

CaaX: NPFREKKFFCLIS) showed a translocation (t1/2= 233±8 s) which is nearly similar to the WT G3 as well as closer to the Gγ9-gly-gly→phe-phe mutant (t1/2= 169±7 s) (Fig.

4-5C, D, Table B-1). This mutant also exhibited a ~ 70% enhanced rate of PIP3 generation

(0.00072 s-1) compared to WT Gγ9 (0.00025 s-1) (Fig. 4-5G, H, Table B-1). These data collectively indicate that the pre-CaaX region is the primary regulator of Gβγ signaling over the type of prenylation. Data also show that positively charged residues in the pre-

CaaX region only have a minor effect on Gβγ signaling regulation, while the hydrophobic residues play a major role. These data show that, while the pre-CaaX region controls the

PM-affinity of Gβγ and Gβγ-mediated signaling, the phe-duo is essential for Gβγ signaling activation, and regardless of the type of prenylation, its effect prevails.

4.2.6. Terminal cys-attached phe-duo is crucial for activation of key kinases in the chemokine pathway and invasion of cancer cells:

Since Gβγ is reported to promote cancer cell invasion and thus metastasis (Fig. 4-

6), we examined if and how the nature of pre-CaaX region of Gγ affect the activity of Gβγ- regulated kinases as well as subsequent invasion (143, 144). Breast cancer cell line MDA-

MB-231 was used for this study since it is considered as a useful experimental model for metastasis (145, 146). Based on initial experimental results, two Gγ3 and two Gγ9 mutants, which showed significant deviations from the behaviors of the WT Gγ3 and Gγ9 expressing cells, were selected for the study.

Fig. 4-6: Gβγ-driven cell invasion. Signal propagation from Gβγ through kinases to trigger cell-invasion.

4.2.6.1. Screening MDA-MB-231 cells for an endogenous chemokine receptor:

MDA-MB-231 cells were first screened for the endogenous expression of Gi- pathway chemokine GPCRs by examining the ability of a variety of ligands that induce

Gβγ9 translocation (Fig. 4-7). Upon addition of 100 μM norepinephrine, MDA-MB-231 cells exhibited a robust translocation of GFP-Gγ9 indicating the presence of endogenous

α2-ARs at levels that can induce physiological responses. Other ligands tested, including carbachol, SDF-1α, C5a, and sphingosine-1-phosphate (S1P), did not induce Gγ9 translocation in these cells, suggesting the lack of the cognate receptors (Fig. 4-7).

A. B.

Norepinephrine C5a Carbachol S1P SDF-1α

Before

After

Fig. 4-7: Screening for the endogenous receptors in MDA-MB-231 cells. A. MDA-MB- 231 cells expressing GFP-Gγ9 were examined for Gβγ9 translocation after exposing to 100 μM norepinephrine (NE), 10 μM c5a, 10 μM carbachol, 10 μM S1P, and 50 ng/ml SDF-1α addition (red arrow) respectively. Translocation was only observed in cells exposed to NE indicating the presence of endogenous α2- AR in MDA-MB-231 cells. B. Plot shows the quantification of Gβγ9 translocation by examining the fluorescence changes in IMs with each ligand addition (scale: 5 μm, error bars: SEM, n=8).

4.2.6.2. CT of Gγ regulates phosphorylation of kinase; Akt, in cancer cells:

Since the phe-duo turned the PI3K activity in HeLa cells (Fig. 4-3B), we examined how significant the presence of these residues in Gγ to the cellular events controlled by

Gβγ, including activation of kinases and chemokine pathway-induced invasion (Fig. 4-6).

To examine Gi-pathway and Gβγ mediated PI3K-PIP3 pathway upon endogenous α2-AR activation in cancer cells, MDA-MB-231 cells expressing the selected following Gγ3 and

Gγ9 mutants were used: Gγ3-phe-phe→gly-gly: NPFREKKGGCALL, Gγ3-phe-phe shifted: NPFFFREKKCALL, Gγ9-gly-gly→phe-phe: NPFKEKFFCLIS, and Gγ9 with

Gγ3 pre-CaaX: NPFREKKFFCLIS. Similar to the trends in PIP3 generation observed in

HeLa cells expressing these mutants, MDA-MB-231 cells also showed a similar pattern of

PIP3 generation. For instance, both Gγ3 mutants (Gγ3-phe-phe→gly-gly and Gγ3-phe-phe shifted) expression showed ~ 60% reduced, and Gγ9 mutants (Gγ9-gly-gly→phe-phe and

Gγ9 with Gγ3 pre-CaaX) showed ~ 80% and ~ 90% increased PIP3 generation, compared to their corresponding WT Gγ types expressing cells (Fig. 4-8).

For the Western blot (WB) analysis, a Gβγ effector kinase; Akt, was selected since it controls a cohort of cellular signaling and behavioral events that govern metastasis (147-

152) (Fig. 4-6). Akt is implicated in a variety of human diseases such as prostate, breast,

Akt-PH-mCh Gγ3 Gγ9

Gγ3-FF→GG Gγ9-GG→FF

Gγ3-FF shifted Gγ9 with Gγ3 pre-CaaX

t=0 t=10 mins t=0 t=10 mins Fig. 4-8: Hydrophobic residues in the pre-CaaX of Gγ control Gβγ-driven PIP3 generation in MDA-MB-231 cells. MDA-MB-231 cells transfected with GFP tagged Gγ3, Gγ9, and their mutants as well as Akt-PH-mCherry (PIP3 sensor), before and after activation of endogenous α2AR using 100 μM NE. Images and the plot show that WT Gγ3, Gγ3-FF shifted, Gγ9-GG→FF, and Gγ9 with Gγ3 pre-CaaX expressing cells exhibiting robust PIP3 generations, while Gγ9 WT and Gγ3-FF→GG showing minor to no PIP3 responses (scale: 5 μm, error bars: SEM, n=8).

bladder, liver, and lung cancer, as well as neurodegenerative disorders such as Parkinson's disease and Alzheimer's disease (153-155). We analyzed the activated, phosphorylated form of Akt (p- p-Akt) in MDA-MB-231 cells expressing Gγ3, Gγ9, and their selected mutants. Since expression of the Gγ types are governed by the same T7 promoter and the length of the Gγ genes are nearly similar, we observed a ~ 60% transfection efficiency for 87 all the Gγ types. Twelve hours after transfection, cells were exposed to 100 μM norepinephrine for 30 minutes. Whole cell extracts were subjected to WB analysis to determine the levels of p-Akt. The results showed a 2-fold reduction in the p-Akt level with both the Gγ3 mutants (Gγ3-phe-phe → gly-gly and Gγ3- phe-phe shifted) cells, and a 2- fold increase with Gγ9 mutants (Gγ9-gly-gly → phe-phe and Gγ9 with Gγ3 pre-CaaX) compared to their WT Gγ cells (Fig. 4-9).

A. B.

shifted FF

→

CaaX

GPI

pre

Gγ9 with Gγ3 Gγ3 Gγ9 with

GFP

GFP unstimulated Gγ9

Gγ3

Gγ3 Gγ9

p-Akt

t-Akt

Actin

GFP

Fig. 4-9: Hydrophobic residues in the pre-CaaX of Gγ control Gβγ-driven kinases- activation in MDA-MB-231 cells. A. Pre-CaaX-dependent Akt phosphorylation. Whole cell lysates from MDA-MB-231 cells expressing GFP tagged Gγ3, Gγ9, and their mutants were immunoblotted using phospho-Akt (p-Akt), actin, GFP, and total Akt (t-Akt). B. Plot shows the quantification of each Gγ subunit compared to the GFP-GPI control. (error bars: SD, n=3 independent experiments)

4.2.7. Chemokine pathway directed cancer cell invasion is CT of Gγ dependent:

To confirm that invasion of MDA-MB-231 cells are driven by Gβγ and subsequent

PIP3 signaling, we initially performed invasion control experiments. Invasion was examined using transwell assay. Cells (2x105 in 100 μl) expressing GFP-GPI were seeded on transwell inserts with Matrigel (100 μl). Cells on Matrigel were supplemented either

A. B. Cell GFP-GPI PtX suspension

Transwell insert Matrigel Porous membrane Gallein Wortmannin

Medium Glass-bottomed dish Invaded cells Microscope objective

Fig. 4-10. Gβγ-governed MDA-MB-231 cell- invasion. A. Schematics showing quantification of invaded GFP- expressing cells. B. Images show invaded, live GFP-GPI expressing MDA-MB-231 cells and the dot-plot show number of invaded cells in the vehicle (control), PtX (Gi inhibitor), gallein (Gβγ inhibitor) or wortmannin (PI3K inhibitor) treated cells respectively. Cells without or with an inhibitor were seeded on Matrigel at 2x105 density and allowed overnight to undergo invasion under the influence of NE (100 μM) in the bottom chamber. Attenuated invasion by inhibitors indicates that the invasion is governed by GPCR (Gi), Gβγ→PI3K pathway. with Gi pathway inhibitor: Pertussis toxin (Ptx), Gβγ inhibitor: gallein, or PI3K inhibitor: wortmannin. In order to create a chemical gradient to asymmetrically activate endogenous

α2-ARs, 100 μM norepinephrine was added to the bottom chamber of the transwell filled with cell growth media; DMEM with 10% DFBS. After 16-24 hours, Matrigel was removed and inserts were placed in equilibrated cell culture medium containing glass- bottomed dish (Fig. 4-10A). Using a 40X objective, confocal images of invaded cells expressing GFP fluorescence were counted. All the inhibitors exhibited significant reduction in invasion; 98% with Ptx, 97% with gallein and 95% with wortmannin, compared to untreated cells. This data confirms that that the observed invasion is governed by Gi-pathway through Gβγ and subsequent PI3K activation (Fig. 4-10B).

Next, the influence of Gγ3, Gγ9, and their selected mutants on MDA-MB-231 cell invasion was examined using cells expressing each mutant and WT type Gγ (with a GFP tag) seeded on Matrigel. Invasion was induced by the addition of norepinephrine (100 μM) to the bottom chamber, and the GFP expressing invaded cells were visualized under 40X magnification and counted as described previously. Remarkably, MDA-MB-231cells expressing both Gγ3-phe-phe shifted mutant (NPFFFREKKCALL) and cells expressing

G3-phe-phe→gly-gly (NPFREKKGGCALL) mutant exhibited a ~ 4-5-fold reduction in invasion compared to WT Gγ3 (Fig. 4-11). Likewise, cells expressing Gγ9-gly-gly→phe- phe (NPFKEKFFCLIS) and Gγ9 with Gγ3 pre-CaaX (NPFREKKFFCLIS) showed ~ 4- fold and ~ 8-fold increase in cell invasion respectively, compared to WT Gγ9 expressing cells (Fig. 4-11). These data further demonstrated that the position of the phe residues closer to the prenylated-cys in Gγ strengthens the PM-affinity of Gγ, increases the PM residence time of Gβγ, and facilitates Gβγ-effector interactions, thus increases the signaling activation, modulating the invasive capacity of breast cancer cells.

GFP-GPI - NE GFP-GPI + NE Gγ3 Gγ3-FF→GG

Gγ3-FF shifted Gγ9 Gγ9-GG→FF Gγ9 with Gγ3 pre-CaaX

Fig. 4-11. Pre-CaaX residues of Gγ significantly influence Gβγ- governed MDA-MB-231 cell- invasion. Images and the dot- plot show MDA-MB-231 cell- invasion performed using cells expressing WT or mutant Gγ. Data show the influence of pre- CaaX residues in Gγ on cancer cell invasion (scale: 5 μm, error bars: SD, n=3 independent experiments).

4.2.8. G is uniquely evolved to have a short pre-CaaX to support heterotrimer formation and interaction with GPCRs:

The objective was to explore the relationship between the design of pre-CaaX motifs of Gγ and Gβγ function. The above demonstrated differential signaling of our Gγ mutant expressing cells indicate that pre-CaaX region of Gγ is a crucial signaling switch that turns ‘on’ and ‘off’ Gβγ signaling. Nevertheless, irrespective of its amino acid composition, mutants of Gγ with five gly residues in the pre-CaaX as in mutant Gγ9-lys-

91 glu-lys→gly-gly-gly (NPFGGGGGCLIS) (a) dimerizes with Gβ, (b) forms heterotrimers with Gα, and (c) the resultant heterotrimers interact with active GPCRs to produce GαGTP and Gβγ (Fig. 4-5A, D, E, H). The observed translocation of Gβγ9-NPFGGGGGCLIS from the PM to IMs upon GPCR activation clearly shows that, this Gγ mutant possesses the above a-c properties (Fig. 4-5A, D). Especially, this mutant shows that the sequence properties of the pre-CaaX region is irrelevant for G protein heterotrimer formation and activation.

Similar to Gγ proteins, Ras superfamily G proteins are also prenylated and contain polybasic pre-CaaX motifs. Interestingly, compared to the short pre-CaaX motifs in Gγ, analogous regions in prenylated-Ras proteins contain 3-4 times more (~ 20) residues (Fig.

4-12). Is the relatively short pre-CaaX motif in Gγ a crucial determinant of (i) heterotrimer

A. B.

Fig. 4-12: Short pre-CaaX motifs in Gγ compared to Ras family pre-CaaX. A. C- terminal sequence alignment of twelve Gγ subunits with some Ras superfamily proteins, showing PM-interacting short pre-CaaX regions in Gγ. B. WebLogo3 representation of the sequence prevalence in 12 Gγ CT alignment (top) and Ras family protein alignment (bottom). Note the significant difference in the length of pre-CaaX regions between the two protein types.

92 formation ability of Gγ and, (ii) heterotrimer-GPCR interaction and heterotrimer activation-dissociation? We hypothesized that the short pre-CaaX motifs in Gγ is crucial for the mobility of Gβγ as well as the heterotrimer on the PM and it facilitates Gβγ-effector and heterotrimer-GPCR interactions.

To examine this, we generated two Gγ3 mutants by replacing its pre-CaaX motif as well as the entire CT (both pre-CaaX + CaaX motifs) with analogous regions of the

A. B. GFP-Gγ3 with GFP-Gγ3 with mCh-Gγ9 mCh-Gγ9

GFP-Gγ3 KRas CT KRas pre-CaaX

GFP-Gγ3-KRas pre-CaaX GFP - Gγ3 -KRas CT After Before After

Before After Before After

Fig. 4-13. Activity of Gγ3-KRas mutants. A. HeLa cells expressing Gγ3 mutants with KRas pre-CaaX and the entire KRas-CT show, there resultant Gβγ lacks the translocation ability (contrary to Gγ3 cells), upon endogenous α2-AR activation with 100 μM NE. Plot shows the baseline normalized GFP fluorescence change in IMs over 10 mins. B. HeLa cells transfected with mCherry-Gγ9 and each GFP-Gγ3-KRas mutant (in the same cell) showing only the Gγ9 translocation but not the Gγ3-KRas mutant translocation upon α2-AR activation with 100 μM NE (scale: 5 μm, error bars: SEM, n=10).

GTPase, KRas. The resultant Gγ3-KRas mutants, which contained 20-residue long,

93 primarily polybasic pre-CaaX, were examined for their ability to form GPCR-activatable heterotrimers upon endogenous α2-AR activation in HeLa cells. Before activation of

GPCRs, both Gγ3-KRas mutants exhibited a PM distribution (Fig. 4-13A). Interestingly, only GFP-Gγ3 (black plot-yellow arrow), however not the Gγ3-KRas mutants (red and blue plots), exhibited Gβγ translocation upon GPCR activation (Fig. 4-13A-plot).

Also, when each of the Gγ3-KRas mutant with GFP tag and WT mCherry-Gγ9 are expressed in the same cell, only mCherry-Gγ9 showed translocation, while both Gγ3-KRas mutants failed to do so, indicating that GPCRs in cells are being activated (Fig. 4-13B).

However, upon half-cell photobleaching, GFP-Gγ3-KRas and Gβ1-mCherry showed a similar t1/2 value (t1/2= 77±4 s) for Fluorescence Recovery After Photobleaching (FRAP) in HeLa cells (Fig. 4-14). This indicates that Gγ3-KRas mutants form dimers with Gβ.

Control experiments show that the t1/2 of half-cell FRAP of Gβ1-mCherry is similar to Gγ3 in Gγ3 expressing and similar to Gγ9 in Gγ9 expressing cells, indicating that the associated

Gγ dictates Gβγ shuttling rates (APPENDIX A - Fig. A-1).

To examine if Gγ3-KRas mutants form heterotrimers, we compared t1/2 of half-cell

FRAP rates of GFP-Gγ3-KRas and mCherry-Gαo in the same cells (Fig. 4-14). These data exhibited nearly similar t1/2 for FRAP for the mutants (83±4 s) and Gαo (86±2 s), indicating that Gβγ3-KRas containing Gβγ dimers form heterotrimers with Gα (Fig. 4-14). Therefore, the inability of GFP-Gγ3-KRas to translocate upon GPCR activation shows that, the longer pre-CaaX region, either (a) prevents heterotrimer-GPCR interactions or (b) provides a significantly high PM-affinity for Gβγ, retarding its translocation. To examine these possibilities, we investigated if Gγ3-KRas containing heterotrimers compete with

endogenous heterotrimers to interact with activated GPCRs, since transfected Gγ types

always superseded signaling of endogenous Gγ (116).

A. GFP - Gγ3 - KRas CT Gβ1 -mCherry photobleach

Gαo-mCherry GFP-Gγ3-KRas CT

recovery

Before photobleach recovery Before photobleach recovery B. GFP-Gγ3-KRas pre-CaaX Gβ1-mCherry photobleach

Gαo-mCherry GFP-Gγ3-KRas pre-CaaX recovery

Before photobleach recovery Before photobleach recovery

Fig. 4-14. Gγ3-KRas mutants are capable of heterotrimer formation. A, B. Half-cell instantaneous photo-bleaching was performed in HeLa cells expressing Gαo- mCherry, Gβ1-mCherry, and GFP tagged Gγ3-KRas mutants, using FRAP-PA unit incorporated 8 mW 488 nm and 6 mW 594 nm to photobleach GFP and mCherry, respectively. The recovery of both fluorescent proteins was captured for 10 mins at 1Hz. Plots show the same recovery rate/t1/2 for Gαo, Gβ1, and both Gγ3-KRas mutants indicating that Gγ3-KRas mutants dimerize with Gβ, and the resultant Gβγ are capable of generating the heterotrimer with Gα (scale:5 μm, error bars: SEM, n=12).

Both WT HeLa cells (containing endogenous Gγ) and HeLa cells transfected with Gγ3-

KRas mutants failed to exhibit PIP3 generation (Fig. 4-15A). We previously showed that,

endogenous Gγ types in HeLa cells does not support PIP3 generation, which can be

overcome by expressing a high PM-affinity Gγ3 (116). Since we showed that endogenous

Gγ in RAW264.7 cells support PIP3 generation, we examined if Gγ3-KRas mutants alter 95 the PIP3 production in RAW cells. Upon activation of endogenous c5a receptors, both GPI-

GFP (control) (rate= 0.0029 s-1) and GFP-Gγ3-KRas mutants (0.0027 s-1 and 0.0024 s-1)

A. Gγ3 with KRas Gγ3 with WT Gγ3 pre -CaaX KRas CT

asal asal B NE

NE After B. Gγ3 with KRas Gγ3 with

WT Gγ3 Gγ9 pre-CaaX KRas CT

asal B

c5a

c5a c5a After After

Fig. 4-15. Inability of Gβγ-Gγ3-KRas mutants to transduce GPCR signaling. A. HeLa cells expressing Gγ3 or Gγ3-KRas mutants with Akt-PH-mCherry before and after endogenous receptor α2-AR activation with 100 µM NE. Only Gγ3 WT cells showed PIP3 generation, suggesting that Gγ3-KRas mutants containing heterotrimers or resultant Gβγ are dysfunctional. The plot shows baseline normalized mCherry (594 nm) fluorescence change at the PM. B. PIP3 generation in RAW 264.7 cells upon endogenous c5a receptor activation with 10 μM c5a, suggesting that endogenous Gγ types support Gβγ signaling. While overexpression of Gγ3 enhanced this PIP3 production, Gγ9 expression attenuated it. Interestingly, both Gγ3-KRas mutants did not influence the PIP3 production elicited by endogenous Gγ in RAW cells, suggesting the likelihood of they are forming inactive Gβγ and heterotrimers (scale: 5 μm, error bars: SEM, n=12). transfected RAW cells exhibited a similar PIP3 generation, while overexpression of Gγ3 enhanced and Gγ9 expression attenuated PIP3 production, suggesting that PIP3 generation observed in Gγ3-KRas mutant expressing cells is just due to the endogenous Gγ s (Fig. 4- 96

15B). Collectively, these data suggest that the 5-6 residue short pre-CaaX region of Gγ is uniquely evolved to act as a signaling switch for Gβγ while accommodating stringent functional G protein heterotrimers.

4.2.9. Short pre-CaaX regions of Gγ are evolutionarily conserved:

Since all Gγ types have relatively short pre-CaaX regions compared to other prenylated proteins such as Ras, we examined if this character is conserved beyond the mammalian genome. We investigated multiple Gγ subunit sequences of species ranging from primitive invertebrates/nematodes (Caenorhabditis elegans), insects (Fruit fly-

Drosophila melanogaster, diamondback moth- Plutella xylostella), fish (Zebrafish- Danio rerio) to plants (Thale cress- Arabidopsis thaliana, Asian rice- Oryza sativa) where GPCR signaling has been reported. Phylogenetic analysis of Gγ from those species together with human Gγ types showed that Gγ types with hydrophobic residues in their pre-CaaX cluster together and are divergent from the rest of the Gγ subunits (Fig. 4-16A). To understand this relationship between Gγ types from different species, Gγ CT sequences from multiple species were aligned. Surprisingly, similar to mammalian Gγ, all Gγ sequences examined exhibited 5 to 6- residues short pre-CaaX regions (Fig. 4-16B-red box). Remarkably, Gγ2

(GNG2) and Gγ3 (GNG3) from Zebrafish exhibited the phe-duo that support Gβγ effector activation (Fig. 4-16B-organge circle).

Gβγ signaling has been reported in Zebrafish where Gβγ regulates Rac activity to control cell polarity during the primordial germ cell migration (156). Similar to mammalian

Gγ1 and Gγ9, Zebrafish also possesses two gly residues in their GNGT1 (Gγ1) and GNGT2

(Gγ9) which may exert low PM-affinity for Gβγ, that may likely to support primarily Gα signaling. GNGT1 and GNGT2 of Zebrafish as well as Gγ types from other species possess primarily polybasic residues in the pre-CaaX regions.

Fig. 4-16. PM-interacting short pre-CaaX regions of Gγ are evolutionary conserved. A. Phylogenetic analysis of aligned Gγ sequences from insects, fish, plants, and human show the conserved short pre-CaaX regions, indicating the ability of Gβγ to associate with the conserved 7-TM structure of GPCRs. Further, conserved pre-CaaX properties among Gγ (blue box) in these species also indicate the common regulatory mechanism of Gβγ signaling across species. B. Partial alignment of Gγ sequences from species in A. They all show a short pre- CaaX region (red box) with 5, 6 amino acid residues similar to human Gγ types. Note the conserved hydrophobic character in GNG2 and GNG3 from Zebrafish (Danio rerio) similar to GNG2 and GNG3 in mammals (orange circle), and interestingly Gβγ signaling is reported in Zebrafish.

Interestingly, in species these Gγ types were sequenced, evidence for presence of pronounced Gβγ signaling is unclear. While this may suggest that the lack of hydrophobic residues is an indication for reduced Gβγ signaling compared to GαGTP, further investigations are required to confirm this. Overall, our analysis reveals that, Gγ subunits display conservation of sequence characteristics regardless of differences in the species type, likely to accommodate the conserved signaling architecture in GPCR-G protein signaling. Further, sequence-structural differences of Gγ subunit groups confer their relation to the specificity as well as selectivity of their designated function in heterotrimeric

G protein signaling.

4.3. DISCUSSION:

Activated heterotrimeric G proteins control two distinct types of signaling mechanisms; Gα governed and Gβγ mediated. In addition, literature broadly suggests that, different tissues and organs primarily utilize one of the above signaling types, depending on the functional specialization of cells. For an instance, GPCR-governed vision signaling in rod and cone photoreceptor cells of the eye is controlled by Gα-transducin and we believe that Gβγ only plays a supportive role in visual transduction by aiding heterotrimer formation and activation. We propose that, the reason for human eye to expresses low PM- affinity Gγ9 and Gγ1 is to achieve selective Gα signaling. The reduced PM-affinities of

Gβγ9 and Gβγ1 also likely to prevent unwanted and possibly deleterious Gβγ signaling in photoreceptor cells (157, 158). In GPCR-governed chemokine cell migration, the role of

Gαi/o signaling is relatively unclear while Gβγ plays the prominent role (159). The abundant expression of Gγ3 in macrophages clearly demonstrates that, cells utilize Gγ3 to

99 enhance the PM-affinity of Gβγ. Abundant expression of Gγ3 in the brain is another example where Gβγ signaling regulates functions including neuronal development (160).

Similar to mammals, evidence for recruitment of Gγ subtypes to achieve signaling selectivity can also be seen in C. elegans, that employ GPC-1, a Gγ type closely related to vertebrate Gγ1 and Gγ9, in sensory neurons to sense a wide variety of olfactory cues (15,

161). Fourteen nematode-specific Gi-like gpa genes are expressed in chemosensory neurons of C. elegans (162), collectively suggesting that Gγ types with a low PM-affinity are universally employed when Gα signaling is in demand.

We previously showed that, signaling and cell behaviors of Gβγ are Gγ-type dependent and also the PM-affinity and signaling efficacy of Gβγ are proportional (24, 39,

116). Although we have indicated the involvement of pre-CaaX region of Gγ controlling the PM-affinity of Gβγ, it was unclear how Gγ is evolved to act as a molecular tuner for

Gβγ activity in a Gγ-dependent manner, while equally supporting heterotrimer-GPCR interactions. It was also unclear what the molecular features in the Gγ family to impart distinct signaling abilities to Gβγ are, for cells to use expression regulation to gain the required signaling specificity of heterotrimeric G proteins. Current study reveals the crucial nature of the pre-CaaX region of Gγ as a multi-modal regulator of Gβγ in particular and

GPCR-G protein signaling in general.

The crystal structure of Gβγ-GRK shows that Gβγ not only laterally moves away from the activated receptor, spatially accommodating GRK2, but also it rotates by ~90 degrees to re-orient and interact with the Gβγ binding site on GRK2 (136-138). Over the simple notion that Gβγ stays bound to the PM, these observations indicate a more complex regulation of Gβγ behavior at the PM, diversely controlling Gβγ-effector interactions.

100

Among the remaining obvious questions, (i) could the relative orientation with the PM be important for Gβγ-effector interactions? (ii) is that why all Gβγ types are not activating effectors to a similar extent? (iii) what makes Gγ3 and Gγ2 to be more effector friendly?

(iv) is effector-interaction cross-section exploration by Gβγ governed by the CT of Gγ? are prominent. These questions led to the inquiry of unique sequence properties of Gγ3 and

Gγ2 compared to the rest Gγ-pool and resulted in identification of a phe-duo adjacent to the prenylated-cys, as a key regulator of Gβγ signaling. Interestingly, even several geranylgeranylated Gγ types such as Gγ5, γ7, γ10, and γ12, that are lacking hydrophobic residues, specially phe, adjacent to prenylated-cys also exhibited a significantly reduced

Gβγ signaling, compared to Gγ2 and Gγ3 (116). Therefore, we hypothesized that this phe- duo is as a must-have character in Gγ types with high Gβγ-effector activity. Systematic perturbation of these residues then revealed intricate molecular details of pre-CaaX residues and their relative location in the Gγ-CT in Gβγ signaling regulation.

The observed significant attenuation of Gβγ signaling after shifting phe-duo away from the prenylated-cys indicates that, being located away from the prenyl-anchor and thus the resultant reduced proximity of methyl-phenyl side chains of the shifted phe-duo to the

PM reduce the PM-affinity of Gβγ. When the phe-duo is next to the prenylated-cys, the lipid anchor is likely to pull Gγ towards the PM, facilitating methyl-phenyl groups to form strong hydrophobic interactions with the PM. This reinforcement appears to be a must- have criterion in the pre-CaaX of Gγ for it to enhance the PM-affinity and effector- interaction ability of Gβγ. This crucial involvement of the phe-duo was further confirmed by the reduced Gβγ signaling observed in cells expressing Gγ mutants with gly or Gγ types lacking phe residues adjacent to prenylated-cys of the CaaX motif (as in Gγ9).

101

Additionally, data also indicate a moderate enforcement of Gβγ-PM interactions by other hydrophobic residues in the pre-CaaX (e.g. leu, as in Gγ1), likely through hydrophobic interactions with the PM. However, data also collectively indicate a less obvious impact of charged residues in the pre-CaaX including lys, arg, and glu on Gβγ signaling and function of Gβγ overall. This is significant because, Ras family proteins primarily use lys residues to enforce PM-interactions.

Our results also show that when the hydrophobic residues are absent in the pre-

CaaX region, Gβγ dimer loses the ability to stay on the PM and thus translocates to IMs.

However, with the anchoring assistance from Gα, these Gγ subunits with weak PM-affinity can stay bound to the PM in the heterotrimer and undergo activation by the receptor. For instance, mutant Gγ9-lys-glu-lys→gly-gly-gly (NPFGGGGGCLIS), in which the pre-

CaaX region entirely composed of gly residues, produces functional heterotrimers, although the signaling of the resultant Gβγ is compromised. Selection of Gγ with such reduced PM-affinity is ideal for a cell that only desires Gα signaling upon GPCR activation.

Our data also indicate that, the influence of the type of prenylation on the PM- affinity as well as the signaling of Gβγ is not as strong as initially predicted. For instance, predictably farnesylated Gγ9 mutant with Gγ3 pre-CaaX expressing cells exhibited a nearly similar Gβγ induced PIP3 generation to that of WT Gγ3 cells. Our findings collectively show that the sequence properties of the pre-CaaX region of Gγ allow cells to have a range of Gβγ activities from completely “on” to “off” paradigms. Overall, they indicate that Gγ types are uniquely evolved and evolutionary conserved to have a relatively short PM- interacting pre-CaaX regions compared to other prenylated proteins (i. e., Ras family proteins) and by installing phe residues at unique locations, they manage to have a higher

102

PM-affinity and Gβγ activation ability when required. The ability of pre-CaaX residues to significantly change signaling of MDA-MB-231 cells, including alteration of kinase signaling in the chemokine pathway and invasion potential establishes the physiological relevance of pre-CaaX region of Gγ as well as the significance of the existence of twelve

Gγ types with distinct PM-affinities.

4.4. MATERIALS AND METHODS:

4.4.1. Reagents:

The reagents; Gallein (TCI AMERICA), Wortmannin (Cayman Chemical, Ann

Arbor, MI), c5a (Eurogentec), were initially dissolved in DMSO and then diluted with

HBSS (Gibco laboratories) before adding to cells. Norepinephrine (Sigma-Aldrich),

Carbachol (Fisher Scientific), Spingosine-1-phosphate (Cayman Chemicals) were dissolved in DI water and further diluted with HBSS during experiments. PtX (lyophilized powder) (Sigma-Aldrich) was reconstituted with DI water. SDF-1α (PeproTech) was reconstituted in DI water (100 μg/mL) and further diluted with a buffer containing 0.1%

BSA before adding to cells. Antibodies used were purchased from Cell Signaling

Technology, Santa Cruz Biotechnology, and Bio-Rad.

4.4.2. DNA constructs and cell lines:

Engineering of DNA constructs used; GFP-Gγ3, GFP-Gγ9, mCh-Gγ9, Gβ1- mCherry, Gαo-mCherry, Akt-PH-mCherry, and mCh-KRas have been described previously (99, 110, 111). GFP-GPI was a kind gift from Professor. N. Gautam’s lab,

Washington University, St. Louis, MO. Gγ3 and Gγ9 mutants were generated by PCR amplifying the parent constructs (GFP-Gγ3 and GFP-Gγ9) with overhangs containing

103 expected nucleotide mutations, DpnI (NEB) digestion (to remove the parent construct) followed by Gibson assembly (NEB) (112). Cell lines (HeLa and RAW 264.7 cells) were originally purchased from the American Tissue Culture Collections (ATCC) and authenticated using a commercial kit to amplify 9 unique STR loci.

4.4.3. Cell culture and transfections:

HeLa cells used in Gβγ translocation and PIP3 generation experiments were cultured in minimum essential medium (MEM; from CellGro) supplemented with 10% dialyzed fetal bovine serum (DFBS; from Atlanta Biologicals) and 1% Penicillin−Streptomycin (PS) in

60 mm tissue culture dishes. MDA-MB-231 cells used for invasion and PIP3 generation experiments were maintained in Dulbecco’s modified eagle medium (DMEM; from

CellGro) with 10% DFBS and 1% PS. RAW 264.7 macrophage cell line (PIP3 generation experiments with Gγ3-KRas mutants) were cultured in Roswell Park Memorial

Institute (RPMI) 1640 (10-041-CV; Corning, Manassas, VA) with 10% DFBS, and 1% PS supplementation. When the cells reach ~ 80% confluency, cells were lifted from the dish using versene-EDTA (CellGro) and resuspended in their growth medium at a cell density of 1×106 /mL. For imaging experiments (translocation and PIP3 generation), cells were seeded on 35 mm glass-bottomed dishes (1×105 cells on each) with 15 mm inner diameter, prepared using #1 German cover glasses.

Before cell seeding, dishes were incubated with 2 N NaOH for 30 min, ethanol washed, and sterilized for 1 hour using UV irradiation in a transilluminator. The following day of cell seeding, cells were transfected with appropriate DNA combinations using

Lipofectamine 2000 transfection reagent (Invitrogen), according to the manufacturer’s

104

0 protocol and stored in a 37 C, 5% CO2 incubator. Cells were imaged after 16 hours of the transfection.

4.4.4. Live cell imaging to monitor Gβγ translocation and PIP3 generation:

A spinning-disk XD confocal TIRF (total internal reflection) imaging system with a Nikon Ti-R/B inverted microscope, a Yokogawa CSU-X1 spinning disk unit (5000 rpm), an Andor FRAP-PA (fluorescence recovery after photo-bleaching and photo-activation) module, a laser combiner with 40−100 mW four solid-state lasers (with 445, 488, 515, and

594 nm wavelengths) and iXon ULTRA 897BV back-illuminated deep-cooled EMCCD camera was used to capture time-lapse image series of live cells. In Gβγ translocation and

PIP3 generation experiments, imaging was performed using a 60X, 1.4 NA (numerical aperture) oil objective. To examine the Gβγ translocation, GFP fluorescent tags on Gγ subunits (WTs and mutants) were imaged in every 1 s interval using 488 nm excitation−515 nm emission for 10 mins. In PIP3 generation experiments, mCherry tagged PIP3 sensor;

Akt-PH, was imaged using 594 nm excitation−630 nm emission red laser.

4.4.5. Western blot analysis:

MDA-MB-231 cells were, transfected with WT or mutant GFP-Gγ3, GFP-Gγ9, were exposed to 100 μM norepinephrine for 30 mins. After the incubation, cells were lysed and the whole cell lysate was run on a SDS polyacrylamide gel. Proteins were transferred to a PVDF membrane, blocked with 5% non-fat milk, and incubated with the primary antibodies specific for p-Akt, Actin, and GFP (Cell Signaling: #4060S, #9211S, Santa

Cruz: sc-47778), followed by the appropriate HRP-conjugated secondary antibodies (Bio-

Rad: #1705047, #1705046). The membrane was incubated with chemiluminescent detection solutions and exposed to an X-ray film. The protein band intensities on X-ray

105 films were quantified and normalized to the Actin and GFP level. Experiments were performed in triplicate, and quantification and statistical analysis of WBs was performed on three independent biological replicates.

4.4.6. Cell invasion assay:

1X Matrigel (100 μl) was set on the insert (with 8 μm membrane pores) in the transwell plate, and the following day MDA-MB-231 cells (2X105 cells) expressing WT and mutant Gγ types were seeded on the solidified Matrigel. To asymmetrically activate endogenous α2-ARs, 100 μM norepinephrine was added to the bottom chamber of the transwell, and after 16-24 hours, the inserts were placed on a glass-bottomed dish containing equilibrated cell culture media (200 μL). Using a 40X objective, confocal images of GFP tag in invaded cells expressing fluorescent Gγ and Gγ mutants were captured and the number of invaded cells were counted.

4.4.7. Statistical data analysis:

Results of all quantitative assays (Gβγ translocation, PIP3 generation, cell invasion) are expressed as standard error of mean (SEM) from n numbers of cells (indicated in the figure legends) from multiple independent experiments. Statistical analysis of cell invasion and

WB data of Gγ3, Gγ9 WT and their mutants were performed using two-tailed unpaired t- test. P value < 0.05 was considered as statistically significant.

106

Chapter 5

Engineering of Cone Opsin-based Chimeric Receptors with Improved Trafficking and Signaling

5.1. INTRODUCTION:

G protein coupled receptors (GPCRs) are considered as the largest family of cell surface receptors which control the majority of cellular signaling in the body (163). GPCRs share a seven transmembrane (TM) domains connected through three extracellular and three intracellular loops (ILs). They sense a wide array of extracellular/external signals such as peptides, hormones, lipids, small molecules, which transiently interact with the ligand binding pocket of the GPCR (1, 99). In contrast, light sensing GPCRs; rhodopsin and opsins, covalently bind to their ligand/inverse agonist; 11-cis retinal by forming a

Schiff base with a Lys residue (Lys296 in Rh) in the seventh TM helix of the receptor (3,

4). Upon photon absorption, receptor bound 11-cis-retinal (11-cis-retinylidene chromophore) isomerizes to all-trans-retinal which triggers conformational change in the receptor and activates it (5, 6). Binding of agonist results in the relaxation of intramolecular interactions and the formation of new interactions between the residues resulting movements of TM helices (164, 165). This TM rearrangements disrupt the salt bridge

107 between TM3 and TM7 and triggers the primary conformational changes not only in opsins, but also in all other GPCRs (164, 166-169). A greater movement of TM6 and TM3, exposes the hidden or previously less exposed internal surfaces of TM2, TM3, TM6, TM7, and the carboxy terminus (CT) of the receptor which enables the G protein interaction (5,

164, 165, 170, 171). G proteins which interact with the intracellular regions of the GPCR, especially with the IL3 and the CT (172-174), then exchange the bound nucleotide GDP to

GTP on the Gα subunit due to conformational changes, leading to G protein heterotrimer dissociation (8, 12, 14, 175, 176).

According to available crystal structures and conserved amino acid sequence patterns of GPCRs in different classes, it is evident that the seven transmembrane helices are highly conserved among GPCRs, which suggests a common activation mechanism (5,

164, 177-180). However, extracellular loops (ELs) and ILs as well as both N and C termini of GPCRs are highly divergent, and determine the signaling specificity of the receptor (5).

Although the light-induced activation and conformational changes in rhodopsin-driven signal propagation to the intracellular side of the receptor has been extensively studied

(163, 170, 181), how they recognize the appropriate G protein subtype based on the Gα type is not fully understood. To address this, several studies have been conducted so far by replacing the ILs of rhodopsin with those of other ligand binding GPCRs (i.e., muscarinic acetylcholine receptor, adrenergic receptor, prostaglandin receptor, and endothelin receptor) to identify the roles of each loop for selective G protein activation (182). Majority of the studies identified the IL3 as the region that interacts with the Gα subunit of G protein and have shown the switching of G protein recognition to the GPCR of the new IL3 (182).

A reduced activity has been observed with the IL2 replacement of bovine rhodopsin, which

108 led to the identification of seven residues including the “ERY” triad at the N terminus of

IL2 as a crucial motif for G protein activation rather than for selective recognition of G protein type (182).

Later, several other rhodopsin-based chimeras have been engineered by replacing all three ILs and the CT of rhodopsin with ligand activatable GPCRs (179, 183, 184). A set of rhodopsin-based chimeras were generated by replacing the IL1, IL2, IL3, and CT from hamster beta 2-adrenergic receptor (β2-AR) (179). The generated chimeras were purified and tested for their ability to bind the chromophore, to activate transducin, as well as for

Gαs signal transduction. Although those chimeras were designed to retain the conserved residues of rhodopsin, the chimeras with longer and multiple loop replacements were reported to have lower expression, lower chromophore formation as well as misfolding compared to the WT rhodopsin. Only the chimeras with shorter loop replacements were stated to show the activity similar to WT rhodopsin. This indicated that the size and number of loop replacements affect the folding of the chromophore which is crucial for the receptor functionality.

Two other bovine rhodopsin chimeras (opto-XRs) have been developed later with

ILs and the CT from Gq coupled human alpha 1a-adrenergic receptor (α1a-AR) and Gs coupled hamster β2-AR (184). Both opto-α1aAR and opto-β2AR were optimized for in vivo expression in mammalian cells and have been tested for Gq coupled phospholipase C activation induced intracellular calcium concentration and Gs coupled adenylyl cyclase activation mediated cAMP generation, respectively. Additionally, optical stimulation induced behavioral responses in mammals (mice) also have been tested with both opto-XR expressions and have reported to have nearly similar activities to their WT counterparts

109

(184). A photoactivatable-chemokine C-X-C receptor 4 (PA-CXCR4) also has been engineered to achieve effective trafficking of T cells to target tissues (183). PA-CXCR4 chimera also was designed by replacing the rhodopsin ILs and the CT with those regions of CXCR4, in order to switch the G protein signaling from Gαt to Gαi. Both intracellular signaling pathways (i.e., intracellular calcium level) and downstream physiological responses (i.e. cell migration) have been tested with the PA-CXCR4 chimera (183).

However, this PA-CXCR4 failed to show a good receptor activation thus signaling responses similar to chemokine receptor mediated signaling in vivo.

Other than rhodopsin-based chimeras, non-rhodopsin opsin-based chimeras also have been developed later to achieve much precise spatiotemporal control. A human color opsin-based chimera (CrBlue) with intracellular regions (ILs and the CT) from Jellyfish opsin has been shown to be optically controllable in mammalian cells (99). Likewise, several other chimeras have been developed over time using amphioxus opsin, scallop opsin, long-wave-sensitive and short-wave-sensitive cone opsins as well, which possessed different levels of activities (172). However, no chimera generated to date have been reported to have the complete switching of the signaling with a good efficiency. Although the reduced or loss of activity has been reported with IL1 and IL2 loop replacements in few studies (179, 182), the cause for reduced or loss of activity is not completely understood so far.

5.2. RESULTS:

5.2.1. Screening rhodopsin-based chimeras and their limitations:

110

In this study, we first examined the activity of current rhodopsin-based chimeras by characterizing their G protein activating abilities. We focused on the Gαt to Gαs switched chimeras and selected opto-β2AR which has been shown effective optically controllable Gs activity (184). Opto-β2AR has been developed in 2009 by introducing all the ILs as well as the CT of β2AR to rhodopsin (Fig. 5-1A). With opto-β2AR chimera, we measured both GPCR-G protein activation and downstream signaling as well. To detect the GPCR-G protein activation, we used the Gγ9 assay which measures the real time Gβγ- translocation from the plasma membrane (PM) to internal membranes (IMs) upon receptor activation in live cells (185). Gαs signaling was measured by mini Gs translocation from the cytosol and recruitment to the PM (186). Mini Gs is the GTPase activating domain of

Gαs subunit which is genetically engineered to improve protein stability in vitro with a truncated N-terminus, and few key modifications (i.e., deletion of the helical domain) to stabilize the receptor-mini G protein complex (186).With a Gs coupled GPCR activation, mini Gs which is initially in the cytosol of a cell translocate to the PM to interact with the activated receptor by forming the G protein hetereotrimer with endogenous Gβ and Gγ.

Initially, to check the receptor and G protein activation with the current chimera, opto-β2AR chimera was expressed in HeLa cells along with Gγ9 and checked for the Gβγ translocation from the PM to IMs upon opto-β2AR activation. To activate the chimeric receptor opto-β2AR, 50 μM 11-cis retinal was added in dark and 445 nm blue light was exposed on cells which resulted in Gβγ translocation. However, the level/extent of translocation was significantly lower (~80%) (Fig. 5-1B) compared to the translocation observed with WT β2AR expressing cells activated with 10 μM isoproterenol (Fig. 5-1C).

Next, we checked the mini Gs translocation with opto-β2AR activation. HeLa cells

111

Rhodopsin β2AR Opto-β2AR

B. C. Opto-β2AR-YFP mCh-Gγ9 β2AR-Dronpa mCh-Gγ9

t=0 t=200 s t=0 t= 200 s

Iso

D. E. Opto-β2AR-YFP mini Gs-mCh β2AR-Dronpa mini Gs-mCh

t=0 t= 300 s t=0 t= 300 s Iso

112

Fig. 5-1: Limitations of rhodopsin-based chimeras: A. Crystal structures of rhodopsin (PDB ID: 1F88), β2AR (PDB ID: 3SN6) and SWISS-MODEL predicted structure of opto-β2AR representing the opto-β2AR chimera engineering. B, C. Images of a HeLa cell expressing mCh-Gγ9 with B. opto-β2AR-YFP or C. β2AR-Dronpa. β2AR activation with 10 μM isoproterenol (Iso) showed a profound Gβγ translocation from the PM to IMs while opto-β2AR activation with 50 μM 11-cis retinal addition followed by 445 nm blue light exposure showed relatively minor Gβγ translocation. Plots show the relative mCh florescence accumulation in IMs over time with each receptor activation. D, E. HeLa cells expressing mini Gs-mCh either with D. opto-β2AR-YFP or E. β2AR-Dronpa. β2AR activation with isoproterenol displayed a vibrant PM recruitment of mini Gs-mCh, measured by the reduction of mCh fluorescence in the cytosol whereas opto-β2AR activation with retinal and blue light resulted a very faint response (error bars: SEM, n=8, scale: 5 μM). expressing mini Gs-mCherry and opto-β2AR-YFP were activated and monitored the mCherry fluorescence of mini Gs accumulation on the PM upon opto-β2AR activation with

11-cis retinal and blue light. Although a clear mini Gs accumulation on the PM was observed with WT β2AR (Fig. 5-1E), the accumulation was barely detectable with opto-

β2AR (Fig. 5-1D). Another significant change observed between the WT β2AR and the opto-β2AR chimera was the receptor expression on the PM. The WT β2AR was well expressed on the PM (Fig. 1C, E), while the opto-β2AR was not (Fig. 5-1B-E).

5.2.2. Blue opsin is an efficient signaling activator:

Since the rhodopsin-based chimeras failed to show proper receptor activation and

G protein signaling, and due to the difficulty of maneuvering rhodopsin due to its extreme light sensitivity, we decided to employ a color opsin as the backbone of our study as they display a defined spectral selectivity. Out of the color opsins (red, green, and blue) we selected blue opsin considering its spectral characteristics. Absorption maximum of blue opsin is at 414 nm which is further away from the absorption and emission spectrum

113 maximum wavelengths of fluorescent proteins used for live cell imaging. Therefore, we can simultaneously monitor the dynamics of molecules (i.e., effectors, sensors) tagged with a fluorescent protein without activating the receptor blue opsin (Table 5-1).

Table 5-1: Excitation and emission wavelengths of common fluorescent proteins Table 5-1: Excitation and emission wavelengths of common fluorescent proteins Fluorescent protein Excitation wavelength Emission wavelength Green fluorescent protein (GFP) 488 507 Cyan fluorescent protein (CFP) 456 480 mTurquoise 434 474 Yellow fluorescent protein (YFP) 513 527 Venus 515 528 mCherry 587 610

Although blue opsin fulfilled the requirements of a typical light activatable GPCR, upon expression in live cells, they showed a limited lifespan on the PM, which results in receptor internalization after 1 day of DNA transfection in to cells (Fig. 5-2A). Therefore, it was challenging to conduct experiments the following day of transfection due to limited receptor expression on the PM. Hence, we engineered the receptor blue opsin by introducing mutations to the C terminal phosphorylation sites. With multiple rounds of extensive mutations and screening, we were able to obtain an enhanced version of the receptor blue opsin (E-blue opsin) with improved expression on the PM as well as signaling even after 5-7 days of transfection (Fig. 5-2A). Comparative screening of WT blue opsin and E-blue opsin showed that E-blue opsin is a superior light activatable GPCR which showed signaling nearly similar to the signaling levels obtained with ligand inducible

GPCRs (Fig. 5-2B). Therefore, we used this E-blue opsin as the backbone for chimera generation in this study.

114

A. BO WT - mCh BO - E - mCh B. BO-E-mCh GFP-Gγ9

t=0 300 s Day 7 Day 1 Day 7 Day

Fig. 5-2: Enhanced blue opsin (BO-E) is as efficient as ligand activatable GPCRs. A. Images of HeLa cells expressing WT BO and BO-E captured on the first day and seventh day of transfection. Note the PM localization of BO-E compared to the internalized BO WT on day 7. B. BO-E and GFP-Gγ9 expressing Hela cell shows prominent Gβγ9 translocation even after seven days of translocation upon BO-E activation with 11-cis retinal and 445 nm blue light exposure (scale: 5 μM).

5.2.3. IL3 is the most crucial region of the receptor for G protein interactions:

According to the available crystal structures (PDB ID:3SN6) as well as the literature reports, the IL3 is the region of a GPCR which shows to be interacting with the

Gα subunit of G protein (187). The crystal structure of β2AR interacting with a G protein shows that the Gα subunit of the G protein is embedded in the barrel of transmembrane helices of the receptor with a close proximity to the IL3. We anticipated that the extended

IL3 compared to the length of other loops (IL1 and IL2) could make it extra flexible making

115 it enable to re-orient accordingly in order to interact with the Gα subunit of the G protein.

Therefore, we decided to first start with an IL3 replacement of blue opsin.

5.2.3.1. IL1 and Il2 replacements cause defective receptor trafficking to the PM:

In a preliminary study of chimeric receptor engineering, we had generated a blue opsin chimera with IL3 replaced from the IL3 of CXCR4 (BO-CXCR4-IL3-18 AA) which showed a good G protein activation (Fig. 5-3C). However, due to its poor trafficking on to the PM, we decided to replace other loops also from CXCR4 to the blue opsin.

Nevertheless, since a crystal structure is not yet available for color opsins, we had to rely on the sequence homology between selected receptors when designing the chimeras

(APPENDIX C). Individual replacements of blue opsin IL1 and IL2 with those of CXCR4

(BO-CXCR4-IL1-9 AA and BO-CXCR4-IL2-12 AA) resulted in a significant loss of the receptor activation which was measured by Gβγ9 translocation (Fig. 5-3A, B). Plasma membrane trafficking of both chimeras were also impaired where majority of the chimeric receptors were trapped in IMs, suggesting a protein misfolding (Fig. 5-3A, B).

When the chimeras were overexpressed in HeLa cells with E-blue opsin, only the

E-blue opsin was expressed on the PM while the chimeras were trapped in IMs (Fig. 5-4).

To make sure that the impaired trafficking of chimeras to the PM is not due to the overexpression of other DNA which we used to measure the receptor activity (i. e., Gγ9,

Akt-PH-Venus), we expressed the chimeras alone in HeLa cells and observed their distribution pattern. Even with the individual expression, the receptor distribution on the

PM was not improved (Fig. 5-4C). Also, we checked the chimera expression after 6-8 hours of the transfection because we suspected that the receptors might have gotten internalized

116

A. BO-CXCR4-IL1 CXCR4: IVGNGLVILVMGYQKKLRSMTDKYRLHLS BO: FPLNAMVLVATLRYKKLRQPLNYILVNVS

Mutant 1

CXCR4: IVGNGLVILVMGYQKKLRSMTDKYRLHLS BO: FPLNAMVLVATLRYKKLRQPLNYILVNVS

Mutant 2

B. BO-CXCR4-IL2 CXCR4: FISLDRYLAIVHATNSQRPRKLLAEKVVY BO: FLAFERYIVICKPFGNFR----FSSKHAL

Mutant 1

CXCR4: FISLDRYLAIVHATNSQRPRKLLAEKVVY BO: FLAFERYIVICKPFGNFR----FSSKHAL

Mutant 2

C. BO-CXCR4-IL3 CXCR4: YCIIISKLSHSKGHQK------RKALKTTV BO: YTQLLRALKAVAAQQQESATTQKAEREVSRMVV

Mutant 1

CXCR4: CIIISKLSHSKGHQK------RKALKT BO: TQLLRALKAVAAQQQESATTQKAEREVSRM

Mutant 2 117

Fig. 5-3: Engineering of blue opsin-CXCR4 chimeras: HeLa cells expressing each BO-CXCR4 A. IL1, B. IL2, or C. IL3 chimeras with different loop lengths along with GFP-Gγ9 show different level of Gβγ9 translocation with chimera activation upon 50 μM 11-cis-retinal addition followed by 445 nm blue light exposure. SWISS-MODEL predicted crystal structures of chimeras and sequences substituted with both receptors (blue opsin-E and CXCR4) are also shown for each chimera (error bars: SEM, n=8, scale: 5 μM).

over time. But even after 6-8 hours of transfection, the receptors were trapped in IMs which eliminated the possibility of receptor internalization (Fig. 5-4D).

5.2.3.2. Size of the loop replacements determine proper folding and expression of receptor:

Although the original loop length of blue opsin was conserved in IL1 and IL2 chimeras, considering that the loop replacement could have introduced a strain on the loop

(IL) flanking transmembrane helices which could result in improper folding of the receptor, we next shortened the size of the loop replacement. According to the literature reports, the chimeras with a longer loop replacement showed a higher level of misfolding which resulted in instability of the chromophore compared to that of the shorter replacement which had been measured by comparing the A280/A500 ratio with WT rhodopsin which reflects the extent of regeneration; a measure of protein folding (179).

A BO-CXCR4-IL3 with 13 residue IL3 replacement, showed a better PM localization with increased Gβγ9 translocation which was nearly similar to that with both

WT blue opsin and CXCR4 (Fig. 5-3C). Shortening of IL1 and IL2 (BO-CXCR4-IL1-6

AA and BO-CXCR4-IL2-5 AA) also increased the receptor-G protein activation which

118

A. BO - CXCR4 - C. BO-CXCR4- BO-CXCR4-

IL1-mCh E-BO-GFP Overlay E-BO-GFP IL1-mCh IL2-mCh

B. BO-CXCR4-

IL2-mCh E-BO-GFP Overlay Cell 3 Cell 2 Cell 1 Cell 2 Cell Cell 3

D. BO-CXCR4- BO-CXCR4- IL1-mCh IL2-mCh

Fig. 5-4: Defective PM trafficking of BO-CXCR4 chimeras: Images show the HeLa cells expressing A. BO-CXCR4-IL1, B. BO-CXCR4-IL2 chimeras with E-BO- GFP in the same cell. Note the defective trafficking of BO-CXCR4 chimeras while E-BO is properly expressed on the PM. C. Individual expression of only one construct in HeLa cells also shows that BO-CXCR4 chimeras are trapped in IMs compared to E-BO which is expressed on the PM. D. HeLa cells expressing individually expressing each BO-CXCR4 chimeras after 6-8 hours of transfection. Note the defective PM trafficking of both BO-CXCR4-IL1 and BO-CXCR4-IL2 (scale: 5 μM). was measured by Gβγ9 translocation, but not to the same extent as WT receptors (Fig. 5-

3A, B). Also, no improvement of the chimera expression on the PM was observed (Fig. 5-

3A, B).

Since BO-CXCR4-IL3-13 AA chimera showed an improvement in both expression as well as G protein activation, we next tested Gβγ mediated PIP3 generation ability of the

IL3 chimera. To monitor the PIP3 generation, the PIP3 sensor Akt-PH-Venus was used which gets accumulated on the PM upon receptor activation induced PIP3 generation on the PM. RAW 264.7 macrophage cells transfected with the PIP3 sensor and BO-CXCR4- 119

IL3 (13 AA)-mCh showed PIP3 sensor accumulation on the PM upon optical activation of the chimera by retinal addition and 445 nm blue light exposure (Fig. 5-5A). Also, RAW cells expressing BO-CXCR4-IL3 (13 AA) chimera showed directional cell migration nearly similar to that with WT blue opsin expression upon localized optical activation of the receptor (Fig. 5-5B).

A. B. Akt-PH-Venus E-BO-mCh BO-CXCR4-IL3-mCh t=0 t=0

t=10 min t=10 min

t=0 t=20 mins t=0 t=20 mins

Fig. 5-5: Improved BO-CXCR4-IL3(13 AA) chimera induces PIP3 generation and macrophage migration. A. RAW 264.7 cells were transfected with either WT blue opsin or blue opsin-CXCR4-IL3 chimera along with the PIP3 sensor Akt- PH-Venus. Receptor blue opsin and the chimera were activated with 50 μM 11-cis retinal addition followed by 445 nm light exposure. Cells show the PIP3 accumulation on the PM. Plot shows the quantified PIP3 level on the PM (scale: 5 μm, error bars: SEM, n=10). B. RAW 264.7 cells expressing mCherry tagged E-BO or blue opsin-CXCR4-IL3, supplemented with 50 µM 11- cis- retinal. Receptors were activated in confined regions using a 445 nm laser in every 3 s interval (white boxes). Images show cells before and after 20 mins of receptor activation. Bar graph shows the relative displacement of cells’ leading and trailing edges, with receptor activation (scale: 5 μm).

120

Considering the functionality observed, we next introduced the shortened IL1 (6

AA) and IL2 (5AA) to the BO-CXCR4-IL3 (13 AA) chimera expecting an increased activity with good receptor trafficking. We first introduced the IL1 and IL2 individually and screened the receptor activation by measuring the Gβγ9 translocation. None of the introductions provided the expected result, but they reduced the functionality of BO-

CXCR4-IL3 chimera instead. This unexpected result further confirmed that the IL3 is the region which is crucial for signaling and suggested that the IL1 and IL2 may be involved in maintaining the receptor integrity. Since the IL1 and IL2 are relatively shorter compared to the IL3, based on the experimental results observed, we believed that even a very small change could affect the receptor architecture due to the limited flexibility of IL1 and IL2.

Therefore, in order to check if an increased flexibility of the loops would make a difference, we introduced 3 gly residues at the PM-IL interface to both sides of the IL1 and IL2 to increase the flexibility to those two loops. However, this extra base addition also did not improve the receptor expression on the PM nor G protein signaling, which directed us to manipulated without disturbing the receptor integrity.

5.2.4. Engineering of β2AR chimeras:

Applying the lessons learnt from the BO-CXCR4 chimeras, we next started generating a blue opsin-β2AR chimera with signaling switched from Gi to Gs. Since Gs pathway is the exact opposite of the Gi pathway, a complete opposite G protein activity was expected which could be easily detected and differentiated.

5.2.4.1. Light-dependent β2AR-specific signaling by chimeric receptors:

121

Since it was established that the IL3 is the most crucial region which determines the G protein selectivity as well as signaling, both from the literature reports and our experimental results, we first started replacing the IL3 of BO with that of β2AR. According to the available crystal structure of β2AR (PDB ID: 3SN6), the loop regions of β2AR were recognized and BO loop regions were identified by considering the homology with rhodopsin, in which the crystal structure has been resolved (PDB ID: 1F88). Taking precautions to not to disturb the receptor integrity by introducing a large number of residues at once, we first selected only two residues in the middle of the BO IL3 and replaced them with the middle two residues from the IL3 of β2AR. The two residues replaced BO-β2AR-

IL3 chimera managed to show a good PM distribution, however with no Gβγ9 translocation from the PM to IMs upon optical activation of the chimera (Fig. 5-6A). Next, we replaced four more residues, two from each side of the loop, expecting a receptor activity. To our surprise, the resultant BO-β2AR-IL3 chimera with 6 residues exchanged also showed no activity, which was measured by Gβγ9 translocation, even a considerable level of chimeric receptor expression on the PM was observed (Fig. 5-6B). However, we kept increasing the size of the loop replacement by substituting two residues at a time to both sides of the last loop replacement. The chimera generated with a 10-residue replacement showed a good trafficking of the chimera on the PM, which also showed a reasonable Gβγ9 translocation

(Fig. 5-6C). The next chimera generated had a 14 amino acid replacement and it had the best receptor expression on the PM. Also, the BO-β2AR-IL3 (14 AAs) chimera showed a

Gβγ9 translocation nearly similar to the WT BO receptor (Fig. 5-6D). Also, we introduced the whole IL3 of β2AR, which is 52 residues long, to BO and examined its Gβγ translocation ability. Although a Gβγ9 translocation was observed with BO-β2ARIL3 (52

122

A. B. BO-β2AR-IL3 BO-β2AR-IL3 2 AA)-mCh GFP-Gγ9 6 AA)-mCh GFP-Gγ9

t=0 t=200 s t=0 t=200 s

C. D. BO-β2AR-IL3 BO-β2AR-IL3 (10 AA)-mCh GFP-Gγ9 (14 AA)-mCh GFP-Gγ9

t=0 t=200 s t=0 t=200 s

E. BO-β2AR-IL3 52 AA)-mCh GFP-Gγ9

t=0 t=200 s

123

Fig. 5-6: Loop length dependent receptor activation: Images show the HeLa cells expressing GFP-Gγ9 and each BO-β2AR-IL3 chimera with A. 2 AA, B. 6 AA, C. 10 AA, D. 14 AA, and E. 52 AA residue IL3 replacements. Upon each chimera activation with 50 μM 11-cis retinal addition followed by 445 nm blue light exposure showed a prominent Gβγ9 translocation with BO-β2AR- IL3(14 AA) chimera, relatively weak activation with BO-β2AR-IL3(10 AA), BO-β2AR-IL3(52 AA) chimeras, and almost no activation with BO-β2AR- IL3(2 AA) and BO-β2AR-IL3(6 AA) chimeras. Plots show the relative Gβγ9 translocation measured by GFP fluorescence accumulation in IMs over time. The gray line in plots shows the comparative Gβγ9 translocation observed with Gs coupled β2AR activation with 10 μM isoproterenol (error bars: SEM, n=10, scale: 5 μM).

AAs) chimera, it was considerably lower than the translocation observed with BO-

β2ARIL3 (14 AAs) (Fig. 5-6E). Also, we introduced the whole IL3 of β2AR, which is 52 residues long, to BO and examined its Gβγ translocation ability. Although a Gβγ9 translocation was observed with BO-β2ARIL3 (52 AAs) chimera, it was considerably lower than the translocation observed with BO-β2ARIL3 (14 AAs) (Fig. 5-6E).

Since the IL3 14 and 52 residue replacements showed a good receptor-G protein activation, we next decided to check the Gs specific signaling with those two chimeras.

HeLa cells expressing the above-mentioned chimeras (mCherry tagged) and mini Gs-

Venus were activated with 11-cis retinal addition followed by 445 nm blue light exposure and monitored the mini Gs recruitment to the PM. With both chimeras, a slight mini Gs translocation from the cytosol to the PM was observed denoting the Gs coupled GPCR

activation which was measured by the reduction of cytosolic Venus fluorescence (Fig. 5-

7). However, the overall mini Gs recruitment data observed were not comparable with the extent of Gβγ translocation observed. These results led us to question the Gβγ translocation with BO-β2AR-IL3 mutants, which could have occurred due to Gαi heterotrimer

124

A. BO-β2AR-IL3 (14 AA)-mCh

t=0 t=300 s 0 B. BO-β2AR-IL3 (52 AA)-mCh

t=0 t=300 s 0

Fig. 5-7: Gαs activity of BO-β2AR-IL3 chimeras. HeLa cells expressing A. BO- β2AR-IL3(14 AA) + mini Gs-Venus, B. BO-β2AR-IL3(52 AA) + mini Gs- Venus show the PM recruitment of mini Gs upon each chimera activation with 50 μM 11-cis retinal addition followed by 445 nm blue light exposure. Mini Gs translocation to the PM was measured by the fluorescence loss of Venus in the cytosol (error bars: SEM, n=8, scale: 5 μM). activation. Therefore, in order to eliminate the possibility of Gαi heterotrimer activation from remaining BO IL3 regions and due to the promiscuity of β2AR which can interact with Gαi containing heterotrimers also to a certain extent, we treated BO-β2AR-IL3 chimeras (with 10, 14 and 52 AA IL3 replacements) expressing cells with 1 μg/mL of

Pertussis toxin (PtX), which inhibit the Gi pathway and checked their Gβγ translocation.

Surprisingly, PtX treatment reduced the Gβγ translocation with all three chimeras and could observe only a minor activity (Fig. 5-8) which explained the marginal mini Gs recruitment to the PM.

125

A. BO-β2AR-IL3 (10 AA)-mCh GFP-Gγ9

t=0 200 s

B. BO-β2AR-IL3 (14 AA)-mCh GFP-Gγ9

t=0 200 s

C. BO-β2AR-IL3 (52 AA)-mCh GFP-Gγ9

t=0 200 s

Fig. 5-8: Reduced activation of BO-β2AR-IL3 chimeras upon PtX treatment. HeLa cells

expressing GFP-Gγ9 and each BO-β2AR-IL3 chimera with A. 10 AA, B. 14 AA, and C. 52

AA residue replacements show reduced Gβγ9 translocation upon each chimera activation with

50 μM 11-cis retinal addition followed by 445 nm blue light exposure. Note: All the cells were

treated with PtX overnight to inhibit Gi activation due to the β2AR promiscuity (error bars:

SEM, n=8, scale: 5 μM).

In order to make sure that the PM trafficking and signaling issues obtained with

BO-CXCR4 chimeras are not specific only to Gi coupled receptors (i. e., CXCR4), we 126 generated two blue opsin chimeras with the IL1 and IL2 from β2AR. However, similar to

BO-CXCR-IL1 and BO-CXCR4-IL2 chimeras, none of the BO-β2AR-IL1 and BO-β2AR-

IL2 chimeras showed significant Gαs signaling nor G protein activation. Also, both chimeras trafficking to the PM was not good which further supporting that the IL1 and IL2 replacements affect the receptor folding and thus signaling.

5.2.4.2. Carboxy terminus of the receptor further strengthens the β2AR specific signaling of chimeric receptors:

Although the BO-β2AR-IL3 (14AAs) and BO-β2AR-IL3 (52 AAs) chimeras showed Gs activation, the responses observed with both chimeras were significantly lower compared to the responses with Gs coupled WT β2-AR (Fig. 5-1E, 5-7). Since it has been reported that the CT of β2-AR is also involved in generating the Gαs binding pocket by making a pseudo loop which results from the Cys341 palmitoylation (188), we decided to introduce the β2AR CT to the BO-β2AR-IL3 chimeras which showed slight activity (10,

14, and 52 AA), expecting an enhanced Gαs activity. The generated BO-β2AR-IL3-CT chimeras were first tested for the receptor-G protein activation ability by measuring the light-induced Gβγ9 translocation with PtX treatment. As expected, all three chimeras with the β2AR CT introduction showed an increased Gβγ translocation (Fig. 5-9A-C).

Interestingly, the CT replacement improved the mini Gs recruitment to the PM suggesting an enhanced Gs activity (Fig. 5-9D-F).

Also, we observed that the PM bound mini Gs reverses back to the cytosol with the cessation of blue light exposure to inactivate the receptor which was represented by the decreased mini Gs fluorescence in the cytosol. Re-activation of the receptor resulted in

127 mini Gs fluorescence reduction in the cytosol again denoting that the mini Gs recruitment is reversible (Fig. 5-10). Since the mini Gs recruitment to the PM was quantified by

A. B. C. BO-β2AR-IL3 BO-β2AR-IL3 BO-β2AR-IL3 (10 AA)-CT-mCh (14 AA)-CT-mCh (52 AA)-CT-mCh

t=0 t=200 s t=0 t=200 s t=0 t=200 s GFP0 -Gγ9 GFP-Gγ9 GFP-Gγ9

D. E. F. BO-β2AR-IL3 BO-β2AR-IL3 BO-β2AR-IL3 (10 AA)-CT-mCh (14 AA)-CT-mCh (52 AA)-CT-mCh

t=0 t=250 s t=0 t=250 s t=0 t=250 s mini Gs-Venus mini Gs-Venus mini Gs-Venus

128

Fig. 5-9: Gαs signaling reinforcement by the introduction of the CT. HeLa cells expressing GFP-Gγ9 and each BO-β2AR-IL3 chimera with A. 10 AA, B. 14 AA, and C. 52 AA residue IL3 and complete CT replacements from β2AR. Each chimera was activated with 50 μM 11-cis retinal addition followed by 445 nm blue light exposure. Note: All the cells were treated with PtX overnight to inhibit Gi activation due to the β2AR promiscuity. D-F. HeLa cells expressing each of the BO-β2AR-IL3-CT mutant along with mini Gs- Venus show the Gαs activity with each chimera, which was measured by the fluorescence loss of Venus in the cytosol. The gray line in plots (in A-F) shows the responses (i.e., Gβγ9 translocation, PM recruitment of mini Gs) observed with Gs coupled β2AR activation with 10 μM isoproterenol (error bars: SEM, n=10, scale: 5 μM). measuring the reduction of mini Gs fluorescence in the cytosol, a control experiment was conducted in the same way however without retinal addition to make sure that the

fluorescence loss is not due to photo bleaching of the fluorescent tag on mini Gs. That

Fig. 5-10: Reversibility of mini Venus

- Gs translocation: Images show the HeLa t=0 300 s 600 s 800 s cells expressing mini mini Gs mini Gs-Venus and BO- β2AR-IL3(52AA)-CT. Upon chimera Dar activation with 11-cis k retinal addition followed by 445 blue light exposure, mini Gs translocase to the PM resulting Venus fluorescence loss in the cytosol. With the cessation of receptor activation (by activation (by incubating cells in dark), the mini Gs reverses back to the cytosol which re-translocates to the PM upon receptor re-activation by shinningincubating blue light cells(scale: in5 dark), the mini Gs μM). reverses back to the cytosol which re- translocates to the PM experiment resulted in no significant loss of cytosolic fluorescence other than simple upon receptor re- activation by shinning fluctuations, verifying the mini Gs PM recruitment results obtained with the chimeras. blue light (scale: 5 μM). 129

Overall, from the generated chimeras, BO-β2AR chimera with 52 AAs IL3 and complete CT replacement showed the best expression as well as the Gαs activity so far.

5.3. DISCUSSION:

The objective of this study was to design and develop a light activatable Gs coupled chimeric receptor which shows Gs signaling and can be controlled by light in a spatial and temporal manner. Due to the limitations associated with so far developed current chimeric receptors based on the rhodopsin, we selected a non-rhodopsin color opsin; blue opsin, to obtain much precise spatiotemporal control of the system. By comparing the signaling of rhodopsin-based chimeras with their wild type counterparts, it was apparent that the rhodopsin-based chimeras have a relatively poor signaling ability compared to ligand activatable receptors. Therefore, the non-rhodopsin-based blue opsin was selected as it has specific spectral characteristics which becomes advantageous when to be used with other fluorescent proteins in experiments. The enhanced blue opsin we generated due to the limited life time of WT blue opsin on the PM showed good PM localization as well as signaling even after 5-7 days, was a good candidate for the PM trafficking issue of the rhodopsin-based chimeras. Also, it showed almost similar G protein activation to that of ligand induced GPCRs. Therefore, we used the enhanced blue opsin as the backbone to develop the chimeras in this study.

The IL3 is been reported as the central region of a GPCR that interacts with the G protein, which is also supported by the available crystal structures of GPCR-G protein complexes. Therefore, our initial work of chimeric receptor engineering by substituting the

130

IL3 of blue opsin with IL3 from CXCR4, revealed that only the IL3 can be modified but not IL1 and IL2 because IL1 and IL2 replacements significantly lost or eliminated the receptor activity. Even a functional IL3 chimera can be made non-functional just by introducing a change to the IL1 or IL2. Nevertheless, since the functional BO-CXCR4-IL3 chimera failed to traffic well to the PM, we predicted that a long loop replacement can introduce a strain to the neighboring/flanking TM regions, leading to misfolding of the receptor, which has been reported in the literature as well. Better PM localization of the

BO-CXCR4-IL3 chimera obtained with a much shorter IL3 replacement which retained/increased the activity also validated that hypothesis. However, the shortening of neither IL1 nor IL2 showed any improvement of PM localization of chimeras but some minor receptor activity which is not significant compared to the WT receptor activity. This further validated that the IL1 and IL2 are not involved in receptor signaling but are important determinants of maintaining the receptor integrity. It is possible that this diminished G protein activity is most likely caused by the misfolding of the receptor

(chimeras) which is evidenced by altered PM trafficking. This suggests that the interactions between the ILs and TM regions have a prominent role in stabilizing the receptor structure which ultimately affects the chromophore as well. Also, our results are in agreement with literature reports, which identified conserved residues in IL2 (including E134, V138, and

C140) which were reported to have distinct roles which may be important for maintaining the receptor structure by forming intramolecular interactions between ILs and TM domains

(182).

However, since any signaling switch could not be achieved or detected from the

BO-CXCR4-IL3 chimeras as both BO and CXCR4 interact with the same G protein type

131

Gαi, we next moved to the BO-β2AR chimera development. Since a crystal structure is available only for β2AR but not for blue opsin we had to consider the sequence homology between the two receptors to determine the regions to be swapped without affecting the membrane integrity. By applying the results obtained from the BO-CXCR4 chimeras, we started replacing 2 residues at a time to the IL3 of blue opsin starting from the middle of the loop. With gradual replacements, the best IL3 chimeras obtained which showed both

PM trafficking and the Gs activity were further enhanced with the CT replacements. Upon gradual increment of the loop length, the maximum light-dependent Gs activity was observed with BO-β2AR chimera containing both 52 residue IL3 and whole CT replacement. Light dependent activation of that BO-β2AR-IL3 (52 AA)-CT chimera showed a 30% of the Gαs activity (which was measured by the mini Gs recruitment to the

PM), shown by β2AR with isoproterenol. That in vivo response is remarkable as a chimera and superior compared to the rhodopsin-based chimeras considering both signaling as well as distribution. Additionally, if the receptors are extracted and conducted experiments in vitro as reported in the literature (179), it may be possible to observe much greater activity.

Moreover, there may be receptor specific (i.e., β2AR) conformational changes involved in chromophore and/or G protein interacting/binding pocket formation, which could be a possible reason for these functional BO-β2AR chimeras to not to show 100% activity as their WT counterparts, and further fine-tuning of the sequence boundaries may enhance characteristic Gαs signaling. Overall, from this study we could obtain properly folded opsin-based β2AR chimeras containing IL3 and CT from β2AR which show clear light dependent Gαs signaling and the results obtained can serve as a tool which can be useful in studies of GPCR signaling in vivo.

132

5.4. MATERIALS AND METHODS:

5.4.1. Reagents:

The reagents; Isoproterenol (Cayman Chemicals), was initially dissolved in DI water and further diluted with HBSS during experiments. PtX (lyophilized powder)

(Sigma-Aldrich) was reconstituted with DI water. SDF-1α powder (PeproTech) was reconstituted in DI water to a concentration 100 µg/mL and further diluted to 10 μg/mL with a buffer containing 0.1% BSA before adding 5 μl to cells. 11-cis retinal (National Eye

Institute) was originally resuspended in absolute ethanol to make 50 mM retinal solution and 1 µl from that was introduced to cells in 1 mL media under dark conditions to obtain

50μM final retinal concentration.

5.4.2. DNA constructs and cell lines:

Engineering of DNA constructs used; GFP-Gγ9, mCh-Gγ9, and Akt-PH-Venus have been described previously (99, 110, 111). Blue opsin-mCh was a kind gift from

Professor. N. Gautam’s lab, Washington University, St. Louis. Mini Gs DNA was also a gift from Dr. Nevin Lambert from Augusta University, GA. Blue opsin-mCh enhanced version was generated by site directed mutagenesis of potential phosphorylation sites at the

CT of blue opsin. For the mutagenesis, the template blue opsin-mCh plasmid was PCR amplified with overhangs containing expected nucleotide mutations, DpnI (NEB) digested

(to remove the parent construct) and performed DNA ligation. To generate the chimeric receptors also, the template DNA (i.e., enhanced-blue opsin) was amplified with primers containing the desired IL sequence containing overhangs. After the DpnI digestion, Gibson assembly (NEB) was performed to the PCR product according to the manufacturer’s instructions (112).

133

Cell lines (HeLa and RAW 264.7 cells) were originally purchased from the

American Tissue Culture Collections (ATCC) and authenticated using a commercial kit to amplify 9 unique STR loci.

5.4.3. Cell culture and transfections:

HeLa cell line was cultured in minimum essential medium (MEM; from CellGro) supplemented with 10% dialyzed fetal bovine serum (DFBS; from Atlanta Biologicals) and

1% Penicillin−Streptomycin (PS) in 60 mm tissue culture dishes. RAW 264.7 macrophages

(PIP3 generation and cell migration experiments) were cultured in Roswell Park Memorial

Institute (RPMI) 1640 (Corning, Manassas, VA) with 10% DFBS, and 1% PS supplementation.

When the cells reach ~ 80% confluency, media in the in the tissue culture dish were aspirated, versene-EDTA (CellGro) added and incubated in the 37 0C incubator for 3 mins.

After the incubation, cells were lifted from the dish and centrifuged for 3 mins at 290 rpm.

Next, versene was aspirated and the cell pellet was resuspended in their growth medium at a cell density of 1×106 /mL. For imaging experiments, cells were seeded on 35 mm glass- bottomed dishes (1×105 cells on each) with 15 mm inner diameter.

Before cell seeding, dishes were incubated with 2 N NaOH for 30 mins, ethanol washed, and sterilized in a UV transilluminator for 1 hr. The following day of cell seeding, cells were transfected with appropriate DNA combinations using Lipofectamine 2000 transfection reagent (Invitrogen), according to the manufacturer’s protocol and stored in a

0 37 C, 5% CO2 incubator. Cells were imaged the following day of transfection (after 16 hours of the transfection).

134

5.4.4. Live cell imaging to monitor Gβγ translocation, mini Gs translocation, cell migration, and PIP3 generation:

A spinning-disk XD confocal TIRF (total internal reflection) imaging system was used for live cell imaging experiments. The imaging system contains a Nikon Ti-R/B inverted microscope, a Yokogawa CSU-X1 spinning disk unit (5000 rpm), an Andor

FRAP-PA (fluorescence recovery after photo-bleaching and photo-activation) module, a laser combiner (with 40−100 mW four solid-state lasers; 445, 488, 515, and 594 nm wavelengths), and an iXon ULTRA 897BV back-illuminated deep-cooled EMCCD camera. In all experiments (Gβγ translocation, mini Gs translocation, cell migration, and

PIP3 generation), imaging was performed using a 60X, 1.4 NA (numerical aperture) oil objective.

In Gβγ9 translocation experiments, mCherry or GFP fluorescent tag on Gγ9s were imaged for 10 mins in every 1 s interval, using 594 nm excitation−630 nm emission and

488 nm excitation−515 nm emission, respectively. In cell migration experiments, mCherry tagged receptor blue opsin was imaged using 594 nm excitation−630 nm emission. To activate blue opsin, 50 µM 11-cis retinal was added and incubated 3-5 min in dark. After incubation, the fluorescent sensor in cells (i.e., blue opsin-mCherry) was imaged to capture basal signaling in cell migration, and then receptor blue opsin was activated by shinning

445 nm blue light every 2 s with 0.1% transmittance on a selected region of interest in one side of the cell, and mCherry imaging was continued for 20 mins. In PIP3 generation experiments, Venus tagged PIP3 sensor; Akt-PH, was imaged using 515 nm excitation−

528 nm emission yellow laser.

135

5.4.5. Statistical data analysis:

Results of all quantitative assays (Gβγ9 translocation, mini Gs translocation, PIP3 generation, cell migration) are expressed as standard error of mean (SEM) from n numbers of cells (indicated in the figure legends) from multiple independent experiments.

136

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Appendix A

I. II. III. Gβ1-mCh Gβ1-mCh GFP - Gγ9 Gβ1 -mCh GFP - Gγ3

photobleach photobleach photobleach

Fig. A-1. Gβ forms Gβγ dimers with overexpressed Gγ types. Half of the HeLa cells which are expressing I) Gβ1-mCherry alone, II) Gβ1-mCherry with GFP-Gγ9, III) Gβ1-mCherry with GFP-Gγ3 were exposed to high energy 488 nm green and 594 nm red light to photobleach GFP and mCherry, respectively, after baseline imaging for 20 s. The recovery of both fluorescent proteins was captured for 10 mins in every 1 second interval. Plots show similar t1/2 values for FRAP of Gβ1 to Gγ3 and Gγ9 recovery t1/2 values, respectively, indicating that Gβ1 forms Gβγ dimers with overexpressed Gγ types (scale: 5 μm, error bars: SEM, n=10).

161

Appendix B

Table B-1: Translocation and PIP3 generation properties of Gγ mutants

Gγ type Sequence t1/2 (s) Rate of PIP3 -1 generation (s )

Gγ3 WT NPFREKKFFCALL 290 ± 7 0.00069±0.00004 Gγ3-FF→GG NPFREKKGGCALL 92 ± 8 0.00042±0.00003 Gγ3-FF shifted NPFFFREKKCALL 68 ± 5 0.00032±0.00001 Gγ3-KK→GG in FF shifted NPFFFREGGCALL 47 ± 5 0.00025±0.00002 Gγ3-F65→G NPGREKKFFCALL 142 ± 9 0.00056±0.00003 Gγ3-RE→GGGG NPFGGGGKKFFCALL 204 ± 5 0.00067±0.00004 Gγ9 WT NPFKEKGGCLIS 37 ± 4 0.00025±0.00003 Gγ9-GG→FF NPFKEKFFCLIS 169 ± 7 0.00038±0.00003 Gγ9-KEK→GGG NPFGGGGGCLIS 25 ± 3 0.00023±0.00003 Gγ9 withGγ3 pre-CaaX NPFREKKFFCLIS 233 ± 8 0.00068±0.00003

162

Appendix C

CXCR4 ---MEGISIYTSDNYTEEMGSGDYDSMKEPCFREENANFNKIFLPTIYSIIFLTGIVGNG Rhodopsin MNGTEGPNFYVPFSNKTGVVRSPFEA---PQYYLAEP-WQFSMLAAYMFLLIMLGFPINF BO MRKMSEEEFYL-FKNISSV--GPWDG---PQYHIAPV-WAFYLQAAFMGTVFLIGFPLNA . .:* . : . ::. * : : : .: ::: *: *

CXCR4 LVILVMGYQKKLRSMTDKYRLHLSVADLLFVI--TLPFWAVDAVANWYFGNFLCKAVHVI Rhodopsin LTLYVTVQHKKLRTPLNYILLNLAVADLFMVFGGFTTTLYTSLHGYFVFGPTGCNLEGFF BO MVLVATLRYKKLRQPLNYILVNVSFGGFLLCIFSVFPVFVASCNGYFVFGRHVCALEGFL :.: . **** : ::::...::: : . .. . : ** * .:

CXCR4 YTVNLYSSVLILAFISLDRYLAIVHATNSQRPRKLLAEKVVYVGV---WIPALLLTIPDF Rhodopsin ATLGGEIALWSLVVLAIERYVVVCKPMSNFR----FGENHAIMGVAFTWVMALACAAPPL BO GTVAGLVTGWSLAFLAFERYIVICKPFGNFR----FSSKHALTVVLATWTIGIGVSIPPF *: : *..::::**:.: :. .. * :..: . * * .: : * :

CXCR4 I------FANVSEADDRYICDRFYPNDLWVVVFQFQHIMVGLILPGIVILSCYCIIIS Rhodopsin VGWSRYIPEGMQCSCGIDYYTPHEETNNESFVIYM----FVVHFIIPLIVIFFCYGQLVF BO FGWSRFIPEGLQCSCGPDWYTVGTKYRSESYTWFL----FIFCFIVPLSLICFSYTQLLR . : * . * * .: :. : ::. :*:* :* .* ::

CXCR4 KLSHSKGHQK------RKALKTTVILILAFFACWLPYYIGISIDSFILLEIIKQGCE Rhodopsin TVKEAAAQQQESATTQKAEKEVTRMVIIMVIAFLICWLPY------AGVAFYIFTHQGSD BO ALKAVAAQQQESATTQKAEREVSRMVVVMVGSFCVCYVPY------AAFAMYMVNNRNHG :. .:*: .:. . .:::: :* *::** .. : . :..

CXCR4 FENTVHKWISITEALAFFHCCLNPILYAFLGAKFKTSAQHALTSVSRGSSLKILSKGKRG Rhodopsin FG---PIFMTIPAFFAKTSAVYNPVIYIMMNKQFRNCMVTTLCCGKNPLGDDE------BO LD---LRLVTIPSFFSKSASIYNPIIYCFMNKQFQACIM-KMVCGKAMTDEADTCAAQKT : ::*. :: . **::* ::. :*. . : . . .

CXCR4 G----HSSVSTESESSSFHSS Rhodopsin -----ASTTVSKTETSQVAPA BO EVATVAATQVGPNETAQVAPA :: .*::.. .:

Fig. C-1. Sequence alignment showing the homology between rhodopsin, blue opsin, and CXCR4.

163