engineering synthetic control over rho gtpases using ca ... · engineering synthetic control over...

Engineering Synthetic Control Over Rho GTPases Using Ca2+ and Calmodulin Signaling

by

Evan Mills

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Institute of Biomaterials and Biomedical Engineering University of Toronto

© Copyright by Evan Mills 2012

ii

Engineering Synthetic Control Over Rho GTPases

Using Ca2+ and Calmodulin Signaling

Evan Mills

Doctor of Philosophy

Institute of Biomaterials and Biomedical Engineering University of Toronto

2012

Abstract

Engineered protein systems have been created to impart new functions, or “re-program”

mammalian cells for applications including cancer and HIV/AIDS therapies. The successful

development of mammalian cells for re-programming will depend on having well-defined,

modular systems. Migration is a particularly important cell function that will determine the

efficiency and efficacy of many re-programming applications in vivo, and Rho proteins are

responsible for regulation of cell migration natively. While there have been several reports of

photo-activated Rho proteins, no strategy has been developed such that Rho proteins and cell

migration can be controlled by a variety of extracellular stimuli that may be compatible with

signaling in large organisms. Here, several methods are described for engineering Ca2+-sensitive

Rho proteins so that the large, natural toolbox of Ca2+-mobilizing proteins can use the Ca2+

intermediate to activate Rho proteins in response to a variety of exogenous stimuli, including

chemicals, growth factors, and light.

First, an unreported calmodulin binding site was identified in RhoA. This knowledge was used

to create a tandem fusion of RhoA and calmodulin that mediated Ca2+-sensitive bleb retraction in

response to a variety of Ca2+-elevating chemicals. Ca2+-mobilizing modules including

iii

channelrhodopsin-2 and nicotinic acetylcholine receptor α4 were used for light- and

acetylcholine-dependent bleb retraction.

Second, a more robust morphology switch was created by embedding a calmodulin binding site

into RhoA to enable Ca2+-responsive bleb formation. A wider range of Ca2+-mobilizing modules

were also used here including LOVS1K/Orai1 and vascular endothelial growth factor 2.

Combining Ca2+-mobilizing and Ca2+-responsive modules increased amoeboid-like cell

migration in wound closure and transwell assays.

Finally, the embedded peptide design was applied to Rac1 and Cdc42 to enable control of new

morphologies and migration modes. The modular Ca2+ control over Rho proteins developed here

is an important contribution to cell re-programming because it shows that control over cell

migration can be rewired in a way that is flexible and tunable.

iv

Acknowledgments

My supervisor, Dr. Kevin Truong,

for being an incredibly supportive, inspiring and creative teacher and mentor.

My lab mates and colleagues, especially Elizabeth Pham and Seema Nagaraj,

for being there to guide, listen and help without asking anything in return.

My supervisory committee,

for helping me to focus, suggesting new ideas, and challenging me.

My family and friends, especially my parents,

for years of support and encouragement.

My wife, Jessica,

for giving me goals, a broader vision and balance.

v

Table of Contents

Abstract.......................................................................................................................................... ii

Acknowledgments ........................................................................................................................ iv

Table of Contents .......................................................................................................................... v

List of Publications ...................................................................................................................... ix

List of Abbreviations .................................................................................................................... x

List of Figures............................................................................................................................... xi

List of Movies ............................................................................................................................. xiii

1 Introduction .............................................................................................................................. 1

1.1 Motivation.......................................................................................................................... 1

1.2 Specific Research Objectives ........................................................................................... 2

1.3 Organization ...................................................................................................................... 2

2 Background............................................................................................................................... 4

2.1 Re-programming Mammalian Cells ............................................................................... 4

2.1.1 Engineering Genetic and Protein Networks for Therapeutic Applications .... 5

2.1.2 Optogenetics to Control Cellular Behaviour ...................................................... 6

2.2 Rho GTPases in Cell Migration....................................................................................... 8

2.2.1 The Principal Rho Proteins RhoA, Rac1 and Cdc42......................................... 9

2.2.2 The Role of Rho Proteins in Cell Migration ..................................................... 11

2.2.3 Tools for Studying Rho Proteins and Cell Migration...................................... 12

2.3 Ca2+ Signaling.................................................................................................................. 14

2.3.1 Ca2+-mobilizing Proteins .................................................................................... 15

2.3.2 Ca2+-sensitive Proteins ........................................................................................ 17

2.3.3 Tools for Sensing Ca2+ in Living Cells .............................................................. 18

3 Experimental Procedures ......................................................................................................20

vi

3.1 Gene Construction .......................................................................................................... 20

3.1.1 Subcloning Reagents and Materials .................................................................. 20

3.1.2 Generating a Cassette Vector from PCR.......................................................... 22

3.1.3 Combining Two Cassettes .................................................................................. 24

3.1.4 Removing the Fluorescent Protein Tag............................................................. 26

3.1.5 Performing Point Mutations .............................................................................. 26

3.2 In Vitro Protein Analysis................................................................................................ 27

3.2.1 Protein Production and Extraction ................................................................... 27

3.2.2 Pull-down Assays and SDS-PAGE .................................................................... 28

3.2.3 Fluorescence Spectroscopy for FRET measurements ..................................... 30

3.3 Cell Culture and Live-cell Fluorescence Imaging........................................................ 30

3.3.1 Cell Culture Reagents, Stimulatory Reagents and Equipment ...................... 30

3.3.2 Cell Culture Protocols ........................................................................................ 31

3.3.3 Fluorescence Microscope and Accessories........................................................ 32

3.3.4 Time-lapse Imaging Experiments...................................................................... 33

3.4 Migration Assays............................................................................................................. 33

3.4.1 Scratch Wound Closure Assays ......................................................................... 33

3.4.2 Transwell Migration Assays............................................................................... 35

3.5 Data Analysis ................................................................................................................... 36

3.5.1 Statistical Comparisons ...................................................................................... 37

3.5.2 Fluorescence Signal Colocalization ................................................................... 37

3.5.3 Area Calculations................................................................................................ 37

4 CaM-RhoA Fusion Protein.................................................................................................... 38

4.1 Chapter Aims and Motivation ....................................................................................... 38

4.2 Results .............................................................................................................................. 39

4.2.1 A CaM Binding Site in RhoA............................................................................. 39

vii

4.2.2 Design of a Ca2+-dependent RhoA-based Morphology Switch ....................... 41

4.2.3 Ca2+-dependent Bleb Retraction by CaM-RhoA(DP)-YFP ............................ 43

4.2.4 Light- and ACh-dependent Bleb Retraction .................................................... 47

4.3 Discussion......................................................................................................................... 49

4.4 Chapter Summary and Conclusion ............................................................................... 50

5 CaRQ: A New Ca2+-Sensitive RhoA Chimera..................................................................... 52


5.2 Results .............................................................................................................................. 53

5.2.1 Rational Design of CaRQ, a Ca2+-Sensitive RhoA Chimera ........................... 53

5.2.2 Characterization of CaRQ ................................................................................. 56

5.2.3 The Effect of Ca2+-mobilizing Modules and Spatial Localization on CaRQ . 61

5.2.4 The Effect of Temporally Distinct Ca2+ Signals on CaRQ Activation ........... 65

5.2.5 Regulation of Cell Migration by CaRQ ............................................................ 67

5.3 Discussion......................................................................................................................... 71


6 RACer: A Ca2+-Sensitive Rac1 Chimera ............................................................................. 77


6.2 Results .............................................................................................................................. 78

6.2.1 Design of RACer, a Ca2+-Sensitive Rac1 Chimera........................................... 79

6.2.2 Characterization of RACer ................................................................................ 80

6.2.3 The Effect of Ca2+-mobilizing Modules on RACer Activation ....................... 86

6.2.4 Regulation of Cell Migration by RACer ........................................................... 88

6.2.5 Design and Characterization of a Ca2+-sensitive Cdc42.................................. 92

6.3 Discussion......................................................................................................................... 94


7 Summary and Conclusion ................................................................................................... 100

viii

8 Future Directions ................................................................................................................. 102

8.1 Further Development of Cell Re-programming......................................................... 102

8.2 Refining Artificial Ca2+ Signaling Networks .............................................................. 105

8.3 Studying Rho Proteins and Cell Migration ................................................................ 107

9 References ............................................................................................................................. 109

Appendices................................................................................................................................. 120

Appendix A: Gene and Protein Sequences ........................................................................ 121

Appendix B: List of Oligonucleotide Primers and Sequences ......................................... 127

Appendix C: Supplemental Movie Legends ...................................................................... 128

Appendix D: Additional Control Experiments for CaM-RhoA(DP) .............................. 130

Appendix E: Further Colocalization Analysis of RBD/PBD Probes ............................... 132

Appendix F: Behaviour of CaRQ in HeLa and CHO Cells ............................................. 133

Appendix G: Mutations in Rac1 and Cdc42 to Introduce a CaM Binding Site............. 135

Appendix H: Ca2+ Control Designs for Rac1 and Cdc42 with PBD................................ 137

Appendix I: Area Thresholding Analysis Applied to Blebbing Cells.............................. 139

ix

List of Publications

1. Mills E, Pham E, Nagaraj S and Truong K. Engineered networks of natural and synthetic

proteins to control cell migration. ACS Synth Biol. Accepted, in press.

2. Mills E, Chen X, Pham E, Wong S and Truong K (2012). Engineering a photo-activated

caspase-7 for rapid induction of apoptosis. ACS Synth Biol. 1(3): 75-82.

3. Wong S, Kotera I, Mills E, Suzuki H and Truong K. (2012) Split-intein mediated re-assembly

of genetically encoded Ca2+ indicators. Cell Calcium. 51(1): 57-64.

4. Mills E and Truong K. (2011) Ca2+-mediated synthetic biosystems offer protein design

versatility, signal specificity and pathway re-wiring. Chem Biol. 18(12): 1611-1619.

5. Pham E, Mills E and Truong K. (2011) A synthetic photo-activated protein to generate local

or global Ca2+ signals. Chem Biol. 18(7): 880-890.

6. Mills E, Pham E and Truong K. (2010) Structure-based design of a Ca2+ sensitive RhoA

protein that controls cell morphology. Cell Calcium. 48(4): 195-201.

7. Li IT, Mills E and Truong K. (2010) A Computational Tool for Monte Carlo Simulations of

Biomolecular Reaction Networks Modeled on Physical Principles. IEEE Trans Nanobiosci. 9(1):

24-30.

8. Mills E and Truong K. (2010) Engineering Ca2+/calmodulin-mediated modulation of protein

translocation by overlapping binding and signaling peptide sequences. Cell Calcium. 47: 369-

377.

9. Mills E and Truong K. (2009) Cascading signaling pathways improve the fidelity of a

stochastically and deterministically simulated molecular RS latch. BMC Syst Biol. 3(1): 72.

10. Mills E and Truong K. (2009) Rate and extent of protein localization is controlled by

peptide-binding domain association kinetics and morphology. Protein Sci. 18(6): 1252-60.

x

List of Abbreviations ATP Adenosine 5’-triphosphate ACh Acetylcholine CaM Calmodulin CaRM Ca2+-activated Rho protein with embedded MLCKp CaRQ Ca2+-activated Rho protein with embedded IQp CaM Calmodulin CBP Calmodulin binding peptide CDZ Calmidazolium CFP Cyan fluorescent protein CKKp Calmodulin binding peptide from Ca2+-CaM kinase kinase ChR2 Channelrhodopsin-2 DMEM Dulbecco’s modified Eagle’s medium ER Endoplasmic reticulum FBS Fetal bovine serum FMN Flavin mononucleotide FRET Förster resonance energy transfer GAP GTPase accelerating protein GEF Guanosine nucleotide exchange factor GDI Guanosine dissociation inhibitor GDP Guanosine 5’-diphosphate GST Glutathion-s-transferase GTP Guanosine 5’-triphosphate IP3 Inositol 1,4,5-triphosphate IP3R Inositol 1,4,5-triphosphate receptor IQp Calmodulin binding peptide from myosin Va MLCKp Calmodulin binding peptide from myosin light chain kinase mRFP Monomeric red fluorescent protein nAChR-α4 Nicotinic acetylcholine receptor, α4 subunit PAK p21-activated kinase PAV Parvalbumin PLC Phospholipase-C PBD p21-binding domain from p21-activated kinase PBS Phosphate-buffered saline PM Plasma membrane RACer Rac1 activated by Ca2+ chimera RBD RhoA-binding domain ROCK Rho kinase Tg Thapsigargin TnC Troponin C TRPC Transient receptor potential channel SOCE Store-operated Ca2+ entry VEGF-A Vascular endothelial growth factor A VEGFR2 Vascular endothelial growth factor receptor 2 YFP Yellow fluorescent protein

xi

List of Figures

Figure 2.1: Schematic of a simplified Rho nucleotide exchange cycle. ......................................... 8

Figure 2.2: Simplified protein interaction network for the three principal Rho GTPases. ............. 9

Figure 2.3: Role of Rho GTPases in two modes of cell migration. .............................................. 11

Figure 2.4: Schematic of a typical cycle of events in a Ca2+ signaling pathway. ......................... 14

Figure 2.5: Cartoon of Ca2+-mobilizing proteins used in this work.............................................. 15

Figure 2.6: Overview of FRET principles in dual fluorophore Ca2+ sensors. .............................. 19

Figure 3.1: Overview of the pCfvtx vector. .................................................................................. 21

Figure 3.2: Flowchart of the three procedures for generating a cassette-based gene library........ 25

Figure 3.3: Two ways to combine cassettes A and B to create A-B............................................. 26

Figure 3.4: Cartoon depiction of wound assays analysis. ............................................................. 35

Figure 3.5: Cartoon depiction of transwell migration analysis..................................................... 36

Figure 4.1: Overview of fusion proteins used in Chapter 4. ......................................................... 39

Figure 4.2: Identification of a CaM binding site in RhoA............................................................ 40

Figure 4.3: RhoA-dependent morphologies.................................................................................. 41

Figure 4.4: Structural modeling of the CaM-RhoA(DP) fusion protein. ...................................... 43

Figure 4.5: Ca2+ transients elicited by ATP or UTP in CHO, HeLa and HEK293 cells. ............. 44

Figure 4.6: Ca2+-induced bleb retraction in CHO, HeLa and HEK293 cells................................ 46

Figure 4.7: Summary of bleb retraction in CHO cells. ................................................................. 47

Figure 4.8: Light- and ACh-mediate bleb retraction.................................................................... 48

Figure 5.1: Overview of main fusion proteins created in Chapter 5............................................. 53

Figure 5.2: Modeling of CaRQ. .................................................................................................... 55

Figure 5.3: Colocalization analysis of CaRQ and RBD. .............................................................. 56

Figure 5.4: Stimulation of CaRQ in HEK293 cells. ..................................................................... 58

xii

Figure 5.5: Determination of the Ca2+ EC50 value for pLyn-CaRQ. ............................................ 60

Figure 5.6: A caspase-3/7 sensor uncleaved in stimulated CaRQ-expressing cells. .................... 60

Figure 5.7: Activation of CaRQ and pLyn-CaRQ by ACh/nAChR-α4. ....................................... 62

Figure 5.8: Activation of pLyn-CaRQ and CaRQ by ATP and blue light....................................63

Figure 5.9: Localization of CaRQ to the ER with Stim1.............................................................. 64

Figure 5.10: Blebbing duration relative to duration of Ca2+ signals............................................. 65

Figure 5.11: Effect of PAV on blebbing duration......................................................................... 66

Figure 5.12: Light-dependent migration by CaRQ using wound assays. ..................................... 67

Figure 5.13: VEGFR2 is a Ca2+-mobilizing mode that activated CaRQ. ..................................... 69

Figure 5.14: Multi-signal migration of CaRQ cells using transwell assays. ................................ 70

Figure 6.1: Overview of main fusion proteins created in Chapter 6............................................. 78

Figure 6.2: Design of RACer. ....................................................................................................... 80

Figure 6.3: Pre-stimulation effects of RACer. .............................................................................. 81

Figure 6.4: Effect of ionomycin on cells expressing RACer. ....................................................... 84

Figure 6.5: The Ca2+ EC50 of RACer was 24 µM. ........................................................................ 84

Figure 6.6: Effect of Ca2+ stimulation of RACer in various cell lines.......................................... 85

Figure 6.7: RACer can be activated by three distinct Ca2+-mobilizing modules.......................... 87

Figure 6.8: Light-dependent migration by RACer using wound assays. ...................................... 89

Figure 6.9: Multi-signal migration of RACer cells using transwell assays. ................................. 91

Figure 6.10: Characterization of CaM-Cdc42 chimera................................................................. 93

Figure 7.1: Overview of selected artificial Ca2+ signaling networks developed here.................101

xiii

List of Movies

Movie 4.1: CaM-RhoA(DP)-mediated bleb retraction in CHO cells in response to ATP

Movie 4.2: CaM-RhoA(DP)-mediated bleb retraction in HeLa cells in response to UTP

Movie 4.3: CaM-RhoA(DP)-mediated bleb retraction in HEK293 cells in response to ionomycin

Movie 5.1: CaRQ-mediated blebbing in HEK293 cells in response to ionomycin

Movie 5.2: CaRM-mediated bleb retraction in HEK293 cells in response to ionomycin

Movie 5.3: YFP control cells in response to ionomycin

Movie 5.4: pLyn-CaRQ/nAChR-α4-mediated blebbing in HEK293 cells in response to ACh

Movie 5.5: pLyn-CaRQ(T19N)/nAChR-α4 cells stimulated with ACh

Movie 5.6: Stim1-CaRQ cells blebbing in response to engineered SOCE

Movie 6.1: RACer-mdiated lamellipodia formation in response to ionomycin

Movie 6.2: RACer(T17N) cells stimulated with ionomycin

Movie 6.3: RACer/LOVS1K/Orai1 cells stimulated by illumination

Movie 6.4: RACer(T19N)/LOVS1K/Orai1 cells stimulated by illumination

Movie 6.5: Ca2+ stimulation of CaM-Cdc42 chimera by ionomycin

Movie 6.6: Ca2+ stimulation of CaM-Cdc42(T17N) chimera by ionomycin

1

1 Introduction

This chapter provides the reader with the broad motivation for the work and specific research

goals are introduced. An organizational overview of the document is given.

1.1 Motivation

Engineered protein and genetic systems have been created to impart new functions, or “re-

program” mammalian cells to facilitate cancer immunotherapies (1-2), HIV treatment (3-4), and

as tools for basic scientific research (5-7). While these reports demonstrate interesting proof-of-

concept systems, the widespread use of re-programmed mammalian cells for therapeutic

applications will require the modular combination of genetic and protein parts that can carry out

well defined functions such as cell migration, programmed cell death, protein secretion or gene

transfer between cells, both in vitro and in vivo.

Cell migration is a particularly important cell function that will determine the efficiency and

efficacy of many re-programming applications involving the delivery of a population of cells to a

particular location in vivo. A robust system to regulate cell migration should be capable of

responding to a variety of physical and chemical cues, depending on the requirements of a

particular application. Synthetic proteins to regulate cell migration that have been reported have

been designed to respond to only one signal, such as blue light for PARac (5). However, should

migration be required in response to some other stimulus such as red light or a chemical signal,

PARac would not be a suitable functional module for the new application. There is a growing

kernel of work in engineering the cell migration-regulating Rho GTPase family, which provides

additional motivation for working in this area.

One cellular pathway that can be re-configured to respond to different chemical and physical

cues is the Ca2+ signaling pathway (8). Ca2+ is a versatile second messenger that regulates many

physiological pathways through Ca2+-sensitive proteins and can be regulated by a large “tool-

box” of Ca2+-mobilizing proteins. This tool-box of Ca2+-mobilizing proteins can control the

cytoplasmic Ca2+ concentration in time and space in response to a variety of chemical cues

including neurotransmitters (9) and growth factors (10), and physical signals such as light (11-

12) and voltage (13). The Ca2+-sensitive protein calmodulin (CaM) represents a good starting

2

point for designing Ca2+-sensitive chimeras because it has been well characterized (14) and

widely used in the design of Ca2+ biosensors (15-18).

1.2 Specific Research Objectives

The overall goal is to develop a Ca2+-based system where exogenous input signals can regulate

cell migration through Rho GTPases. Given the motivation for the work, this thesis is thereby

objective-oriented. To achieve the overall objective, three specific research objectives were

identified:

1. Create chimeras where Rho-associated morphologies are switched by changes in the

intracellular Ca2+ concentration.

2. Construct artificial Ca2+-signaling networks through combinations of Ca2+-mobilizing

proteins and Ca2+-sensitive chimeras developed in (1) so that exogenous signals, such as

vascular endothelial growth factor (VEGF-A) or blue light, can control cell morphology.

3. Determine whether the artificial Ca2+-signaling networks developed in (2) can be used to

regulate cell migration in response to exogenous signals.

1.3 Organization

This thesis consists of nine chapters organized so as to demonstrate the progression of design

strategies and ideas that were employed to engineer Ca2+-sensitive Rho proteins. The first

chapter motivates the work as a contribution to the field of cellular re-programming and

discusses specific research objectives such as regulating cell migration using versatile input

signals. The second chapter gives the reader background information in three relevant areas: cell

re-programming, the role of Rho proteins in cell migration and the Ca2+ second messenger as the

foundation for a complex signaling network. The third chapter describes the experimental

techniques used in this work and is comprised of five sections: gene construction, in vitro protein

analysis, live-cell fluorescence imaging, cell migration assays and data analysis.

Chapters four, five and six describe three chimeric proteins that were constructed, characterized,

and tested for their ability to regulate cell migration. Chapter four describes a tandem fusion

protein of CaM and RhoA, CaM-RhoA, which mediated Ca2+-dependent bleb retraction.

Chapter five describes a novel chimera where RhoA was fused with CaM and two CaM-binding

3

peptides to create CaRQ, a more robust Ca2+-dependent switch of RhoA-associated

morphologies. Chapter six describes the design strategy of chapter five applied to the closely

related Rho protein Rac1 to create RACer. Chapters five and six include assays that demonstrate

that CaRQ and RACer successfully regulated cell migration. Throughout, insights into the

modularity and spatiotemporal fidelity of Ca2+ signaling were used to modify the chimeras and

their behaviour.

The final chapters summarize the document and body of work, and suggest reasonable future

directions. Chapter seven recapitulates the contributions described and chapter eight discusses

several ways to expand upon the work both using these specific chimeras and generally using

insights gained from the work. Chapter nine is a list of references, and is followed by relevant

additional information in several appendices.

4

2 Background

This chapter will review pertinent background related to the objective of developing a modular

system for regulating Rho GTPases with Ca2+ signaling. First, the idea of cellular re-

programming will be explored by reviewing studies that report engineered proteins and cells

designed to ameliorate a disease state. Photo-activatable proteins will also be discussed as useful

laboratory tools to study re-programming of different pathways, including manipulating Ca2+

signals and Rho proteins. In the second section, Rho proteins and their role in cell migration will

be reviewed. The focus will be on the three principal Rho proteins RhoA, Rac1 and Cdc42 and

their regulators and effectors. Models for two modes of cell migration, mesenchymal and

blebbing-based amoeboid-like will be summarized from the literature. Key tools and techniques

to study Rho proteins will be introduced, with an emphasis on those used in this study. Last,

selected aspects of Ca2+ signaling networks will be discussed. In particular, Ca2+-mobilizing

proteins and Ca2+-sensitive proteins used in this study will be described, with some discussion of

how the cell uses networks of proteins to appropriately interpret Ca2+ signals. Tools used to

study Ca2+ signaling in living cells will be introduced, with emphasis on the ones used here.

2.1 Re-programming Mammalian Cells

Imparting novel functions to mammalian cells, or “re-programming” mammalian cells by

delivering engineered transgenes is being pursued in experimental therapeutic strategies (1-4)

and to create tools for basic research (5-7). This is a broad area of investigation that combines

protein engineering, work in synthetic biology and cell-based therapies. If research in this field

is successful, it will ultimately lead to a view of the cell as a platform from which therapeutic

strategies can be deployed, similar to current views of biomaterials or nanoparticles (19-20). The

word “re-programming” in this context should be delineated from the concept of cell type re-

programming where adult cells are de-differentiated to induced pluripotent stem cells by the

Yamanaka factors (21) or are directly transferred to some other adult cell type (22). Similarly,

“re-programming” should be distinguished from conventional gene therapy where corrected

copies of defective genes are delivered to cells (23). In both of the latter cases (cell type re-

programming and gene therapy) the goal of research is to alter a population of cells to a different,

but naturally occurring, population, whereas cellular re-programming as the term is used here

seeks to introduce novel, useful functions to cells.

5

This section will briefly review several recent studies where engineered or synthetic transgenes

were delivered to cells, in vivo or in vitro, to endow the cells with novel functionalities. The

discussion is separated into two groups: first, where the aim was to use cells as a therapeutic

platform, and second, where light-sensitive tools were developed for laboratory research.

2.1.1 Engineering Genetic and Protein Networks for Therapeutic Applications

Three recent studies have successfully demonstrated the ability to deliver engineered or synthetic

transgenes to re-program mammalian cells into therapeutic platforms.

Engineered oncolytic viruses re-program cancer cells to initiate an immune response: Oncolytic

viruses based on the vaccinia virus have been created that preferentially infect certain cancer cell

types. In the case of JX-594, the virus has been engineered to proliferate only in cells with

elevated EGFR/Ras signaling by deleting the viral thymidine kinase and virally-encoded growth

factors. Additionally, the virus has been engineered to include a trangene for GM-CSF, a

neutrophil chemoattractant and macrophage maturation factor. Cancerous cells, which do not

normally secrete GM-CSF, were re-programmed to do so constitutively, upregulating immune

response and improving tumour suppression in vivo (1). This study demonstrated that delivery

of transgenes in vivo in humans by a suitable vector could be a viable therapeutic strategy.

Chimeric antigen receptors re-program T-cells: Chimeric antigen receptors are engineered

transmembrane proteins comprised of an extracellular antigen-binding domain and the

intracellular CD3-ζ chain, which is sufficient to activate T-cell receptor signaling transduction

pathways. In particular, engineered CART19 cells have an extracellular anti-CD19 antibody,

fused to CD3-ζ intracellularly, to target B-cell lymphoma, CD19 being an accepted antigen for

B-cell malignancies (2). In this instance of cell re-programming, T-cells were removed from

patients by apheresis, modified ex vivo and re-injected resulting in disease remission in 2 of 3

trial patients. This study is contrasted with the previous one in a number of ways: cells were

transfected (or infected) ex vivo rather than in vivo, the re-programmed cells were not the targets

of the therapy and the engineered protein was a novel chimera. Conceptually, however both

studies fit within the frame of cellular re-programming because a population of cells was

endowed with a specific, useful functionality that does not naturally occur for that cell type.

6

Engineered transgenes re-program lymphoid cells for HIV resistance: HIV resistance has been

achieved in different cell populations by disrupting the CCR5 gene, which encodes a co-receptor

for many strains of HIV. In CD4+ T-cells, engineered zinc fingers were used to disrupt the

CCR5 gene; when re-programmed cells were delivered to HIV-infected mice, the mice

developed lower viral loads and higher numbers of circulating T-cells (3). CCR5 has also been

suppressed by shRNA in CD34+ cells, also in a mouse model, where the HIV infection rate in a

CCR5-disrupted cell population was significantly less than in control cells (4). These studies

show that re-programming cells may become an effective therapeutic strategy for HIV infection;

the treatment of other diseases may also benefit from cellular re-programming.

2.1.2 Optogenetics to Control Cellular Behaviour

Optogenetics refers to a class of laboratory tools where light is used as a stimulus to control an

engineered device (usually a synthetic protein) to alter cellular physiology (24). When applied to

large organisms, light-based or photo-activatable tools suffer from light’s poor tissue penetration

due to absorption and scattering (24). However, photo-activatable proteins are convenient

laboratory tools because light can be easily controlled in time and space, and insights gained into

a system’s behaviour can be applied to design control mechanisms that may be more relevant for

large organisms. Native light-sensing protein domains are genetically combined with effector

proteins in such a way that conformational changes in the light-sensing domain regulate the

activity of the effector protein: LOV domains (5, 12, 25-26), channelrhodopsins (11, 27-30) and

the phytochrome-PIF system (6) have been used to build optogenetic tools.

The LOV2 domain from A. Sativa phototropin-1 (31-32) is a light-sensitive domain that has been

used to construct optogenetic tools (5, 12, 25-26). LOV2 associates with a flavin

mononucleotide (FMN) chromophore in the dark and incident photons with frequencies in the

blue light range excite FMN to form a transient covalent bond with Cys450 (33). The FMN-

Cys450 bond perturbs the LOV2 structure resulting in undocking of the Jα helix, a peptide that is

carboxy-terminal to LOV2 in phototropin-1. The thermal relaxation of the FMN-Cys450 bond

has an approximate half-life of 30-60s, so after illumination ceases, the Jα helix will re-dock with

the LOV2 domain (34). Three optogenetic tools created with LOV2 are discussed in relation to

potential re-programming applications.

7

• LovTAP is a photoactivatable transcription factor for the trp repressor from E. Coli (25). In

LovTAP LOV2 is an allosteric inhibitor of trp in the dark; inhibition is released by Jα

undocking resulting in a 65-fold increase in DNA binding (25). While LovTAP was

designed to investigate allosteric regulation, and later optimized (26), the ability to inhibit

gene expression by light is intriguing. If the mechanism of LovTAP were applied to a

promoter, repeated illumination could potentially drive gene expression in many applications.

• PARac is a photoactivatable Rho GTPase (Rac1) that controls lamellipodia formation and

cell migration (5). In PARac, LOV2 “cages” Rac1 activity by hydrogen bonding between

LOV2 and F37/W56 on Rac1 (5). Illumination releases the caging and sustained

illumination of PARac results in local lamellipodia formation and directed migration. One

could imagine directing engineered cells by light toward sites of injury or disease in vivo as a

potential application of PARac or similar systems.

• LOVS1K is a photoactivatable switch that regulates Ca2+ influx (12). LOVS1K is a tandem

fusion of LOV2, residues 233-450 from Stim1, which is an endogenous protein that monitors

the ER Ca2+ concentration (35), and mRFP. Undocking of the Jα helix from LOV2 in

LOVS1K presumably allows the Stim1 fragment to associate with Orai1 (12), a PM

transmembrane Ca2+ channel (35). Repeated activation results in a steadily increasing Ca2+

concentration via influx through Orai1. The modular nature of Ca2+ signaling means that

LOVS1K, in combination with other engineered proteins, could be used for light-based

control of many pathways (discussed in detail below in section 2.3).

Channelrhodopsins and the phytochrome-PIF system have also been used to create

photoactivatable proteins. Channelrhodopsin-2 (ChR2) is a light-gated ion channel where

permeability is increased when the associated all-trans-retinal is photoisomerized to 13-cis-

retinal, leading to a conformational change in the seven transmembrane channels (36). ChR2 has

been engineered for increased permeability, kinetics and sensitivity (29-30), and to “re-wire”

illumination to activation of other second messengers such as IP3 or cAMP (28). A naturally

occurring red-shifted variant, VChR1, also exists (27). Artificial neuronal pacing with ChR2

could enable re-programmed behaviour or learning at the organism level. With phytochrome-

PIF, light-regulated cell morphology and migration was achieved by a different mechanism than

PARac. Phytochromes are naturally occurring red-light sensors from A. Thaliana where incident

light (650 nm) photoisomerizes a phycobilin chromophore covalently bound to a phytochrome

8

protein (6). Photoisomerization increases the affinity of the phytochrome-phycobilin complex

for PIF proteins, creating photoactivatable interdomain recruitment. This was used to achieve

light-activated translocation of guanine nucleotide exchange factors, which activate Rho

proteins, to the PM to regulate morphology and migration (6).

This project explores re-programming cells using Ca2+ signaling as a robust and modular way to

regulate cell migration via Rho GTPases. The re-programming capability of a Ca2+-based

signaling network will enable cells to migrate in response to diverse stimuli, representing a

conceptual improvement on Rho GTPase regulation by PARac and the phytochrome-PIF system.

2.2 Rho GTPases in Cell Migration

The Rho protein family is a group of some 20 GTPases within the larger Ras superfamily of

GTPases. The Rho family’s main role in the cell is as regulator of actin-myosin organization,

morphology and migration (37-39). Rho proteins are approximately 190 amino acid globular

domains with a carboxy-terminal tail bearing a CAAX prenylation motif. They share a common

regulation scheme (Figure 2.1). The exchange of guanosine 5’-diphosphate (GDP) for guanosine

5’-triphosphate (GTP) results in a conformational change in two regions known as Switch I

(residues 28-44) and Switch II (residues 62-70) (40). Conformational change in these regions

alters the binding surfaces of Rho proteins for various effector proteins, thus driving the overall

switching behaviour. The net effect of a Rho protein’s specific guanosine nucleotide exchange

factors (GEF’s), GTPase accelerating proteins (GAP’s) and guanosine dissociation inhibitors

(GDI’s) determine its nucleotide state, although additional regulation can be applied through

phosphorylation (41) and ubiquitination (42). Similarly, a Rho GTPase’s set of effector proteins

determines its particular effect on the cell.

Figure 2.1: Schematic of a simplified Rho nucleotide exchange cycle.

9

This section will discuss the three principal Rho proteins in more detail, focusing on their

regulation and effector proteins, describe models of two different modes of cell migration and

the role of Rho proteins in these models, and finally describe some common tools that are widely

used to study Rho proteins and migration.

2.2.1 The Principal Rho Proteins RhoA, Rac1 and Cdc42

While there are some 20 Rho proteins, three are studied in the most detail: RhoA, Rac1 and

Cdc42. Each has a distinct set of effector proteins and therefore distinct effects on cell

morphology and migration (Figure 2.2).

Figure 2.2: Simplified protein interaction network for the three principal Rho GTPases.

Input signals (left) are linked to cytoskeletal effects (right) by a simplified Rho network. Lines are shown for stimulatory (arrow) and inhibitory (bar) interactions. Question marks denote interactions that are suspected but not proven. Acronyms: TKR (tyrosine kinase receptor), LIMK (LIM kinase) and MLCP (myosin light chain phosphatase).

RhoA: The primary role of RhoA in terms of the cytoskeleton is to generate contractile forces.

RhoA is regulated by several GEF’s including Vav3, ArhGEF18 and LARG which link the Eph-

Ephrin system (43), G-protein coupled receptors such as lysophosphatidic acid receptor (44) and

tyrosine kinase receptors such as insulin growth factor receptor (45) to RhoA. RhoA, Rac1 and

Cdc42 share several common GAPs, such as p55RhoGAP and p110RhoGAP and common GDIs,

10

RhoGDI1-3 (46-47). RhoA generates contractile forces through binding to the effector Rho

kinase (ROCK) (48). GTP-bound RhoA binds to ROCK releasing an auto-inhibitory domain

that allows ROCK to phosphorylate myosin light chain (MLC) (49), resulting in cross-linking

and contraction. Through ROCK, RhoA also regulates LIM kinase, which inactivates the actin

depolymerization factor cofillin (50). GTP-bound RhoA also activates mDia, a formin that

promotes actin polymerization through association with actin monomers, profillin and the barbed

end of polymeric F-actin (51). RhoA has also been linked to gene expression pathways STAT3

(52) and NF-κβ (53), possibly through activation of PKN1 (54). Together, RhoA effectors enable

RhoA to mediate cytoskeletal contraction and stress fibre formation.

Rac1: The main role for Rac1 in cell migration is the development of lamellipodia at the leading

edge. Rac1 and Cdc42 share similar GEF’s including Tiam1, Vav1 and Intersectin (55-57).

These GEF’s function downstream of growth factor receptors such as platelet derived growth

factor receptor for Tiam1 (55) or integrins (β1 subunit for Tiam1 (57) and β2 for Vav1 (56))

which link the extracellular environment to migration. The role of Rac1 in increasing actin

polymerization and lamellipodia formation at the leading edge is primarily achieved through its

activation of the WAVE complex and subsequent regulation of Arp2/3 (58). In this way

activation of Rac1 directs formation of new branched F-actin structures and an extension of the

cytoskeleton. Rac1 also is known to exert control over RhoA and Cdc42: direct binding to the

GEF DBS allows Rac1 to activate Cdc42 (59-60) while a variety of mechanisms may be

involved in Rac1 antagonism of RhoA including activation of the RhoA-specific GAP

p190RhoGAP (61). Rac1 and Cdc42 both activate p21-activated kinase (PAK) (62) which likely

links them to regulation of the JNK/MAPK pathway (63).

Cdc42: Cdc42’s contribution to cell migration is regulating the formation of filopodia at the

leading edge. Active Cdc42 is directly incorporated into the N-WASP complex which regulates

the Arp2/3 actin polymerization complex (64). N-WASP and WAVE are related complexes and

differences in resultant actin polymerization patterns may be based on differences in their VCA

domains, the parts of the N-WASP and WAVE complexes that interact with actin monomers and

Arp2/3 (65-66). However, it is not currently known how Cdc42/N-WASP is able to produce

straight actin bundles characteristic of filopodia via Arp2/3, which produces branched actin.

Given that formins produce linear actin polymers, some interaction between Cdc42, or its

11

effectors, and formins such as an mDia homologue is suspected (67). Cdc42 is able to activate

Rac1, probably through PAK which is known to activate a Rac1 GEF called αPIX (68).

2.2.2 The Role of Rho Proteins in Cell Migration

Two starkly different modes of cell migration exist: mesenchymal (48, 58, 64, 69-70) and

blebbing-based amoeboid-like (71-76), likely along a spectrum containing aspects of each mode.

The role of each Rho protein in cell migration is dependent on the particular mode of cell

migration, or, perhaps, the relative activities of Rho proteins determine the mode of cell

migration (Figure 2.3). For brevity, the discussion will be limited to the contribution of the

principal Rho proteins to current models.

Figure 2.3: Role of Rho GTPases in two modes of cell migration.

The top row shows a simplified model of mesenchymal cell migration including filopodia and lamellipodia formation and rear-cell contraction. The bottom row shows a model of blebbing-based cell migration where RhoA/ROCK induces blebbing and cytosolic flow propels the cell. Black circles denote transmembrane adhesion receptors linking the cell to the matrix. Rho GTPase activation is colour-coded: RhoA (red), Rac1 (blue) and Cdc42 (green).

Mesenchymal cell migration: Mesenchymal, lamellipodia-based, or fibroblast-like migration

involves the spatiotemporal coordination of Rho proteins and the cytoskeleton on an adhesive

surface. In this model Cdc42 activity at the leading edge directs straight actin bundles in

structures known as filopodia (64) or invadopodia (77). Subsequently Rac1/WAVE initiate

12

branched actin polymerization in a flat, protrusive sheet to create lamellipodia (58). Adhesion

receptors connected to the growing actin network (integrins, cadherins, etc) anchor the cell’s

forward motion (70). Simultaneously, RhoA/ROCK activity in the cell rear (48, 69) initiates

actin-myosin crosslinking providing the contractile force to dislodge focal adhesions and “pull”

the cell forward. This model has recently been supported by several elegant studies where Rho-

family biosensors and photoactivatable proteins were combined to elucidate the dynamic system

(5, 78). This model is complicated by the contribution of several “other” Rho proteins such as

RhoC which interacts with different formins than RhoA leading to distinguishable morphologies

and migration patterns (79), the observation that RhoA is cyclically activated in a narrow band at

the front of protrustions (41, 78), and the appearance of morphological structures that cannot be

classified as lamellipodia or filopodia, such as blebs (see below).

Blebbing-based amoeboid-like cell migration: Also known as actinomyosin contractility-based

migration, blebbing-based amoeboid-like migration is distinguished from other types of

amoeboid-like migration such as the actin polymerization-dependent protrusive migration of

leukocytes (80). Blebbing-based migration has been demonstrated in model organisms (81-82),

and in invasive cancer cell migration (71, 74, 76). In this mode, contractile forces generated by

RhoA/ROCK (75, 83) cause local detachment of the PM from the cytoskeleton known as a bleb.

As the bleb expands, cytosol rushes to fill the new space causing the cell to “run” forward (82).

As a bleb ages (1-2 minutes), actin polymerizes inside the bleb re-linking the PM with the

cytoskeleton (84). This has two effects: first, it enables transmembrane adhesion receptors such

as E-cadherin to anchor the bleb to the extracellular matrix or neighbouring cells, allowing the

cell to realize forward movement (81), and second, retracts the bleb (84). A subsequent bleb in

an area near the first one will allow the cell to continue to move in the same direction. This

mode of cell migration tends to occur in low adhesion contexts, however the causal relationship

between low adhesion and amoeboid-like migration is unknown. The role of Rac1 and Cdc42 in

bleb formation is unclear, although in some blebbing models, such as SH4 domain

overexpression, dominant negative Rac1 and Cdc42 did not interfere with blebbing (75).

2.2.3 Tools for Studying Rho Proteins and Cell Migration

Elucidating the role of Rho proteins in living cells has benefited from several biochemical

techniques including fusions with fluorescent proteins, often used in a colocalization or “pull-

13

down” assay (85), siRNA (79, 86), Förster resonance energy transfer (FRET)-based biosensors

(69, 87-88), optogenetic tools (5-7) and a variety of migration assays (71, 74, 76, 79, 86, 89-92).

Overexpression of dominant negative and positive mutants have been used to gain insight into

Rho proteins: The T17N mutation (T19N for RhoA) renders Rho proteins constitutively inactive

(93) by disrupting binding to a necessary Mg2+ cofactor and the Q61L mutation (Q63L for

RhoA) diminishes GAP activity preventing GTP hydrolysis (94). Distinguishing between GTP-

and GDP-bound wild type Rho proteins in living cells can be done by observing association

between Rho proteins and high-affinity binding domains: the Rho-binding domain (RBD) from

rhotekin for RhoA (95) or the p21-binding domain (PBD) from PAK for Rac1 and Cdc42 (96).

siRNA has been particularly useful in distinguishing the roles of closely related Rho family

proteins such RhoA and RhoC (79). FRET biosensors have been created for each of the

principal Rho proteins (69, 87-88), and have been used to uncover the spatiotemporal

relationships between the key Rho proteins in migrating cells. FRET biosensors are discussed in

more detail in relation to monitoring Ca2+ concentrations below (section 2.3.3).

Optogenetic tools: Photoactivatable proteins such as PARac and others (5-7) have also been

applied to study Rho protein networks and cell migration. Combining photoactivatable proteins

with FRET-based biosensors has allowed for insights into Rho protein regulation to go beyond

noticing correlations to establishing cause and effect relationships. For example, it was shown

that activation of Rac1 results in a striking de-activation of RhoA in time and space around the

localized Rac1 activity (5, 69). PARac has also been used in a D. Melanogaster development

model to show that Rac1 activity in a few cells of a coherent cluster can “direct” the migration of

other cells through cell-cell adhesion and possibly through the secretion of migratory cues (7).

Migration assays: Migration is one of the most physiologically relevant phenotypes of Rho

protein function, and is clinically relevant in terms of metastatic cell migration. Commonly used

techniques for assessing cell migration include scratch wound closure assays (90-92), transwell

assays (71, 79, 89) and 3D invasion assays (71, 74, 76, 86).

• A scratch wound closure assay typically involves a confluent monolayer of cells that has

been disrupted by a straight scratch or “wound” and the rate of cell migration into the open

area is measured. Wound assays are typically used to assess the effect of surface conditions,

protein expression (or suppression) or chemical inhibitors on migration rate. This does not

14

require complex analysis (wound closure is measured by counting cells (91), changes in

wound edge (92), or area (90)), but have the obvious limitations of a 2D environment without

the ability to present migratory cues with any spatial control.

• In a transwell assay, cells are seeded on the apical side of a porous filter and can migrate

either on top of or through the filter. A chemoattractant of some kind is usually placed in the

basal chamber, and the rate at which cells migrate “toward” the chemoattractant is measured

(71, 76, 89). The transwell assay overcomes some limitations of a wound assay by

presenting a chemical concentration gradient, although the gradient cannot be maintained

indefinitely. Still, the 3D environment is simplistic compared to the in vivo cellular milieu.

• An invasion assay is an extension of a transwell assay where a thick gel, often collagen or

Matrigel, is laid down between the seeded cells and the porous filter. The rate at which cells

can “invade” or migrate through the gel is measured, often as a function of matrix

metalloproteinase activity, gel composition or some other cellular property (71, 74, 76, 86).

Invasion assays are often carried out in studies involving cancerous cells.

2.3 Ca2+ Signaling

Ca2+ is a ubiquitous second messenger that has been implicated in the regulation of many cellular

functions from excitation-contraction coupling to gene expression to apoptosis and necrosis (8,

97-98). Ca2+ regulates cellular functions through a three step cycle: first, a stimulus activates

some Ca2+-moblizing protein or pathway that increases the Ca2+ concentration in some part of

the cell for some time, second, a Ca2+-sensitive protein in the region of increased Ca2+ senses the

Ca2+ influx and actuates some physiological process, and third, Ca2+-removal processes are

activated to return the Ca2+ concentration to its resting level (Figure 2.4).

Figure 2.4: Schematic of a typical cycle of events in a Ca2+ signaling pathway.

15

While there is a large toolbox of natural and engineered Ca2+-mobilizing proteins, specific ones

which have been used here will be described. Similarly there are many Ca2+-sensitive proteins

that decode changes in Ca2+ concentration into physiological signals; some putative mechanisms

for this, and specific Ca2+-sensitive proteins used will be discussed. Mechanisms for Ca2+-

removal will not be discussed because they have not been actively modified during this work.

2.3.1 Ca2+-mobilizing Proteins

The specific Ca2+-mobilizing proteins and pathways that have been used in this study can be

grouped into three classes: endogenous, where a cell already contains the protein machinery to

translate a chemical signal into a Ca2+ signal; exogenous, where a particular protein machinery is

introduced to a cell from a specialized cell-type via transgene expression; and synthetic, where

an engineered protein is introduced as a transgene to mediate a Ca2+ signal (Figure 2.5).

Figure 2.5: Cartoon of Ca2+-mobilizing proteins used in this work.

ChR2, nAChR-α4 and P2X channels (light blue) mobilize Ca2+ directly from the extracellular space when their stimuli are present (blue wave for photons, red dots for ACh and green dots for ATP). P2Y and VEGFR2 receptors (orange) initiate a signaling cascade and Ca2+ influx is mediated by IP3R from the ER stores when ATP or VEGFA is present (light blue dots). LOVS1K (dark blue) associates with Orai channels (light blue) when excited by light, leading to Ca2+ influx from the extracellular space.

Endogenous Ca2+-mobilizing proteins: Two chemicals are often used to elicit Ca2+ responses

based on endogenous signaling mechanisms: adenosine 5’-triphosphate (ATP) and ionomycin.

ATP has several possible mechanisms of action depending on the endogenous receptors that are

present. P2X receptors are two membrane-pass ligand-gated cation channels; ATP binding to the

extracellular domain causes a conformational change and Na+ and Ca2+ entry (99). P2Y

16

receptors are seven membrane-pass G-protein coupled receptors; ATP binding causes the Gαq

subunit to activate phospholipase-Cβ (PLCβ) (100) leading to inositol 1,4,5-triphosphate (IP3)

production and Ca2+ influx from the ER through IP3 receptor (IP3R) (101) . Depletion of ER

lumenal Ca2+ stores may then result in store-operated calcium entry (SOCE) via transient

receptor potential channels (TRPC) (102) or through Stim and Orai (35). Depending on the

particular P2X or P2Y subunits present, a given cell may also be activated by other purines such

as uridine 5’-triphosphate (UTP) (99). Ionomycin is also thought to elevate intracellular Ca2+

levels through a SOCE mechanism (103).

Exogenous Ca2+-mobilizing proteins: Three Ca2+ -mobilizing proteins were introduced via

transgenes to enable cell lines used here to respond to exogenous stimuli: nicotinic acetylcholine

(ACh) receptor (nAChR-α4), VEGF receptor (VEGFR2) and ChR2.

• nAChR is a ligand-gated non-specific cation channel with permeability for Na+, K+ and Ca2+:

maximum permeability in the absence of nicotine is achieved with a 1 mM concentration of

ACh (9). In nAChR there are five subunits comprised of some combination of nine α

subunits and three β subunits: the α4 subunit is typically present in the α4β2 nAChR, but

assemblies of nAChR-α4 without any β subunits are functional cation channels (9).

• VEGFR2, also known as KDR or FLK1, is a single-pass transmembrane receptor with seven

Ig-like extracellular domains and an intracellular tyrosine kinase. VEGF-A binding to

VEGFR2 (at concentrations of 5 to 10 ng/mL) results in dimerization and

autophosphorylation of the tyrosine kinase domain (104). There are several downstream

effects of VEGFR2 ligation including MAPK/ERK and Akt1 activation; in the context of

Ca2+ signaling, VEGFR2 ligation activates PLCγ leading to IP3R mediated Ca2+ release from

the ER, and possibly subsequent SOCE (10).

• ChR2 is a ligand-gated proton channel from with some permeability for Ca2+ (discussed

above in section 2.1.2). ChR2 has been used to modulate Ca2+ signals in neurons (29), pace

transgenic cardiomyocytes (105), and activate engineered proteins in cell lines (12).

Synthetic Ca2+-mobilizing proteins: LOVS1K is a synthetic Ca2+-mobilizing protein that

translates blue light (~440 nm) into local and global Ca2+ signals (discussed further in section

2.1.2). Repeated illumination (300 ms every 10-15s) steadily increases cytoplasmic Ca2+:

17

initially the Ca2+ signal is small and only detectable near the PM however with repeated

illumination over several hours, global increases in Ca2+ spread throughout the cytoplasm (12).

2.3.2 Ca2+-sensitive Proteins

A diverse group of proteins are known to be directly affected by Ca2+ including CaM, troponin C

(TnC), phosphatases such as calcineurin and kinases such as protein kinase C, and many others

indirectly by binding to or being enzymatically modified by Ca2+-sensitive proteins. Ca2+ does

not act as a simple switch in all cases: a range of affinities for Ca2+ and requirements for

cofactors, both of which are involved in regulating different behaviours within, for example, the

calpain family (106), provide some complexity to Ca2+ signaling.

Complexity can also be derived from the frequency and time-varying shape of a Ca2+ signal

(107), the modular nature of the protein components in a given environment (8) and spatial

patterning (13, 108-110). The modular nature of Ca2+ signaling pathways is particularly relevant

in this work. Ca2+-sensitive proteins likely do not discriminate between different mechanisms

for mobilizing Ca2+, provided that the spatiotemporal pattern is indistinguishable, and similarly

Ca2+-mobilizing processes operate independently from the downstream Ca2+-sensitive processes

in the absence of feedback. As an example, Ca2+ release from ryanodine receptors and IP3R

mediate contraction via TnC in cardiomyocytes (111) but they enable long term depression and

potentiation in neurons with CaM-dependent kinase II (112). Spatial patterning of Ca2+ signals

is also known to regulate complex physiological responses. Migrating fibroblasts have been

shown to use local micrometer-sized “Ca2+ flickers” to alter their directionality on 2D surfaces

(108). Voltage-regulated Ca2+ channels, themselves regulated by Ca2+ to some extent, can

respond to both the local Ca2+ concentration near the channel and the global cytoplasmic Ca2+

concentration through an associated CaM molecule (13). Buffering proteins such as

parvalbumin (PAV) are thought to help massage the spatiotemporal pattern of a given Ca2+

signal (109). The cell has evolved many ways of achieving complexity using Ca2+, and a clear

understanding of these mechanisms is needed to design protein systems using Ca2+.

Calmodulin and its binding peptides: CaM is an evolutionarily conserved, ubiquitous 150 amino

acid protein with a dumbbell shape. CaM was chosen for this study because it has a large library

of potential binding target peptides, and the structures of CaM bound to many of these peptides

are known. CaM has four EF-hand domains, two in each lobe of the dumbbell and therefore four

18

Ca2+ binding sites, each with a different affinity for Ca2+: the apparent overall affinity is 1-10 µM

(113). Unloaded, or apoCaM, adopts a closed conformation: the EF-hand domains are nearly

parallel to each other obscuring the hydrophobic core of CaM to potential binding partners.

After Ca2+ loading, Ca2+-CaM is in an open conformation: the EF-hands snap open and are

nearly perpendicular to each other, exposing hydrophobic binding sites (14).

CaM converts changes in Ca2+ concentration into physiological actions by binding other proteins

and altering their activity. The location of CaM binding on a target protein, or CaM binding site,

is usually a 20-30 amino acid region with a high propensity for α-helix formation (14, 114).

Specific hydrophobic anchoring residues in the target peptide interact with the exposed

hydrophobic core of Ca2+-CaM in one of several common patterns such as 1-10 or 1-5-8-14,

where the numbers indicate the relative positions of hydrophobic anchors on the target peptide

(14, 114-115). A class of peptides which can bind apoCaM and Ca2+-CaM are the IQ motifs

(116), so-called because they frequently have Ile-Gln (I-Q) residues.

CaM’s ubiquitous role in the cell is related in part to its promiscuous nature: there are hundreds

of known binding targets for CaM on proteins that control processes as diverse as

musculoskeletal contraction and gene expression (115). The CaM target peptide from myosin

light chain kinase (MLCKp) binds to Ca2+-CaM with very high affinity (low nanomolar range)

while there is little affinity between apoCaM and MLCKp (117). Similarly, the CaM target

peptide from Ca2+-CaM kinase kinase (CKKp) has low nanomolar affinity for Ca2+-CaM and

poor affinity for apoCaM, but the CKKp fragment binds CaM in the reverse orientation from

MLCKp (118). In contrast, the IQ motif peptide from myosin Va (IQp) has a low micromolar

affinity for both apoCaM and Ca2+-CaM (119). The ability to access a large library of binding

partners means that engineered proteins based on CaM can be easily refined and tuned for a

range of affinities and behaviours.

2.3.3 Tools for Sensing Ca2+ in Living Cells

The prevalence of Ca2+ signaling in regulating cellular physiology has led to the creation of a

number of tools for studying Ca2+ in living cells. Sensing Ca2+ can be done by measuring Ca2+

indirectly with chloride channel currents (120), more direct measurements using Ca2+-sensitive

dyes such as Fluo-4 (intensity-based) (121) and Fura-2 (ratiometric) (122) or protein-based Ca2+

sensors. Dyes respond quickly to Ca2+ changes and are commercially available in a range of

19

affinities. However, dye toxicity, photobleaching and the difficulty targeting dyes to subcellular

locations are major disadvantages.

Protein-based Ca2+ sensors report on Ca2+ concentrations by changes in the intensity of one or

two fluorophores. Localization-based Ca2+ sensors exist (123-125), however these rely on

inherent cellular trafficking mechanisms to report on Ca2+ and are difficult to quantify. Single

fluorophore Ca2+ sensors such as GCaMP (16) or R-GECO (18) have relatively large dynamic

ranges, can be modified for different Ca2+ affinities (126), and do not require complex optical

setups. Dual fluorophore Ca2+ sensors such as cameleons (15, 17) or TN-XL (127) rely on the

principle of FRET to form an intensity ratio between a FRET donor, typically cyan fluorescent

protein (CFP) and a FRET acceptor, typically yellow fluorescent protein (YFP) (Figure 2.6).

While these sensors have a more limited dynamic range than single fluorophore sensors, they are

less sensitive to artifacts such as changes in cell shape or pH.

Figure 2.6: Overview of FRET principles in dual fluorophore Ca2+ sensors.

A. Overlap between CFP (donor) emission and YFP (acceptor) excitation spectra are required for FRET. B. Jablonski diagram representing the principles of energy transfer between donor and acceptor. Up-pointing arrows represent excitation and down-pointing arrows are for emission, wavy arrows show vibrational energy dissipation, and dashed arrows denote non-radiative processes during energy transfer between donor and acceptor. C. Schematic of a FRET-based protein biosensor in a low FRET-efficiency conformation (top) and a higher FRET efficiency conformation (bottom).

CaM, several of its binding peptides and RhoA will be the initial targets for developing Ca2+

control over Rho GTPases in this work. Ca2+ sensing will primarily be done with the TN-XL

biosensor because it does not introduce the possibility for cross-talk between the Ca2+ sensor and

the engineered Rho chimera. Insights gained into the synthetic regulation of RhoA will later be

applied to Rac1 and Cdc42.

20

3 Experimental Procedures

This chapter will describe the experimental procedures used in this thesis in five sections. First,

gene construction methods will be described including the complete procedures involved in

creating a cassette-based gene library and making point mutations. Second, procedures used for

in vitro protein analysis will be given including fluorescence spectroscopy and pull-down assays

that were used. Third, methods and protocols for cell culture and live-cell fluorescence

microscopy will be provided. Fourth, two migration assays that were used will be described:

wound closure assays and transwell migration assay. Last, several data analysis techniques used

throughout will be discussed including methods for statistical comparisons, colocalization

analysis and cell area analysis.

3.1 Gene Construction

Gene construction in this work is based on using a regular, self-propagating cassette structure

(128). The cassette structure minimizes the number of restriction enzymes that have to be used

during cloning and because most cloning actions regenerate the original cassette structure, genes

can be combined many times to create large, complex fusion products. The cloning vector

contains nested promoters (CMV for mammalian expression and T7 for bacterial expression) so

the same proteins used in live-cell imaging can also be used as markers during the cloning

process to identify successful clones. This section will describe the materials and methods

involved in generating a library using the cassette system described, including generating new

cassettes by inserting genes via PCR, combining existing cassettes and removing fluorescent

protein tags. Point mutations were used extensively in this work and the process for generating

point mutations used here will also be described. Specific genes that were created by cassette

cloning will be discussed in subsequent chapters as they are introduced. Relevant protein and

gene sequences, and oligonucleotide primer sequences are presented in the appendices

(Appendix A and Appendix B, respectively).

3.1.1 Subcloning Reagents and Materials

The vector used as the basis for the cassette cloning method is called pCfvtx (128) and is based

on the pTriex-3 backbone from Novagen (now EMD Chemicals, Missssauga, ON). Ampicillin

resistance enables antibiotic selection of pCfvtx in bacteria; there is no resistance gene for drug

21

selection in mammalian cells. The MCS region of pTriex-3 has been altered (Figure 3.1). Three

pairs of restriction enzyme sites generating compatible cohesive ends flank a stop codon (2a sites

on the 5’ side and 2b sites on the 3’ side) between the Kozak consensus sequence and the YFP

gene: SpeI/NheI, BamHI/BglII and StuI/SmaI (although this last pair is not used due to a

methylation site). Insertions into cassette are made using a pair of non-compatible sites on either

side of the first stop codon, such as BamHI and NheI (section 3.1.2). Combination of existing

cassettes can be done easily by making use of the compatible cohesive end pairs (section 3.1.3)

and the YFP gene can be removed by PmeI digestion (section 3.1.4).

Figure 3.1: Overview of the pCfvtx vector.

The schematic magnifies the MCS region downstream of CMV and T7 promoters. Restriction enzyme sites are written out including 2a sites (SpeI and BamHI) and 2b sites (BglII and NheI). Pairs of sites that generate compatible cohesive ends are marked by the same dashed bars.

Reagents for PCR: PCR was performed in 50 µL reactions containing 5 µL of 5 mM dNTP’s, 5

µL 10x PCR buffer with MgSO4, 5 µL of each forward and reverse primers diluted to 25 pM, 1

µL of cDNA or plasmid, 0.5 µL Pfu polymerase (all preceding from Fermentas, Burlington, ON)

and 28.5 µL RNase/DNase-free water (Invitrogen, Oakville, ON). The PCR reaction was

performed using an Eppendorf MasterCylcer Personal.

Reagents for enzyme digestion and ligation: Enzyme digestion was performed in 30 µL

reactions containing 3 µL 10x enzyme buffer (pre-mixed buffers supplied by the manufacturer),

1 µL 30x BSA, 1 µL of each restriction enzyme (usually two of NcoI, SpeI, BamHI, BglII, NheI

and XhoI) and 1 µg of DNA, up to a maximum of 24 µL, or 25 µL for a PmeI single enzyme

digest (all from NEB Canada, Pickering, ON). The remainder of 30 µL was filled with

RNase/DNase-free water. Ligation was performed in 20 µL reactions containing 2 µL 10x ligase

buffer (supplied by manufacturer), 1 µL T4 ligase (both from NEB) and a mixture of 17 µL DNA

22

containing insert and host in a 4:1 molar ratio. Typically this meant 10-12 µL of insert and 5-7

µL of host DNA solution.

Reagents for gel electrophoresis and DNA purification: Agarose gels were prepared with a 0.5x

TAE solution with 1% agarose (typically prepared in batches of 2 g UltraPure agarose per 200

mL TAE, both from Invitrogen). Prior to loading, DNA was mixed with 6x Orange Loading

Dye (i.e. 10 µL for PCR and 6 µL for enzyme digestion). The O’Gene DNA Ladder (100bp to

10kbp range) was used for mass comparisons (dye and ladder from Fermentas). Purification of

DNA fragments after gel electrophoresis was done with the PureLink Quick Gel Extraction kit

and purification of plasmids after bacterial culture was done with the PureLink Quick Plasmid

Miniprep kit (both from Invitrogen).

Reagents for transformation and plating and fluorescence screening equipment: Transformation

was done using DH5α Subcloning Efficiency chemically competent E. Coli cells stored at -80oC

in 20 µL aliquots to minimize freeze-thaw cycles. Bacterial colonies were grown on LB Agar

plates with 200 µg/mL ampicillin and bacteria were cultured in LB Broth prepared with distilled,

autoclaved water, with or without 100 µg/mL ampicillin depending on the procedure.

Kanamycin and streptomycin were occasionally used to culture E. Coli for plasmids from other

groups at 80 µg/mL and 100 µg/mL, respectively. Colonies were observed using the Illumatool

Tunable Lighting System using excitation filters (approximately 440 nm, 488 nm and 540 nm

peak wavelength transmission for CFP, YFP and mRFP excitation, respectively) and longpass

emission filters (480 nm, 525 nm and 560 nm, respectively).

3.1.2 Generating a Cassette Vector from PCR

The following is the procedure used to insert a gene amplified by PCR into the pCfvtx plasmid

cassette structure. The procedure, and similar procedures of combining cassettes (section 3.1.3)

and removing the fluorescent tag (section 3.1.4), are summarized in a flowchart (Figure 3.2).

1. Primers were designed to amplify the gene of interest, incorporating one restriction enzyme

from either side of the first pCfvtx stop codon into the forward and reverse overhangs (NcoI

and NheI, BamHI and NheI, and SpeI and BglII were common pairs).

2. A 50 µL PCR reaction was prepared as described above. A typical PCR scheme included 10

minutes at 95oC for initial denaturation. The cycling stage usually consisted of 32 cycles of 3

23

minutes at 95oC for denaturation, 3 minutes at 60oC for annealing and 2 minutes per 1 kbp to

be amplified at 72oC for extension. The final extension stage was 10-20 minutes at 72oC

depending on the length of the gene of interest.

3. The PCR product was mixed with loading dye, loaded onto agarose/TAE gels and run at

100V for 20-25 minutes in cold 0.5x TAE buffer. At least one lane per gel was also loaded

with DNA ladder. After electrophoresis, the gel was soaked in ethidium bromide solution for

20 minutes. After ethidium bromide soaking, gels were illuminated with a UV

transilluminator and bands of correct size and sufficient quality were excised. For poor

quality or absent bands, PCR parameters were adjusted and the reaction repeated up to 3

times. In the event of continued failure, primers were usually re-designed.

4. Gels were purified with the PureLink Quick Gel Extraction kit using the manufacturer’s

protocol.

5. Two enzyme digestions were prepared as described above, one for the pCfvtx host and

another for the PCR product. DNA concentrations were measured using the Eppendorf

Biophotometer and the “dsDNA” setting. Reactions were held at 37oC for 3 hours before a

second round of gel purification.

6. A ligation reaction of PCR product and host was prepared as described above. The reaction

was held at 16oC for 1 hour or up to 4 hours for difficult ligations. Troubleshooting at this

stage was common. The insert:host ratio, ligation time and reagent concentrations were

frequently adjusted to find a successful condition

7. Transformation of ligation products was done after E. Coli aliquots were thawed to 4oC on

ice. 1-2 µL of ligation reaction (more for difficult ligations) were mixed with the bacteria,

heat-shocked at 42oC for 45 seconds and then returned to ice for 5 minutes. After chilling, 1

mL of LB Broth was added and the mixture was incubated at 37oC for 2 hours.

8. After 2 hours the mixture was centrifuged for 1 minute at 13,200 rpm, 900 µL of supernatant

was removed and the pellet was re-suspended in the remaining supernatant. This mixture

was then loaded onto pre-warmed agar plates with ampicillin, spread out by gentle rocking,

and incubated at 37oC overnight.

9. The next day, colonies were screened for fluorescence using the Illumatool system and

appropriate filters (the pCfvtx plasmid has CFP and mRFP-bearing variants). If a successful

24

colony was found it was picked off the dish by a clean pipette tip and cultured in 2 mL LB

Broth supplemented with 100 µg/mL ampicillin overnight in a shaking 37oC incubator. If

colonies were not present or not properly fluorescent the procedure was re-started from

progressively earlier steps in the order of steps 6, 5, 2 then 1. Occasionally fluorescent

colonies would appear after 24-48 additional hours, presumably due to slow protein folding.

Additional incubation beyond 48 hours rarely yielded success.

10. Plasmid DNA was extracted from bacterial culture using the PureLink Quick Plasmid

Miniprep kit and manufacturer’s directions. The plasmid was then checked for quality by

enzyme digestion or sequencing (sequencing by TCAG, Toronto, ON).

3.1.3 Combining Two Cassettes

Combining two cassettes, cassette A and cassette B, to create A-B could be done in two ways

(Figure 3.3). Whether cassette A was treated as the host or insert depended on the relative

fluorescent protein tags present: generally fluorescent inserts were ligated with non-fluorescent

hosts. However, YFP-tagged inserts could be ligated with CFP- or mRFP-tagged hosts and CFP-

tagged proteins could be the host or insert for mRFP-tagged proteins, and the converse, limited

by poor spectral separation of YFP fluorescence by the filter sets used. Creating cassette B-A

can be done by selecting restriction enzymes in the opposite way shown in the figure below.

The procedures used to combine two cassettes were very similar to those presented above

(section 3.1.2). The differences in procedure are summarized below.

• The insert was digested with either NcoI and a site 2b enzyme, for example NheI, or XhoI

and a site 2a enzyme, for example SpeI. It follows that the host was digested with NcoI and

SpeI or XhoI and NheI, respectively (Figures 3.1 and 3.3).

• The overall procedure began with step 5 where the host was treated as pCfvtx in the previous

procedure and the insert was treated as the PCR product was.

• During gel electrophoresis there were two bands in the insert lane, one corresponding to the

insert and one to the insert’s vector. The insert’s vector was discarded. Similarly there may

have been a small fragment from the host vector to be discarded, although this was usually so

small that it ran off the gel during electrophoresis.

25

Figure 3.2: Flowchart of the three procedures for generating a cassette-based gene library.

The three main procedures (adding genes by PCR, cassette combination and fluorescent tag removal) are separated by dashed lines and share similar procedures at the end of the process. Decision points are shown in diamond shapes and procedures are rectangular blocks.

26

Figure 3.3: Two ways to combine cassettes A and B to create A-B.

The starting cassettes are shown on the top row, with only the MCS region shown. In the second row, A acts as the host and B as the insert, and the converse in the third row. Restriction enzyme sites that have been digested are shown by a vertical bar.

3.1.4 Removing the Fluorescent Protein Tag

Removing the PmeI-flanked fluorescent tag may be necessary to facilitate a cloning exercise or

because it is undesirable to express a fluorescent protein during an experiment. Again, many of

the procedures used in this process are similar to that of inserting a gene into cassette by PCR, so

only the differences in procedure are summarized below.

• The overall procedure began with step 5. Only one enzyme digestion was performed, on the

vector from which the fluorescent protein tag was to be removed. Only one restriction

enzyme was present, PmeI, so 1 µL extra of DNA-water solution was added.

• In step 6 the ligation was a “self-ligation”: 17 µL of purified digested DNA was used rather

than a mixture of two species of DNA. Exceeding 1 hour at 16oC was rarely needed.

• In step 9 a non-fluorescent colony was selected for culture. This means that it was

imperative to verify that the starting material was free of any non-fluorescent plasmids.

3.1.5 Performing Point Mutations

Point mutations were used extensively in this work as silent mutations to remove restriction

enzyme recognition sites or to introduce amino acid mutations (frequently for dominant negative

or positive mutations to Rho protein chimeras). Point mutations were introduced using overlap

extension PCR with self-hybridizing primers, as described below.

27

1. Assume that there were existing forward and reverse primers to insert the gene to be mutated

into cassette, and call them primers 1 and 3, respectively. Primer 2a was designed such that

it annealed approximately 10-12 base pairs to the 5’ and 3’ sides of the mutation site. The

primer should contain the desired mutant sequence; up to 3-4 consecutive base pairs could be

mutated with a single primer. The primer should have a melting temperature of

approximately 72oC, with equal contributions from the 5’ and 3’ sides of the mutation site.

Primer 2b should be designed as the reverse compliment of primer 2a.

2. Two separate PCR reactions were performed in parallel. In the first, primers 1 and 2b were

the forward and reverse primers, respectively, amplifying fragment F1. In the second,

primers 2a and 3 were the forward and reverse primers, respectively, amplifying fragment

F2. The quality of the PCR’s were checked by gel electrophoresis and purified.

3. A 40 µL PCR reaction was prepared with 5-10 µL F1, 5-10 µL F2, 5 µL dNTP’s, 5 µL buffer,

0.5 µL Pfu and filled with RNase/DNase-free water. This PCR reaction ran for 5 cycles.

4. Then, 5 µL primer 1 and 5 µL primer 3 were added, and this PCR reaction ran for ~30 cycles.

The PCR product was checked for quality and size by gel electrophoresis, then purified.

5. The next step was step 5 of inserting a new gene into cassette by PCR (section 3.1.2).

3.2 In Vitro Protein Analysis

Some protein constructs were analyzed in vitro, using two techniques: pull-down and subsequent

SDS-PAGE, and fluorescence spectroscopy. In each case proteins were cultured in E. Coli and

mechanically extracted before the analytical technique was performed. This section will

describe the protein production and extraction procedures, pull-down assays and SDS-PAGE,

and fluorescence spectroscopy measurements.

3.2.1 Protein Production and Extraction

Recombinant proteins were expressed in E. Coli using the T7 promoter from the pCfvtx

backbone and extracted by mechanical sonication lysis.

1. E. Coli transformed with a plasmid expressing the protein of interest were cultured for 24

hours in 4 mL of LB Broth supplemented with 100 µg/mL ampicillin in a 37oC shaking

incubator. This quantity would often produce approximately 250 µL of ~1 µM protein lysate,

28

judged qualitatively by comparing fluorescence intensity to a set of fluorescent protein

solutions with known concentrations. Scaling up was done using multiple 4 mL cultures.

2. Cells were harvested by centrifugation into a single pellet.

3. The pellet was washed three times in an aqueous solution of 50 mM Tris and 100 mM NaCl.

After the final wash the pellet was re-suspended in ~250 µL of the Tris/NaCl solution, or one

supplemented with MgCl2, CaCl2, EDTA or some other compound depending on the

application. Re-suspension volume was adjusted based on apparent fluorescence intensity.

4. The cell suspension was sonicated for 1 minute using a Branson Sonicator Model 250

(Thermo-Fisher Scientific, Ottawa, ON) with output intensity setting 3 and a 30% duty cycle.

5. After sonication, the suspension was pelleted, and the brightness of the supernatant was

compared to that of the remaining pellet. When the pellet was still much brighter than the

supernatant, an additional 1 minute of sonication was performed.

6. The suspension was centrifuged at 13,200 rpm for 5 minutes. The supernatant was

transferred to a clean 1.5 mL tube and centrifuged again for 5 minutes. The supernatant was

again transferred to a clean 1.5 mL tube and was then ready for downstream applications.

3.2.2 Pull-down Assays and SDS-PAGE

Pull-down assays were used to test for binding interactions between engineered proteins and

standard binding domains. Two types of pull-downs were used: CaM pull-down, and RBD/PBD

pull-downs via glutathione-s-transferase (GST). Fusion proteins were created to mediate the

RBD/PBD pull-downs: RBD-mRFP-GST and PBD-mRFP-GST, respectively. Here, RBD is

residues 7-89 from rhotekin (69, 85, 95) and PBD is residues 67-150 from PAK (85, 96).

For the CaM pull-down, CaM-sepharose beads were used (Stratagene, La Jolla, CA, USA).

1. 100 µL of bead slurry was removed from the stock per condition to be tested (approximately

50% of the slurry volume was beads).

2. The slurry was allowed to settle on the bench top in a 1.5 mL tube. The storage buffer was

removed by pipette and 500 µL of a 50 mM Tris, 100 mM NaCl and 1 mM CaCl2 solution

was added to wash the beads. The beads were allowed to settled, the wash buffer was

replaced, and this process was repeated twice more.

29

3. The final wash buffer was removed and 100 µL buffer of the same composition was added.

25-50 µL of fluorescent protein lysate was added (depending on fluorescence brightness) to

the mixture, and pipetted up and down to mix the lysate, buffer and beads.

4. The beads and proteins were allowed to settle and re-mixed frequently over 1 hour. Protein

translocation to beads was observed using the Illumatool system after beads were settled.

The beads were washed twice in the Tris/NaCl/CaCl2 solution to remove excess proteins.

5. The Tris/NaCl/CaCl2 solution was removed and replaced with Tris/NaCl/EDTA (2 mM

EDTA) to dissociate fluorescent proteins from CaM beads. Re-mixing and settling over 24

hours (kept at 4oC) was sometimes needed to fully dissociate protein from CaM beads. This

solution could then be used for downstream applications (e.g. SDS-PAGE).

For the RBD/PBD pull-down, commercially available immobilized glutathione agarose beads

were used (Novagen).

1. 100 µL of bead slurry was removed from the stock, washed and prepared as described above.

2. An excess of RBD-mRFP-GST or PBD-mRFP-GST was added to the beads, mixed and

allowed to settle. This was repeated several times over 1-2 hours to ensure maximum

loading of the RBD/PBD construct. Construct loading was confirmed by fluorescence.

3. Excess protein lysate was removed from the beads and fresh Tris/NaCl/MgCl2 buffer was

added (Mg2+ being necessary for Rho protein activation). 10 mM GTP was also added to the

buffer at this stage. The protein of interest was added to the beads and allowed to mix/settle

as described above, allowing up to 24 hours at 4oC for translocation to the beads.

SDS-PAGE was performed using pre-cast NuPage 1.5 mm thick 4%-12% Bis-Tris 10-well gels

and the recommended 1x SDS-MOPS buffer provided by the manufacturer (Invitrogen).

NuPage 1x LDS sample buffer and 1x reducing agent were also used. Solutions of 30 µL were

prepared according to the manufacturer’s directions and boiled at 70oC for 10 minutes prior to

loading. Electrophoresis was performed using an Owl EC105 power supply (Thermo-Fisher) at

120 V for 1-2 hours depending on desired separation. After electrophoresis, fluorescent proteins

were visualized using the Illumatool system. PageRuler protein ladder (Fermentas) was run

simultaneously to approximate protein mass. Gels were soaked overnight in Coomassie blue-

based PageBlue stain (Fermentas) to visualize the ladder, according to the manufacturer’s

instructions. Images were taken using a Canon Powershot A350 camera.

30

3.2.3 Fluorescence Spectroscopy for FRET measurements

Fluorescence spectra were collected with a UV/Vis/NIR fluorescence spectrometer model LS55

running the FL WinLab software (PerkinElmer, Woodbridge, ON). Protein lysate samples

(prepared as described above) from the same batch were loaded into UV/Vis cuvettes with either

1 mM CaCl2 or 1 mM EDTA for the “high” and “low” Ca2+ conditions, respectively. Spectra for

CFP-YFP based FRET were collected by exciting the sample at 440 nm and recording the

emission intensity from 460 nm – 560 nm. Spectra for a particular sample were recorded in

triplicate to smooth the signal. Each condition was tested with three independent samples.

3.3 Cell Culture and Live-cell Fluorescence Imaging

Live-cell fluorescence imaging was the primary technique for testing and characterizing chimeric

proteins. Several standard cell lines were used; cell culture reagents and protocols will be

described. The fluorescence microscope and accessory equipment used will also be described as

well as the general approach to time-lapse imaging experiments. Specific experiments that differ

from the general approach will be discussed in Chapters 4-6 as they are introduced.

3.3.1 Cell Culture Reagents, Stimulatory Reagents and Equipment

The main cell lines used in this work were HEK293, HeLa, CHO and COS7. RAW264.7 cells

were also used with some differences in protocol compared to the previous group of cells.

Cell culture and transfection reagents: Cells were cultured in DMEM supplemented with 25 mM

D-glucose, 1 mM sodium pyruvate, 4 mM L-glutamine and 10% FBS (Invitrogen). Culture

medium was also supplemented with 100 U/mL penicillin and 100 µg/mL streptomycin (Sigma,

St. Louis, MO, USA). Cells were passaged with 0.05% trypsin-EDTA (Sigma). Lipofectamine

2000 was used as the transfection reagent (Invitrogen). Pre-mixed cell freezing medium was

used containing DMSO and methylcellulose (Sigma). For RAW264.7 cells, RPMI-1640

medium (Invitrogen) was used, supplemented with 10 mM HEPES, 100 U/mL penicillin, 100

µg/mL streptomycin and 10% FBS. For imaging, cells were often submerged in PBS or PBS

with 1 mM CaCl2 and 1 mM MgCl2 (Sigma).

Cell culture materials and equipment: Cell cultures were maintained in screw-top T-25 flasks

(Sarstedt, Montreal, QC) in a HeraCell incubator maintained at 37oC and 5% CO2 (Thermo-

Fisher). All cell manipulations were performed in a Class IIA biosafety cabinet (Thermo-

31

Fisher). In preparation for imaging experiments, cells were passaged into 35 mm dishes with 14

mm glass bottom wells with No. 0 coverslip thickness (Mattek, Ashland, MA, USA).

Stimulatory reagents and inhibitors: Stimulatory reagents were added from 10x stocks such that

200 µL added to the 2 mL imaging buffer resulted in 1x concentration. Chemicals used during

imaging experiments are stated below at the 1x concentration. Chemicals prepared as aqueous

solutions were ATP and UTP (10 µM, Fermentas), VEGF-A (10 ng/mL, Cell Signaling

Technology, Pickering, ON), ACh (1 mM, Sigma), CaCl2 (1 mM, Sigma), EDTA (2 mM, Sigma)

and Y-27632 (10 µM, Sigma). Compounds dissolved in DMSO were prepared so that the final

concentration of DMSO in the cell culture was no more than 0.1%. Chemicals prepared in

DMSO were ionomycin (1 µM, Sigma), thapsigargin (2 µM, BioShop, Burlington, ON),

calmidazolium (50 µM, Sigma) and blebbistatin (10 µM, BioShop).

3.3.2 Cell Culture Protocols

Cell passaging procedures, especially seeding ratios, were modified depending on the particular

cell line in use, as described below.

1. When cells reached 95% confluency, medium was removed from cells by pipette. Cells were

washed in 5 mL PBS, adding PBS to the side wall, and rocked gently.

2. 1 mL trypsin was added to cells for 5 minutes at 37oC. Cells were loosened by tapping the

flask and transferring the suspension to a clean 1.5 mL tube. For RAW264.7 cells, cells were

detached by cell scraping as follows: after step 1, add 1 mL warm complete cell culture

medium, scrape with a 25 cm handle, 1.7 cm blade cell scraper (Sarstedt) and go to step 4.

3. The tube was centrifuged at 2,000 rpm for 3 minutes to pellet the cells. The supernatant was

removed and cells were re-suspended in 100 µL warm complete cell culture medium.

4. For propagation into a new T-25 flask, the flask was filled with 5 mL warm complete growth

medium and the cell suspension dilution factor was 1:10 (i.e. 10 µL per 100 µL suspension).

This allowed for approximately 3-4 days of culture before the next passage. For seeding into

glass bottom wells, the well was filled with 2 mL medium and the dilution factor was 1:15

for HEK293, 1:12 for COS7, CHO and HeLa, 1:10 for RAW264.7. For a wound assay,

HEK293 cells were seeded at 1:8 dilution. For cell stocking, one half of the suspension was

transferred into a 1.8 mL cryovial (Sarstedt) for the cell freezing procedure, described below.

32

For cell freezing, Styrofoam trays were used to slow the freezing process and prevent ice crystals

formation from damaging cell membranes.

1. Cell freezing medium was added drop-wise to the cell suspension by slowly dripping 0.5 mL

of medium around the edge of the cryovial. Once added, the mixture was gently pipetted up

and down to evenly distribute the cells.

2. After labeling, cryovials were placed in Styrofoam trays and put in a -20oC freezer overnight.

3. Cells in the tray were then transferred to a -80oC freezer overnight. The next day cells were

removed from the Styrofoam tray and put in cardboard freezer boxes for long term storage.

Transfections were performed using Lipofectamine 2000 according to the manufacturer’s

directions, which are summarized below.

1. One 1.5 mL tube was filled with 100 µL DMEM (without serum) and 1 µg plasmid. Another

1.5 mL tube was filled with 100 µL DMEM and 5 µL Lipofectamine 2000.

2. After 5 minute incubation, the contents were mixed together, and incubated for 25 minutes.

3. After the second incubation, the medium was removed from the cells in the glass-bottom dish

that are the target of the transfection procedure and replaced with 0.8 mL warm DMEM.

4. The liposome-DNA complex mixture was added to the cell dish and incubated at 37oC for 4

hours. Then, the medium was removed, 2 mL warm complete growth medium was added

and the cells were tested for transgene expression after 18-24 hours.

3.3.3 Fluorescence Microscope and Accessories

Imaging was performed using an inverted IX-81 microscope (Olympus, Markham ON) with a

Lambda DG4 xenon lamp and QuantEM 512SC CCD camera. There were 10x (air) and 20x,

40x, 60x and 100x (oil immersion) objectives. Bandpass filters (Semrock, Rochester, NY) for

CFP, YFP and mRFP excitation were 438/24 nm, 500/24 nm and 580/20 nm, respectively, and

for emission were 482/32 nm, 542/27 nm and 630/60 nm, respectively. For FRET imaging, a

dual bandpass filter was used (480/30 nm for CFP and 535/30 nm for YFP) with a Photometrics

DV2 505 nm beamsplitter such that CFP and YFP emission channels were monitored

simultaneously. The light intensity at the microscope stage, for CFP excitation light, was 25

mW/cm2. Overnight illumination for some migration assays was provided by an iPod 3

33

programmed to flash blue light (1s per 15s) with a blue light intensity of approximately 1

mW/cm2. Experiments were performed in a desktop incubation chamber set at 37oC (Solent

Scientific, Segensworth, UK). Image acquisition was done using MetaMorph Advanced

(Olympus) and image analysis was performed using MetaMorph and NIH ImageJ.

3.3.4 Time-lapse Imaging Experiments

Most imaging experiments in this work were 30 minutes in duration: a 5 minute control period

preceded stimulant addition and a 25 minute observation window.

1. When inhibitors (calmidazolium, Y-27632 or blebbistatin) were used, they were added to the

cells in the appropriate imaging medium 1 hour before the experiment.

2. For light-activation experiments cells were imaged in complete growth medium. For

experiments where stimulatory chemicals were used, cells were washed with 1 mL PBS to

remove serum and then 1.8 mL PBS or PBS with CaCl2/MgCl2 was added.

3. For FRET biosensor experiments, 1 frame per second was captured. For morphology

experiments, 1 frame per 5 seconds was captured. For illumination of LOVS1K or ChR2,

CFP excitation light was flashed for 300 ms every 10 s.

4. Stimulatory chemicals were added after a 5 minute control period as 200 µL of 10x

concentration to improve even diffusion throughout the dish.

3.4 Migration Assays

Two migration assays were applied to accomplish the third research objective, to determine

whether the engineered protein networks developed here could regulate cell migration. The

procedures used to conduct and analyze scratch wound closure assays (for brevity, wound assay)

and transwell migration assays are provided in this section.

3.4.1 Scratch Wound Closure Assays

Wound assays were performed as described below, based on similar assays reported in the

literature (90-92). In some studies a chemical was added to the medium to inhibit cell division,

such as mitomycin C (91) or TGF-β, to prevent cell growth from contributing to re-populating

the wound, however addition of extraneous chemicals is not desirable for reproducibility and

may introduce unexpected effects; therefore no growth inhibitors were used (90, 92).

34

1. Cells were passaged into glass-bottom wells and transfected as described above (section

3.3.2), except 2-3 µg of plasmid was used to achieve higher transfection efficiency.

2. The morning after transfection, a micropipette tip (0.5-10 µL) was used to make three

parallel scrapes across the confluent monolayer in the glass-bottom microwell portion of the

dish. The dish was then rinsed twice with PBS to remove debris created from the scraping. 2

mL complete growth medium, with any inhibitors, was then added.

3. Each of the three wounds was imaged using the 10x objective by photographing 5 random

locations along the wound. This sampled approximately one third of the wound.

4. The dish was transferred to the 37oC/5% CO2 incubator for 24 hours. Experiments requiring

periodic illumination were placed on the iPod. A thin, clear and colourless plastic insert was

placed between the dishes and the screen to minimize heat transfer from the iPod to the dish.

Experiments requiring darkness were loosely wrapped in aluminum foil.

5. The following morning, each of the wounds was imaged in the same manner as in step 3. The

assay was repeated twice more on two separate weeks for three independent experiments.

There is little consensus in analysis of wound assay images. Some studies count the number of

cells that have crossed the initial edge of the wound (91), however counting individual cells in a

10x image is difficult due to the low resolution. Calculating the wound area has also been done

by thresholding the image based on pixel intensity (90), however this method introduces user

bias when choosing a threshold. Another technique is to draw parallel lines across the two sides

of the wound edge and then calculate the distance between lines (92). This technique is also

wanting because wound edges are seldom parallel after 24 hours; determining where to draw a

straight line introduces considerable bias. Here, the third technique (based on parallel lines) was

modified to reduce bias due to an uneven wound edge. This method attempts to minimize user

bias because the locations of measurement are proscribed, several locations along an uneven

edge are sampled, and there is no need to make a thresholding judgment (Figure 3.4).

1. Five perpendicular lines were drawn at each quarter interval along the upper wound edge

until they contacted the lower wound edge.

2. The average length of these lines was taken as the wound width. The average wound width

for that condition was the average of 9 wounds (3 wounds x 3 independent experiments).

The average percent wound closure was calculated as shown.

35

Figure 3.4: Cartoon depiction of wound assays analysis.

For each image, x, five lines are drawn at quarter intervals perpendicular to the approximate wound edge. The average length of those lines (dxavg) is the wound width for that image (lines are only drawn for one image to simplify the figure). For each wound, 5 images are averaged.

3.4.2 Transwell Migration Assays

The protocol used here has been adapted from several studies that used transwell assays to

quantify Rho-dependent migration (71, 79, 89). Transwell inserts with polycarbonate filters, 8.0

µm pores and approximately 0.33 cm2 insert area (24-well format) were used (Corning, Lowell,

MA, USA). Wells were not coated with specific adhesive proteins, but were soaked in complete

growth medium before cell seeding to improve adhesion.

1. 24 hours before cell seeding, inserts were soaked in 100 µL complete growth medium. Cells

were passaged and transfected as described (section 3.3.2), with 2 µg DNA for transfection.

2. On the day of seeding, the incubating medium was removed from the inserts, cells were

detached from their 6-well culture dishes by trypsin, and seeded onto the inserts in 100 µL

DMEM + 1% FBS, with any inhibitors, in a 1:30 dilution (to match the reduction in area

from 6-well plate format to the insert membrane area) to the apical chamber. The same

volume of cell suspension was seeded in a well without insert as the positive control.

3. The basal chamber was then loaded with 650 µL DMEM + 1% FBS, with any inhibitors or

chemoattractants. For illumination, 24-well plates were placed on the iPod as for the wound

assays. Similarly, conditions requiring darkness were loosely wrapped in aluminum foil.

4. After 24 hours the transwell insert was removed from the basal chamber (the “Receiver

well”). Excess cells were removed from the apical layer by cotton swabs.

5. The insert was then soaked in the “PBS well” containing 500 µL PBS for 5 minutes to

remove remaining serum. The insert was then transferred to the “Trypsin well” containing

36

500 µL trypsin for 30 minutes. After 30 minutes, plates were tapped to detach cells, the

insert was removed and 500 µL complete growth medium was added to inhibit trypsin.

6. Images were taken of 5 random fields (10x objective) from the Receiver, PBS and Trypsin

wells of each insert after 1 hour incubation to allow for settling (Figure 3.5).

Quantifying the extent of migration is usually done by counting the number of cells, either with

or without a stain to aid visualization (71, 79, 89) or with a colourimetric assay, such as calcein

AM. The cell population here is non-homogenous; only the transfected cells are of interest and

all transfected cells are fluorescently labeled to aid in detection. For each condition, 5 random

images were taken of each of the three wells. The average number of fluorescent cells per image

was summed over the three wells to get the average number of migrated cells per image field for

that condition, Nx. Each condition was performed in triplicate. Total migration was summarized

as the migration index, Nx/Ncontrol, where Ncontrol is the average number of fluorescent cells per

image field in the control well. A migration index of 1 indicates complete migration of every

cell seeded onto the insert whereas 0 means no cells migrated through the filter.

Figure 3.5: Cartoon depiction of transwell migration analysis.

After 24 hours, the average number of cells per image field in the receiver (NXR), PBS (NXP) and trypsin (NXT) wells was calculated. The sum, NX, was used to compute the migration index, a ratio between NX and the average number of cells per image field in the control well, Ncontrol.

3.5 Data Analysis

This section briefly discusses several techniques that were used to analyze data in this work:

statistical comparisons were made throughout, fluorescence signal colocalization was used to

assess protein-protein association during initial characterization of RhoA and Rac1 chimeras, and

cell area calculations were made throughout the characterization of the Rac1 chimera. This

section contains a brief introduction; examples will be provided when the data is presented in

subsequent chapters,

37

3.5.1 Statistical Comparisons

The unpaired Student’s t-test was applied to determine if there was a significant difference in

some measured variable between two experiments. For this study α=0.05, so two-tailed P values

less than 0.05 were considered significant. Time-lapse imaging experiments, wound assays and

transwell migration were performed at least in triplicate and graphed data is presented as the

mean ± standard deviation, unless noted otherwise. For pre-stimulation experiments (e.g.

observing morphology of a cell without any stimulation), at least ten measurements were taken.

3.5.2 Fluorescence Signal Colocalization

Probes based on RBD and PBD were used, in combination with morphological changes, to judge

the activation of engineered Rho chimeras by Ca2+. Translocation of the cytoplasmic RBD and

PBD probes (mRFP channel) to the PM-bound Rho chimeras (YFP channel), were judged

visually and quantified using Pearson’s co-efficicent (PC). PC is used to measure the quality of

a linear fit, without reporting on the parameters of the fit itself: the co-efficient ranges from -1 to

1 where 1 indicates perfect colocalization, -1 indicates perfect exclusion and 0 indicates no

correlation (129). The “Pearson Co-efficient” plug-in in MetaMorph was used to calculate PC

for pairs of mRFP and YFP images before and after Ca2+ stimulation. Prior to PC calculation,

images were cropped to remove non-co-transfected cells and background subtracted by applying

a 32x32 pixel square median filter. PC was used to indicate whether colocalization increased or

decreased, without attempting to make a quantitative connection between PC and colocalization.

3.5.3 Area Calculations

Changes in cell area were used to assess whether Ca2+ stimulation resulted in lamellipodia

formation with RACer. Lamellipodia have an irregular shape, so quantifying their appearance is

difficult, unlike blebs which have distinctive shapes. Changes in cell area have been used as a

surrogate to report on effects of Rac1 in other reports of engineered Rac1 chimeras (5). Cell area

was calculated using the lowest intensity threshold that captured the whole cell area with ImageJ.

For cells too close to be delineated automatically, area was calculated by visual inspection. The

fold area change was normalized to the cell area at time zero (i.e. the first image in an

experiment always had fold area change = 1). A cell was considered to have an area increase if

the peak of the fold area change was at least twice the average noise of the graph after the

stimulus was added up to 20 minutes later.

38

4 CaM-RhoA Fusion Protein

The content of this chapter has been modified from a peer-reviewed journal article (130).

Reprinted from Cell Calcium, vol. 48, no. 4, Mills, E., Pham, E. and Truong, K., “Structure-

based design of a Ca2+ sensitive Rho protein that controls cell morphology,” pages 195-201,

© 2010, with permission from Elsevier.

4.1 Chapter Aims and Motivation

RhoA was chosen as the first target for engineered Ca2+/CaM-control because there were no

known interactions between CaM and RhoA at the time of the study, RhoA had not been

synthetically controlled by other groups in the past, as had been done for Rac1 and Cdc42 (5-6,

131-132), and RhoA-mediated blebbing is suspected in regulating the amoeboid-like cell

migration mode (72-73, 81). At the outset, the amino acid sequence of RhoA was analyzed

using a bioinformatics tool (114) designed to identify putative CaM binding peptide (CBP) sites

by searching for suitably spaced hydrophobic anchors and regions of high α–helical propensity,

among other factors. The discovery of a putative CaM binding site motivated the design of a

Ca2+/CaM-based switch of RhoA by a tandem fusion, without the need for extensive genetic

manipulations or structural analysis.

The four specific aims for the work in this chapter were drawn from the overall research aims:

1. Validate the CaM binding site in RhoA.

2. Design a fusion protein that could enable Ca2+/CaM control over RhoA.

3. Determine if the CaM-RhoA fusion could regulate cell morphology in response to

changes in intracellular Ca2+.

4. Subsequently, determine if Ca2+ signals mediated by exogenous Ca2+-mobilizing proteins,

delivered by transgenes, could also regulate cell morphology via the CaM-RhoA fusion

protein.

Several fusion proteins were created in this chapter during the design and characterization of the

CaM-RhoA fusion protein (Figure 4.1).

39

Figure 4.1: Overview of fusion proteins used in Chapter 4.

Schematic diagrams of fusion proteins (left), with the names that are used to discuss them in the text (right). Domains are shown in rounded rectangles and peptides are shown as black lines.

4.2 Results

The experimental results of this chapter are described below. The CaM binding site in RhoA

was verified by pull-down assays (section 4.2.1), this was used to design a tandem fusion where

CaM could bind RhoA (section 4.2.2), the CaM-RhoA fusion was shown to mediate bleb

retraction on Ca2+ stimulation (section 4.2.3) and light and ACh were used with ChR2 and

nAChR-α4, respectively, to also induce bleb retraction (section 4.2.4).

4.2.1 A CaM Binding Site in RhoA

A bioinformatic tool from the CaM target database identified residues 168-190 of RhoA (168-

REVFEMATRAALQARRGKSGC-190) as a putative CaM binding site. The algorithm attempts to

align key hydrophobic residues from the query sequences with one of the known CaM binding

motifs such as 1-10 or 1-14, while also considering α-helical propensity of residues in the query

and their Kyte-Doolittle values (114). The identified region of RhoA corresponds to the most

carboxy-terminal α-helix of RhoA and its carboxy-terminal unstructured tail, and includes part of

the CAAX prenylation motif (Figure 4.2A). This region of RhoA is surface exposed and so it

may indeed be accessible to CaM. The native α-helical structure of part of the putative binding

site also lends itself to the supposition that this region may be a CaM binding site. RhoA in

cassette format has a slightly different sequence than wild type RhoA towards the carboxy

terminus but the modified sequence of RhoA (RhoACBP) (REVFEMATRAALQARSPGASL) still

returned a putative CaM binding site by the search algorithm.

40

A RhoA-YFP fusion was prepared for a CaM pull-down assay to test whether RhoA could

associate with CaM-sepharose beads in a Ca2+-dependent manner. Since the putative binding

site is not near the Switch I/II regions, RhoA(DP)-YFP and RhoA(DN)-YFP were also tested.

Indeed, all three fusion proteins were pulled-down by CaM-sepharose beads in a 1 mM CaCl2

buffer and dissociated into solution when 2 mM EDTA was added (Figure 4.2B). YFP was used

as a negative control condition. To confirm that the identified sequence was sufficient for CaM

binding, RhoACBP was fused to YFP, YFP-RhoACBP, and this was tested for CaM binding in

the same manner; YFP-RhoACBP was also pulled-down by CaM-sepharose beads.

Figure 4.2: Identification of a CaM binding site in RhoA.

A. Ribbon structure of RhoA, PDB 1A2B (133), with RhoACBP in orange. B. SDS-PAGE of CaM-sepharose pull-down assay with RhoA-YFP (and its mutants) and YFP-RhoACBP.

Visual inspection of RhoACBP suggests that the most likely binding mode is 1-10, possibly 1-5-

10, with Val170, Leu179 and possibly Ala174 forming the key hydrophobic anchors. The 1-10

motif is based on the prototypical 1-10 CBP from Ca2+/CaM-dependent kinase II (134). If this

were the case, then CaM binding would either require the rearrangement of the three nearby β-

strands (residues 5-12 and 39-60) to accommodate the N- and C-terminal lobes of CaM forming

a hydrophobic binding pocket around the binding site, or the binding site itself may be “pulled”

away from its current location to enter CaM’s hydrophobic channel. If the CaM-RhoA

interaction were to occur as discussed, there would likely be a significant hydrophobic-driven

rearrangement of RhoA to account for the repositioning of the helix. This is probably an

energetically expensive interaction, which explains why it may occur rarely, if at all, in nature.

41

4.2.2 Design of a Ca2+-dependent RhoA-based Morphology Switch

Cell morphology, and plasma membrane (PM) blebbing in particular, can be used as an “output”

signal for a RhoA-based morphology switch. While RhoA has been implicated in regulating

several gene expression pathways (52-53), in this study the primary interest is in the relationship

between RhoA and morphology. Since cell morphology regulation is the goal of designing the

switch, it is desirable, and intuitive, to monitor cell morphology as the “output” of the switch.

RhoA natively switches between GTP- and GDP-loaded states to control cell morphology, so

overexpression of dominant positive RhoA, RhoA(DP)-YFP, and dominant negative RhoA,

RhoA(DN)-YFP, would likely inform the expected morphologies for a designed Ca2+-dependent

RhoA switch. When RhoA(DP)-YFP was overexpressed in a set of epithelial cell lines (CHO,

HEK293 and HeLa), dynamic, circular protrusions were present (Figure 4.3A-F), indicative of

literature reports of blebs (72-73, 81-82). Blebs were also present, although less frequently, for

wild type RhoA, but did not occur when YFP, CaM-YFP or RhoA(DN)-YFP were expressed.

The average size of the bleb relative to the cell body differed between cells: in HEK293 cells, the

blebs were much larger than in HeLa and CHO cells (Figure 4.3G). Rhodamine-phalloidin

staining of RhoA(DP)-YFP transfected HEK293 cells showed a circumferential ring of actin at

the edge of the bleb of varying intensity (Figure 4.3H), consistent with the observation that actin

is polymerized in aging blebs whereas only the monomer is present in new blebs (84).

Figure 4.3: RhoA-dependent morphologies.

A, C-F: Overexpression of RhoA(DP)-YFP (white box enlarged in B), RhoA-YFP, RhoA(DN)-YFP, YFP and CaM-YFP in CHO cells, respectively. G, H: RhoA(DP)-YFP in HEK293 cells stained with rhodamine-phalloidin (G is YFP channel, H is rhodamine). Scale bars are 25 µm.

42

Computational modeling and rational design were used to develop a Ca2+-dependent RhoA

switch, based on putative interaction between CaM and RhoACBP. Given that the interaction is

likely low affinity (due to the necessary spatial re-arrangements, and evidenced by the relatively

faint fluorescence in solution after CaM pull-down), a tandem fusion of CaM and RhoA would

increase the local concentrations of the two binding partners thereby making interaction more

favourable. The amino- and carboxy-termini of RhoA are relatively close in 3D space (about

16Å), so with a flexible linker, CaM may be able to access RhoACBP in a CaM-RhoA(DP)

fusion. A hypothetical structure of CaM bound to RhoACBP was made from the reported

structure of CaM bound to its target peptide from Ca2+/CaM-dependnet kinase II (134), which is

also a 1-5-10 motif peptide (PDB 1CDL (135), Figure 4.4A, B).

Using this model, a CaM-RhoA(DP)-YFP fusion was made using the fusion protein modeling

program FPMOD (136). In this larger model (Figure 4.4C), a four amino acid linker between

CaM and RhoA was required for FPMOD to find a conformational arrangement of the fusion

protein with CaM bound to RhoACBP (FPMOD disallows conformations where two atomic

coordinates are forced to overlap). Since the cassette cloning method introduces two amino

acids, and there are several amino acids at the amino terminus of RhoA before the first β-strand,

no additional linker was added to at the CaM-RhoA(DP) fusion site. The particular structure

presented in the figure is one of many possible conformations generated by FPMOD, and is not

necessarily the most likely to occur. The model is also limited because FPMOD considers all

structures as rigid, so FPMOD was instructed where to create a “hinge” at the amino-terminal

side of RhoACBP. In reality nature may choose a different way to re-arrange CaM-RhoA(DP);

the figure depicts one possible way for Ca2+ loading to affect the fusion protein.

A FRET-based probe, CFP-CaM-RhoA(DP)-YFP was created, and a repeatable, significant

change in the fluorescence spectrum was observed after Ca2+ loading (Figure 4.4D). The FRET-

based probe was used as a way to determine whether any Ca2+-dependent change occurred in the

overall conformation of CaM-RhoA(DP); the result does not report on any change in RhoA-

effector interactions, or the specific nature of the conformational change, simply that one

occurred. The change in the fluorescence spectrum was not due solely to EF-hand

rearrangement in CaM itself because CFP-CaM-YFP, did not exhibit any detectable changes in

spectrum before and after Ca2+ loading.

43

Figure 4.4: Structural modeling of the CaM-RhoA(DP) fusion protein.

A, B: The CaM-peptide interaction from 1CLD (with 1-5-10 hydrophobic anchors highlighted, and the re-drawing of 1CLD with the RhoACBP anchors. C: Structural models of CaM-RhoA(DP)-YFP generated using FPMOD: CaM (blue), RhoA (red), RhoACBP (orange), YFP (green), amino-terminal linker (black). D: Fluorescence spectra for apo (grey line) and Ca2+-loaded (black) CFP-CaM-Rhoa(DP)-YFP. The error bars show standard deviation of triplicate measurements. The difference was significant (P=0.002).

4.2.3 Ca2+-dependent Bleb Retraction by CaM-RhoA(DP)-YFP

When CHO, HeLa and HEK293 cells overexpressed CaM-RhoA(DP)-YFP, cells had dynamic,

circular blebs, similar to overexpression of RhoA(DP)-YFP. The fraction of transfected cells

with blebs (about 50%), the size of the blebs and their rate of expansion and retraction were all

qualitatively similar on visual inspection between RhoA(DP)-YFP and CaM-RhoA(DP)-YFP,

suggesting that at basal Ca2+ concentrations the fusion with CaM did not interfere with

RhoA(DP)’s role in establishing the blebbing morphology.

Stimulation by the purines ATP and UTP resulted in Ca2+ transients in CHO, HeLa and HEK293

cells (Figure 4.5). Before testing the effect of Ca2+ on CaM-RhoA(DP)-YFP, the FRET-based

TN-XL sensor was used to report on typical Ca2+ transients in these cell lines in response to ATP

and UTP. In CHO cells, ATP treatment led to a Ca2+ transient that oscillated around a plateau

44

above basal level for 8-10 minutes, while in HEK293 cells ATP stimulation resulted in a sharp

Ca2+ spike that lasted for 30-60 seconds. Both ATP and UTP successfully elicited responses in

HeLa cells of similar shape: a bimodal response with approximate duration of 3 minutes was

typical. The amplitude of the Ca2+ signals was determined by scaling each of the signals

between Rmax and Rmin. Rmax was established after the transient by treatment with 1 µM

ionomycin in Ca2+-PBS while Rmin was established by adding 2 mM EDTA after ionomycin.

Figure 4.5: Ca2+ transients elicited by ATP or UTP in CHO, HeLa and HEK293 cells.

The emission ratio (ratio of YFP/CFP emission intensities for CFP excitation) is reported between Rmax and Rmin. ATP or UTP was added at 0 minutes in these graphs; differing delays to Ca2+ transient onset are likely due to diffusion of the chemical in the imaging buffer.

For transfected cells overexpressing CaM-RhoA(DP)-YFP, ATP, UTP and ionomycin treatment

(for CHO, HeLa and HEK293, respectively) resulted in bleb retraction over 5-10 minutes (Figure

4.6 and Movies 4.1-4.3 and Appendices C and D). In CHO cells stimulated with ATP, dynamic

blebbing continued for approximately 10 minutes before the blebs appeared to become less

dynamic and eventually about 70-80% of the blebs retracted (Figure 4.6A, D-G). When CHO

cells were stimulated with ionomycin, the bleb retraction occurred much more quickly (1-2

minutes after stimulation) and was more complete (all blebs typically retracted). There was no

discernable pattern or change in the number of blebs on unstimulated CHO cells throughout the

20 minute experiments. The number of blebs on a cell throughout representative 20 minute

experiments is shown over time (Figure 4.6A-C). The number of blebs per cell was reported,

rather than the number of cells with bleb retraction, to distinguish between cells where a few,

many or all of the individual blebs were retracted. For these graphs the number of blebs per cell

was normalized to range between 0 and the maximum number of blebs per cell over the 20

minute window.

HeLa cells overexpressing CaM-RhoA(DP)-YFP stimulated with UTP showed partial bleb

retraction (Figure 4.6B, H-K), initially similarly to ATP stimulation in CHO cells, but then blebs

45

began to reform and by the end of the 20 minute window, the number of blebs had increased 20-

30% from its trough after stimulation. The re-appearance of blebs after experiencing a minimum

number of blebs was common in HeLa cells and may be due to several factors. The shorter Ca2+

transient compared to the ATP-induced transient in CHO cells may lead to a smaller magnitude

of RhoA inhibition, or the CaM-RhoACBP interaction itself may be reversible which enabled

bleb re-formation after the expiration of the Ca2+ transient. Similarly to CHO cells, ionomycin

stimulation resulted in rapid, nearly complete bleb retraction, and in the absence of stimulation

there was little change in the number of blebs.

For HEK293 cells ATP stimulation had no apparent effect on the large, dynamic blebs

characteristic of these cells, however ionomycin stimulation, as with CHO and HeLa cells,

resulted in near complete bleb retraction (Figure 4.6C, L-O). The inability of ATP stimulation to

induce bleb retraction in HEK293 cells may have been due to the short duration of the Ca2+

transient in these cells.

Blebbing and bleb retraction in stimulated cells expressing CaM-RhoA(DP)-YFP was dependent

on Ca2+/CaM and RhoA (Figure 4.7 summarizes results for CHO cells and Appendix D for HeLa

and HEK293 cells). Introducing the T19N mutation to create CaM-RhoA(DN)-YFP resulted in

no cells blebbing at basal Ca2+, and as expected, Ca2+ stimulation had no effect on these cells.

As more instructive controls, cells overexpressing RhoA(DP)-YFP were stimulated without

significant effect on the number of blebs per cells, suggesting that the fusion with CaM was

necessary to mediate the bleb retraction. For CaM-RhoA(DP)-YFP cells, incubation with the

CaM inhibitor CDZ, which occupies CaM’s hydrophobic binding pocket (137), prevented bleb

retraction. Additionally, the MLCKp CaM-binding peptide was introduced in the CaM-RhoA

fusion as a peptide inhibitor for the CaM-RhoACBP interaction. The CaM-MLCKp-RhoA(DP)-

YFP construct caused blebbing at basal Ca2+ but stimulation had no significant effect on the

number of blebs per cell. Finally, CaM-RhoA(DP)-YFP overexpressing cells were incubated

with Y-27632, a ROCK inhibitor. Under this condition, few cells (approximately 5% of cells)

still blebbed at basal Ca2+, but there was no significant bleb retraction on stimulation in those

cells. Interpreting this result is confounded by the fact that ROCK is known to both cause (72,

74, 82) and retract (73) blebs. It is possible that blebbing in this population of cells was due to

some other phenomenon, such as apoptosis (138), or that the RhoA-ROCK activity in those cells

was so high as to not be inhibited at the concentration of Y-27632 used here.

46

Figure 4.6: Ca2+-induced bleb retraction in CHO, HeLa and HEK293 cells.

A-C: The number of blebs is shown as a function of time for CHO, HeLa and HEK293 cells, respectively, transfected with CaM-RhoA(DP)-YFP after no stimulation (open squares), ATP stimulation (UTP for HeLa) (grey squares) or ionomycin stimulation (black squares). While these data are not continuous, a line has been drawn between data points to help the reader follow the trend. D-G: Time course of bleb retraction in CHO cells showing images immediately before stimulation, at the start of the morphology change, when the morphology change was almost complete, and then 20 minutes after stimulation, respectively. The white arrow indicates the area enlarged in the inset. H-K: A similar time course of images for HeLa cells, and L-O: HEK293 cells. Scale bars are 30 µm (D-O) and 6 µm in the insets (D-K).

47

Figure 4.7: Summary of bleb retraction in CHO cells.

The number of blebs was counted before (black bars) and 20 minutes after stimulation (grey bars); the percentage change was calculated (white bars). All data are the mean of at least 10 cells over at least 3 independent experiments and the error bars are the standard deviation. * denotes P < 0.001 by pair t-test and ** denotes P < 0.001 by unpaired t-test. Data is mean ± standard deviation, n=3 independent experiments.

4.2.4 Light- and ACh-dependent Bleb Retraction

Bleb retraction in response to light and ACh was realized in CHO cells overexpressing CaM-

RhoA(DP) and an appropriate Ca2+-mobilizing protein module (Figure 4.8). One of the primary

motivations for re-programming cells using Ca2+ signaling and Ca2+-responsive proteins was to

leverage the inherent modularity of Ca2+ and the diverse set of Ca2+-mobilizing protein modules

that exist. Light is a convenient laboratory signal to deliver and control: when illuminated by

blue light, ChR2 increases its permeability for Ca2+ and may be a suitable Ca2+-mobilizing

protein module. ACh is an exogenous chemical for HEK293 cells: when nAChR-α4 is bound by

ACh, its permeability for Ca2+ also increases.

Plasmids coding for ChR2 and nAChR-α4, both tagged with YFP were co-transfected with a

mRFP variant of CaM-RhoA(DP)-YFP, CaM-RhoA(DP)-mRFP, which was verified to function

similarly to CaM-RhoA(DP)-YFP prior to experimentation. Successful co-transfection was

verified by labeling of cells with both fluorescent proteins. For ChR2 co-transfections, cells

48

were illuminated with CFP excitation light flickering (a band pass filter centred on 438 nm) and

bleb retraction was observed gradually over approximately 20 minutes. When illumination was

performed solely with mRFP excitation light (a band pass filter centred on 580 nm) no bleb

retraction was observed. Similarly, when cells expressing only CaM-RhoA(DP)-mRFP were

illuminated with flickering CFP excitation light blebs continued to form dynamically. For

nAChR-α4 co-transfected cells, 1 mM ACh application resulted in bleb retraction that began 5-

10 minutes after stimulation. For cells expressing only CaM-RhoA(DP)-mRFP stimulated with

ACh there was no apparent effect on blebbing behaviour.

Figure 4.8: Light- and ACh-mediate bleb retraction.

A: Schematic diagram of Ca2+-based bleb retraction: the Ca2+ dependent process can be made sensitive to a wide range of stimuli due to the prevalence of Ca2+ signaling in nature. Here, we have demonstrated this with blue light (via ChR2) and ACh (via nAChR), but many other stimuli may be candidates. B-E: CHO cells co-transfected with ChR2 plasmid and CaM-RhoA(DP)-mRFP demonstrate bleb retraction after stimulated with blue light (0, 5, 10 and 20 minutes after irradiation onset, respectively). F-I: CHO cells co-transfected with nAChR plasmid and CaM-RhoA(DP)-mRFP demonstrate bleb retraction after stimulation with ACh (0, 5, 10 and 20 minutes after addition of ACh). Scale bars in B-I are 30 µm and 6 µm in the insets.

49

4.3 Discussion

A putative 1-10 motif CaM binding site in the carboxy-terminal region of RhoA was identified

using an algorithm that considers the arrangement and chemical characteristics of amino acids in

a query sequence. While the target region is surface exposed, significant rearrangements of

RhoA would be required to facilitate CaM binding, and so the physiological significance of the

interaction, if any, is an open question. The affinity between CaM and RhoACBP was strong

enough to pull-down YFP in detectable quantities. Applying the CaM binding motif search

algorithm to Rac1 revealed no potential binding sites; for Cdc42 a site was found (amino acids

147-167), but the location is part of a β-sheet and is unlikely to support CaM binding. Mutations

to the carboxy-terminal regions of Rac1 and Cdc42 may support CaM binding, and this may

provide a method for engineering Ca2+ control over these proteins in the future (see Chapter 6).

ATP and UTP were used to induce bleb retraction in CHO and HeLa cells, respectively. ATP

was unable to elicit a similar response in HEK293 cells, although bleb retraction in those cells

was achieved with ionomycin stimulation. The short duration of the ATP-induced Ca2+ transient

was likely why bleb retraction was not observed. This suggests that RhoA-ROCK activity must

be silenced for a certain duration of time for there to be an effect on blebbing. It was also

observed that HeLa cells partially resumed blebbing after UTP treatment, approximately 3-5

minutes after the blebbing morphology was minimized. UTP-induced Ca2+ transients were

shorter than ionomycin signals, or ATP-transients in CHO cells. Taken together, these two

observations suggest a relationship between the duration of the Ca2+ transient and the duration of

morphology changes: shorter Ca2+ signals have little effect on morphology while longer Ca2+

signals disrupt blebbing for longer periods. Resumption of blebbing after the expiration of the

Ca2+ transient suggests that CaM interaction with RhoA in the fusion is reversible.

Several cellular mechanisms are responsible for RhoA inhibition leading to bleb retraction. At

the global cell level, since RhoA-ROCK phosphorylates its targets, ROCK inactivation would

not have immediate effect on its targets until they are dephosphorylated. Phosphatases, such as

myosin light chain phosphatase, are then able to dephosphorylate RhoA-ROCK’s targets, namely

myosin regulatory light chain (139). This leads to a reduction in cytoskeletal tension and

prevents new bleb formation (74-76). In order for a bleb to retract, a successive recruitment of

cytoskeletal proteins to the bleb was shown in an elegant study (84). Ezrin is recruited to the PM

50

by ankyrin B, a protein constitutively present in blebs; ezrin recruits actin monomers. The

mechanism of actin nucleation was unclear in the study because neither Arp2/3 or mDia were

shown to be significant factors in bleb retraction. Subsequent to actin recruitment, bundling

proteins, such as α-actinin and coronin, are recruited, followed by myosin adaptors like

tropomysoin, and finally myosin II. Myosin II-actin cross-linking powers the contraction, which

is ironically also the source of tension that generated the bleb. Whether the recruitment process

is constitutive, or is initiated by a signaling event during the growth of the bleb, is unknown (84).

The experiments with light/ChR2 and ACh/nAChR-α4 showed that, in principle, it is possible to

activate engineered Ca2+-responsive proteins using exogenous stimuli, provided that suitable

Ca2+-mobilizing modules are present to interpret those stimuli: simple regulation networks based

on Ca2+ signals were created. Other Ca2+-mobilizing modules may be used for increased

signaling versatility such as VEGFR2 (10) or synthetic modules such as LOVS1K (12). In terms

of controlling cell migration, the CaM-RhoA fusion protein developed here may be used to

facilitate a “stop here” signal, to use the language of cell clearance researchers (140). For

example, blebbing, randomly migrating re-programmed cells, upon sensing an extracellular

stimulus that generates a Ca2+ transient, would stop blebbing and remain at a target site to

perform some function. However, prolonged RhoA-ROCK activity may have undesirable

effects on the cell, and from an investigative standpoint it is more desirable to activate a

phenomenon than de-activate it, so as to clearly establish cause-and-effect relationships.

4.4 Chapter Summary and Conclusion

In this chapter, CaM and RhoA were combined in a fusion protein to regulate bleb retraction in

response to various Ca2+ signals.

1. A CaM binding site in RhoA, RhoACBP, was identified with a bioinformatic algorithm

and was confirmed by CaM-sepharose pull-down assays and SDS-PAGE.

2. The CaM-RhoA(DP)-YFP fusion protein was designed using FPMOD, a structural

modeling program.

3. Bleb retraction was observed in response to ATP, UTP and ionomycin, chemicals that

elevated intracellular Ca2+, in cells overexpressing CaM-RhoA(DP)-YFP. The bleb

retraction process was dependent on Ca2+, CaM and RhoA/ROCK.

51

4. Light and ACh were used, in combination with ChR2 and nAChR-α4, respectively, to

cause bleb retraction in cells co-transfected with CaM-RhoA(DP)-mRFP, demonstrating

that Ca2+ signals mediated by exogenous Ca2+-mobilizing proteins could regulate

engineered Ca2+-responsive proteins.

In relation to the research objectives outlined in Chapter 1, the work in this chapter addresses the

first and second objectives. 1. A chimeric protein of CaM and RhoA was created using

computational modeling and rational design that controlled cell morphology, bleb retraction in

particular, in response to changes in intracellular Ca2+. 2. Control over bleb retraction was

achieved using two exogenous input signals, light and ACh, by introducing suitable Ca2+-

mobilizing proteins to create a synthetic two-node protein network.

52

5 CaRQ: A New Ca2+-Sensitive RhoA Chimera


Reprinted from Chemistry & Biology, vol. 18, no. 12, Mills, E. and Truong, K., “Ca2+-mediated

synthetic biosystems offer protein design versatility, signal specificity and pathway rewiring,”

pages 1611-1619, © 2011, with permission from Elsevier.


The work of the previous chapter with the CaM-RhoA fusion protein demonstrated that Rho

proteins could be engineered such that Ca2+ signals were able to regulate cell morphology and

that exogenous signals, in concert with Ca2+-mobilizing protein modules, were also able to

regulate cell morphology. However, as a tool to for re-programming cell migration, the CaM-

RhoA fusion is limited in two key ways: first, Ca2+ only switched RhoA activity from “on” to

“off”, whereas an “off” to “on” switch would be more useful for regulating RhoA, and, second,

because the interaction relied on a naturally occurring amino acid sequence, the ability to tune

the interaction or apply it to other GTPases is limited. Both of these limitations could be

overcome by introducing a CaM binding peptide as part of a RhoA/CBP chimera. Introducing a

CBP as a chimera with RhoA would allow the interaction between CaM and RhoA to be tuned

by using CBP’s of different affinity and binding modes for Ca2+-CaM. This would allow a Ca2+-

based switch to operate from “off” to “on” or “on” to “off”, and based on the structural similarity

between them, may allow the same strategy to be applied to regulate other Rho proteins.

The five specific aims for the work in this chapter were drawn from the overall research aims:

1. Determine a suitable design for the RhoA/CBP chimera.

2. Characterize the RhoA/CBP chimera in terms of its overall Ca2+ affinity, response to

different Ca2+ transient-inducing chemicals, and characteristic morphology.

3. Investigate how Ca2+-mobilizing modules with distinct spatial properties to their Ca2+

signals can be used to regulate morphology with the RhoA/CBP chimera.

4. Similarly, determine the effect of the temporal properties of Ca2+ signals on the

RhoA/CBP chimera-induced morphology changes.

53

5. Determine whether activation of the RhoA/CBP chimera affects cell migration, and

characterize the effect if there is one.


RhoA/CBP chimera (Figure 5.1).

Figure 5.1: Overview of main fusion proteins created in Chapter 5.

Schematic layout of the fusion proteins are shown on the left and their names as used in the text are given on the right. All of the RhoA carboxy-portion fragments in the figure have the Q63L mutation. Dominant negative mutants were made bearing the T19N mutation in the RhoA amino-portion fragment, but these have been omitted from the figure for simplicity.

5.2 Results

The experimental results of this chapter are described here. The rational design of the

RhoA/CBP chimera is described (section 5.2.1) and the chimera, CaRQ, was characterized by

cellular pull-down and stimulation by various Ca2+-elevating chemicals (section 5.2.2). CaRQ

was sensitive to Ca2+ signals with different spatial patterns, and variants of CaRQ with different

subcellular localizations had distinct responses to Ca2+ signals (section 5.2.3); similarly, there

was a reproducible relationship between the duration of the input Ca2+ signal and the resultant

duration of the cellular morphology change (section 5.2.4). Prolonged activation of CaRQ,

either by light flickering or chemical concentration gradient presentation, increased cell

migration in wound and transwell assays, respectively (section 5.2.5).

5.2.1 Rational Design of CaRQ, a Ca2+-Sensitive RhoA Chimera

A RhoA/CBP chimera was designed such that an IQ-motif peptide (IQp) was embedded between

residues 49 and 50 of RhoA (Figure 5.2). Embedding a CBP into RhoA has two main strengths

in terms of designing a switch. First, the large library of CBP’s means that the CaM-binding

characteristics of the chimera can be tuned for kinetics, binding mode and Ca2+-dependence,

54

assuming linearity of CBP function once the fusion is constructed. Second, the chimera design

will likely be applicable to other Rho proteins, owing to the structural similarity within the Rho

family. To retain RhoA’s native structure and function in the absence of Ca2+-CaM interaction, a

CBP should be added to RhoA either at the amino or carboxy termini, or within one of the

surface exposed loops so as to not disrupt secondary structure elements. Throughout Chapters 5

and 6, RhoA is RhoA(Q63L) so that RhoA is not subject to normal GTP/GDP cycling.

There are 11 surface exposed loops in RhoA, and three were selected for construction (Figure

5.2A). The embedded peptide used to construct the chimeras was an IQ motif peptide (IQp)

from myosin V (see Appendices A and B for sequences). Since IQp binds apoCaM, this would

allow CaM to bind RhoA/IQp at basal Ca2+ concentrations. This would presumably disrupt

RhoA’s ability to bind effector proteins and establish the “off” state for the morphology switch.

Three sites were initially chosen for chimera construction: RhoA/IQp-1, -2 and -3 corresponded

to IQp insertions between amino acids 26/27, 49/50 and 71/72, respectively. The insertions at

amino acids 26/27 and 71/72 correspond to the Switch I and II regions of RhoA, respectively and

were chosen because inserting peptides here would surely disrupt effector binding when CaM

was bound to IQp. The insertion site at amino acids 49/50 was chosen because it is sandwiched

between regions of well-defined secondary structure and it is near the amino terminus in 3D

space, to facilitate access to IQp from a possibly fused CaM. YFP was added to each chimera as

a carboxy-terminal fusion for visualization. When these chimeras were constructed, only

RhoA/IQp-2 was well expressed in COS7 and HEK293 cells, and this one was selected for

further study. The others showed punctuate fluorescence distributions in localizations that

resembled the ER or Golgi apparatus, which suggested the chimeras were not properly folded.

An amino-terminal fusion of CaM-MLCKp was added to RhoA/IQp-2 to facilitate Ca2+-induced

switching (Figure 5.2B, C). Most IQ-motif peptides bind CaM independently of CaM’s Ca2+-

loading state, so to encourage CaM’s dissociation from IQp on Ca2+-loading, a higher affinity

binding partner was presented in the form of MLCKp. IQp has low micromolar affinity for CaM

(119), while MLCKp has low nanomolar affinity for Ca2+-CaM (117). CaM itself was also

included in the chimera to increase its local concentration for the two CBP’s, and possibly also to

limit the chimera from acting as a sink for native CaM in the cell. All together, the intended

functioning of the chimera is that CaM binds IQp at basal Ca2+ disrupting RhoA-effector

interaction, and upon Ca2+-loading, Ca2+-CaM will dissociate from IQp in favour of the nearby

55

MLCKp, releasing RhoA to interact with its effectors (Figure 5.2B). For brevity the chimera

will subsequently be referred to as CaRQ (Ca2+-responsive RhoA with embedded IQp).

A ribbon structure of CaRQ in the absence and presence of Ca2+ was created using MODELLER

(142) to visualize possible conformations of CaRQ (Figure 5.2C). Structures of RhoA (PDB:

1A2B (133)), CaM bound to MLCKp (PDB: 2BBM (143)) and CaM bound to IQp (PDB: 2IX7

(144)) were used to generate the model, and unbound CBP’s were assumed to be random loops.

MODELLER uses known structures as a basis for computing the atomic spatial arrangements of

homologous proteins by minimizing the difference between the model and a library of

probability distributions for Cα-Cα distances, main-chain N-O distances and main-chain and

side-chain dihedral angles (145). The model suggested that CaM switching between the two

CBP’s is possible without violating spatial restraints however it did not provide insight into

whether CaM switching will impact RhoA-effector binding.

Figure 5.2: Modeling of CaRQ.

A. Ribbon diagram of RhoA highlighting the surface-exposed loops (red) and those that were used for initial construction (blue, numbered according to discussion in text). B. Cartoon illustrating the intended function of CaRQ. C. Ribbon diagram of the CaRQ model built with MODELLER in the absence (left) and presence (right) of Ca2+. The colour coding is equivalent to the legend in B. The YFP tag has been omitted for simplicity.

56

5.2.2 Characterization of CaRQ

Ionomycin stimulation and Ca2+ influx in cells co-expressing CaRQ and a fluorescent RBD

probe increased colocalization between CaRQ and the probe (Figure 5.3 and Appendix E). Ca2+-

dependent activation of CaRQ could not be measured strictly in vitro; when CaRQ was cultured

in E. Coli and mechanically extracted, no association was observed between RBD-mRFP-GST-

labelled glutathione-agarose beads and CaRQ under conditions of high or low Ca2+, probably due

to improper folding of CaRQ in the E. Coli host. Instead, RBD-mRFP-GST and a membrane

tagged version of CaRQ, pLyn-CaRQ, were co-expressed in COS7 cells. This way pairs of YFP-

and mRFP-channel images could be compared by calculating their PC as a way of detecting

changes in fluorescent protein colocalization. COS7 cells were used because RhoA(Q63L)-

dependent blebbing has not been observed in these cells, and so there was little chance of

morphology changes confounding changes in colocalization. Still, CFP was also co-expressed in

these cells as a spectrally distinct volume indicator (146-147). PC correlation between RBD and

CaRQ increased by 26% after stimulation with ionomycin, while the correlation change was less

than 1% for the dominant negative (T19N) CaRQ (Figure 5.3). Correlation between CaRQ and

the volume indicator decreased after ionomycin stimulation, due to a less even distribution of

CaRQ; the increased YFP-mRFP correlation was not a volumetric effect.

Figure 5.3: Colocalization analysis of CaRQ and RBD.

A. Cells co-expressing the RBD probe (mRFP), CaRQ (YFP) and CFP, and B cells co-expressing the RBD probe, CaRQ(T19N) (YFP) and CFP. 0 and 10 minutes refer to immediately before and 10 minutes after addition of ionomycin. PC values for the indicated image pairs are given on the right of the composite images, and the percentage changes over 10 minutes are calculated at the bottom. Scale bars are 30 µm. Images were not falsely coloured to preserve the apparent channel intensities.

57

CaRQ caused Ca2+-induced morphology changes consistent with switchable RhoA activity in

several mammalian cell lines including HEK293, HeLa and CHO cells (Figure 5.4 and Appendix

F). HEK293 cells were chosen for the majority of the characterization because the large blebs in

these cells allowed for easier counting with less chance of error. Cells expressing CaRQ were no

more likely to bleb than YFP-expressing control cells prior to stimulation, and much less likely

to bleb than cells expressing RhoA(Q63L)-YFP, consistent with the design of CaRQ to suppress

RhoA activity at basal Ca2+ (Figure 5.4A). Stimulation of CaRQ-expressing cells with

ionomycin resulted in the appearance of dynamic, circular blebs in approximately 50% of cells;

this phenomenon did not occur in cells expressing YFP alone. The blebbing morphology was

usually maintained for 2-3 minutes before subsiding and the cell morphology returning to normal

(Figure 5.4B-D, Movies 5.1-5.3). While blebs only developed in approximately 50% of

stimulated, transfected cells, RhoA(Q63L) overexpression appeared to cause blebbing in also

about 50% of transfected cells, suggesting that the efficiency of CaRQ-induced blebbing was

similar to the efficiency of RhoA(Q63L)-induced blebbing. Blebbing may not be observed in all

cells due to differences in transgene expression level. A second switch, CaRM, was created by

changing the CBP embedded within RhoA, and is discussed further later in this section.

Ionomycin-induced blebbing did not occur in several control cases that were tested (Figure

5.4F). Cells expressing YFP, CaM-YFP or the dominant negative RhoA mutant CaRQ(T19N)

did not develop blebbing in response to ionomycin. Similarly, there were no morphology

changes in cells that had been pre-incubated with the ROCK inhibitor Y-27632 or the CaM

inhibitor CDZ. The logical effect of the CaM inhibitor in this case is not straightforward because

in CaRQ CaM binds a CBP in both high and low Ca2+ conformations. CDZ is known to

preferentially bind Ca2+-CaM (148), which would explain why the switch from inactivated RhoA

to activated RhoA was prevented. CaRQ was also characterized in HeLa and CHO cells

(Appendix F).

A pair of mutations (E91H, N92H) has been reported to reduce the tendency of Rac1(Q61L) to

act as a sink for cellular GAP’s (5). The analogous mutations were made to CaRQ (E93H,

N94H), however they did not appear to have any affect on the morphology of cells prior to

stimulation or after stimulation with ionomycin. The caging mechanism reported for PARac is

substantially different from the CaM/CBP approach taken here, which may explain why these

mutations had little effect on CaRQ.

58

Figure 5.4: Stimulation of CaRQ in HEK293 cells.

A. Percent of cells expressing the indicated constructs prior to stimulation. Here, n=10 experiments, at least 100 cells per condition. B. Time course showing blebbing morphology versus time for representative cells transfected with CaR-Q (dark line) and CaR-M (light line) after ionomycin stimulus. C-E. HEK293 cells transfected with YFP (C), CaR-Q (D) or CaR-M (E) were stimulated with ionomycin at 5 minutes, with white arrows highlighting dynamic blebs. Scale bars in C-E are 30 µm. F. The percent of cells blebbing after ionomycin stimulation for the indicated conditions. Here, n=3 experiments and at least 10 cells per condition.

59

Replacing the IQp peptide embedded in CaRQ with the MLCKp peptide, or CaRM, resulted in a

switch that appeared to have the opposite Ca2+-induced behaviour than CaRQ (Figure 5.4B, E).

One of the proposed strengths of the embedded peptide approach to engineering control over

RhoA was that changing the peptide would have predictable effects on the behaviour of the

chimera. Replacing the IQp with MLCKp should prevent CaM binding to RhoA at basal Ca2+.

If the amino-terminally fused MLCKp was removed, then CaM should bind the embedded

MLCKp on Ca2+ loading and disrupt RhoA activity, thus switching RhoA from “on” to “off”.

Consistent with this, a similar fraction of cells overexpressing CaRM blebbed before stimulation

as compared to RhoA(Q63L). When CaRM cells were stimulated with ionomycin, blebbing

slowed and blebs retracted in most cells. There may have been some contribution to this effect

from CaM binding to endogenous RhoACBP. However, the nanomolar affinity interaction

between Ca2+-CaM and MLCKp likely meant that RhoACBP interaction was minimal.

There was a dose-dependent relationship between extracellular Ca2+ and the percentage of cells

that developed the blebbing morphology (Figure 5.5). Membrane-tagged CaRQ was used here

because it had a higher maximal response to ionomycin stimulation than cytoplasmic CaRQ; this

allowed for a better fit since the data was spread over a wider range on the dependent axis. The

cell medium was buffered using a mixture of CaEGTA and K2EGTA (Biotium Inc., Hayward,

CA, USA) to maintain a reliable extracellular free Ca2+ concentration when the intended

concentration was below 100 µM; a range from 0.1 to 1000 µM was tested. The paired data was

fit to a model sigmoid expression where the maximal and minimal changes and EC50 value were

parameters. The fit was calculated by minimizing the sum of square differences between the

model and the data using MS Excel. The free concentration of Ca2+ resulting in 50% change

(EC50) was 10-4.56, or 27 µM. The quality of the fit was assessed by calculating the square of the

PC between the data and the model: r2=0.977, indicating a good quality fit. The EC50 value fits

within the expected potential tuning range (0.6 to 160 µM) for Ca2+/CaM-based sensors (149).

The blebbing morphology induced by ionomycin stimulation of CaRQ is not indicative of

apoptosis (Figure 5.6). The appearance of blebs in apoptotic cells is followed by caspase-3/7

activation after 1-2 hours in some cells (138). However, over two hours after CaRQ activation

there was no detectable cleavage of a caspase-3/7 biosensor, based on a reported design (150).

Staurosporine (STS) was used as a positive control. These results suggest that CaRQ-induced

blebbing is not indicative of apoptosis. ROCK activation has been reported in apoptotic

60

cardiomyocytes, however the activation mechanism is irreversible cleavage of the autoinhibitory

domain by caspase-3 (138) and not the typically reversible RhoA-binding autoinhibition.

Figure 5.5: Determination of the Ca2+ EC50 value for pLyn-CaRQ.

Data were fit to a standard sigmoid of the form indicated at the top by minimizing the sum of squares between the fitted curve (dashed line) and the data (filled squares). The error bars are the standard deviation and n=3 independent experiments for each condition. The quality of the fit (r2) is shown, as are the fitted coefficients.

Figure 5.6: A caspase-3/7 sensor uncleaved in stimulated CaRQ-expressing cells.

A. The sensor layout is depicted with the caspase-3/7 substrate in bold. B. Representative cells exposed to 1 µM STS (staurosporine) as a positive control (top) and cells co-expressing the caspase-3/7 sensor and pLyn-CaRQ stimulated with 1.5 µM ionomycin in Ca2+-PBS. The white arrow head shows nuclear translocation. Scale bar is 10 µM. STS control data is from X. Chen.

61

5.2.3 The Effect of Ca2+-mobilizing Modules and Spatial Localization on CaRQ

CaRQ was sensitive to local Ca2+ signals and different subcellular localizations of CaRQ had

specific effects on its activation (Figure 5.7-5.9). This observation came about by way of trying

to activate CaRQ-mediated morphology change using different Ca2+-mobilizing modules. In

particular when HEK293 cells co-expressing CaRQ and nAChR-α4 were stimulated with 1 mM

ACh, blebbing was not observed within a 30 minute imaging window (Figure 5.7A). Similarly,

when CaRQ-expressing cells were stimulated using endogenous signaling pathways with ATP,

there were no apparent morphology changes either. To rescue CaRQ activation by ACh/nAChR-

α4, CaRQ was fused to the PM using the constitutive membrane tag pLyn (Figure 5.7B-D,

Movies 5.4 and 5.5). Two complimentary ideas motivated this. First, the source of the

ACh/nAChR-α4 Ca2+ transient is Ca2+ influx from the extracellular space (9). This means that

the local Ca2+ concentration at the PM (source of Ca2+ entry) may be higher than the average

Ca2+ concentration in the cytoplasm (110) during the ACh-induced transient. Second, because

RhoA is membrane-localized natively (37-39, 69), restoring the membrane localization with

pLyn may increase CaRQ’s ability to interact with RhoA’s effectors. Both ideas are related in

that they increase the ability of CaRQ to translate Ca2+ into RhoA activity.

When HEK293 cells co-expressing pLyn-CaRQ and nAChR-α4 were stimulated with ACh, cells

began to bleb 2-3 minutes after the stimulus was added and the blebbing morphology routinely

lasted for 10-15 minutes before cell morphology returned to normal (Figure 5.7C, D). As with

the ionomycin stimulation experiments, pre-incubation with Y-27632 and CDZ prevented ACh-

induced blebbing. The dominant negative mutation, pLyn-CaRQ(T19N), also abrogated ACh-

induced blebbing. Overexpression of pLyn-CaRQ did not appear to increase the fraction of

transfected cells that were blebbing before stimulation; that is, the membrane localization did not

increase the likelihood of CaRQ producing a RhoA-related morphology at basal Ca2+.

pLyn-CaRQ was more effective at creating morphology changes than CaRQ for ATP-, ChR2-

and LOVS1K-induced Ca2+ signals (Figure 5.8). ATP, ChR2 and LOVS1K each represent an

endogenous, exogenous and engineered Ca2+-mobilizing module. For ChR2 and LOVS1K, the

source of Ca2+ entry is direct influx from the extracellular space (11, 29, 136), while for ATP

there is likely a contribution from both the ER and the extracelluar space (99, 101). For each of

these there was an increase in the fraction of blebbing cells observed after stimulus. For ATP,

62

the stimulus was bolus addition of 10 µM to the imaging well, and for ChR2 and LOVS1K

stimulation was provided by CFP excitation light 300ms every 10s. Stimulation of ChR2 was

more likely to induce blebbing with CaRQ than ATP and LOVS1K, however ChR2 was more

efficient with pLyn-CaRQ than CaRQ. For LOVS1K, the preferentially undocked “bright”

mutant (I539E, bLOVS1K) and preferentially docked “dark” mutant (C450M, dLOVS1K) were

also used as controls (5, 12, 26) to verify that the LOVS1K switching mechanism was

responsible for morphology changes.

Figure 5.7: Activation of CaRQ and pLyn-CaRQ by ACh/nAChR-α4.

A: HEK293 cells co-transfected with nAChR-α4 and CaRQ and stimulated with ACh at 5 minutes. B: Cartoon hypothesizing how locally high Ca2+ activates pLyn-CaRQ but not CaRQ: nAChR-α4 (black), CaRQ (blue and green). C: The percent of morphologically normal, pLyn-CaRQ and nAChR-α4 co-expressing HEK293 cells that developed a blebbing morphology after ACh stimulus, under the indicated transfection and inhibitor conditions (n=3 experiments each for at least 10 cells). D: HEK293 cells co-transfected with nAChR-α4 and pLyn-CaRQ and stimulated with ACh at 5 minutes. White arrow higlights region of bleb formation and indicates area enlarged in inset. Scale bars in A and D are 30 µm and 15 µm in insets.

63

Figure 5.8: Activation of pLyn-CaRQ and CaRQ by ATP and blue light.

HEK293 cells co-expressing no extra constructs (A), ChR2 (B) or LOVS1K/Orai1 (C) were stimulated by ATP (A) or flashing CFP excitation light (B and C). The percentage of blebbing cells in the 25 minute imaging window after the onset of stimulation is shown as the mean ± standard deviation from three independent experiments and at least 10 cells for each condition.

When localized to the ER, CaRQ was able to detect Ca2+ signals from an engineered SOCE

event and cause morphology changes (Figure 5.9). SOCE is a common mode of Ca2+ entry from

the extracellular space. LOVS1K is based on Stim1 and Orai1, proteins that are used by the cell

to induce SOCE, but in the case of LOVS1K, Ca2+ entry through Orai1 is decoupled from

intracellular signaling events. To trigger SOCE on demand, the Ca2+-ATPase inhibitor

thapsigargin (Tg) was used to prevent re-uptake of Ca2+ into the ER lumen. The cell then

initiates Ca2+ influx from the extracellular space, for example, via Stim1 (an ER-bound Ca2+

sensor) and Orai1 (a PM-bound Ca2+ channel) (35, 151). During SOCE Stim1 and Orai1

oligomerize into discrete puncta where the ER is pulled very close to the PM (152). If cells are

exposed to Tg in a Ca2+-free buffer, the Orai1 channel is primed for Ca2+ entry and addition of

CaCl2 to the medium acts as a direct Ca2+ signal. Cells expressing CaRQ and sequentially

stimulated by Tg and CaCl2 had no observable morphology changes (Figure 5.9A). Following

from the reasoning applied to PM-localized CaRQ, CaRQ was localized to the ER, via an amino-

terminal fusion with Stim1 to take advantage of the local source of Ca2+ influx (Figure 5.9B).

Cells co-expressing Stim1-CaRQ and Orai1-Ceru, and sequentially stimulated with Tg and CaCl2

developed blebbing almost immediately after addition of CaCl2 to the medium for 2-3 minutes

(Figure 5.9C, D and Supplementary Movie 5.6). Blebbing was monitored through the CFP

channel which provided a clear PM outline, while puncta formation was observed through the

YFP channel. Tg stimulation alone was not observed to induce blebbing, presumably because,

while Tg acts at the ER, it creates a global rise in Ca2+. As before, the dominant negative

64

mutant, Stim1-CaRQ(T19N) was used as a control. Experiments were also performed with

Stim1-CaRQ using ChR2 as the Ca2+-mobilizing domain. Here, Ca2+ was present in the imaging

buffer and Tg was used to induce puncta formation. However, flickering illumination had no

effect on cell morphology, even though Stim1-CaRQ was incorporated into puncta as before.

This suggests that for Stim1-CaRQ, Ca2+ influx from ChR2 channels is not a sufficient source to

activate CaRQ; apparently in this case there was a strict spatial requirement for local Ca2+ influx,

at the level of protein complexes.

Figure 5.9: Localization of CaRQ to the ER with Stim1.

A: The percent of morphologically normal HEK293 cells that developed a blebbing morphology after Tg (dark bars) and CaCl2 (light bars) stimulus, under the indicated transfection and inhibitor conditions (n=3 experiments each with at least 10 cells). In the latter three conditions, cell were co-expressing Orai1-Ceru in addition to those constructs indicated. B: Cartoon demonstrating how SOCE enables activation of Stim1-CaR-Q: Orai1 (grey), Stim1-CaR-Q (white, blue and green) and the ER (light blue). C: HEK293 cells co-transfected with Orai1-Ceru and Stim1-CaR-Q were stimulated with Tg at 5 minutes and CaCl2 at 10 minutes. The CFP channel is shown because it best outlines the plasma membrane. White arrows indicate regions of blebbing. D: YFP channel of the cells in C-F before and after stimulus. Scale bars in C and D are 25 µm.

65

5.2.4 The Effect of Temporally Distinct Ca2+ Signals on CaRQ Activation

There was a tunable relationship between the duration of the ACh/nAChR-α4-induced Ca2+

signal and the duration of morphology changes observed with CaRQ (Figures 5.10, 5.11). An

apparent pattern emerged during the course of testing the ability of different Ca2+-mobilizing

domains to activate CaRQ. Those stimulants and modules that resulted in relatively long Ca2+

transients (approximately more than 1 minute, such as ionomycin and engineered SOCE) caused

blebbing to last approximately 2-3 minutes on average. However, those stimulants and modules

that resulted in relatively short Ca2+ transients (less than 1 minute, such as ATP and ACh)

routinely caused cell blebbing to last 5-15 minutes. This trend was not due solely to the

difference between CaRQ/Stim1-CaRQ and pLyn-CaRQ, because when pLyn-CaRQ-expressing

cells were stimulated by ionomycin the blebbing also typically had a duration of 2-3 minutes.

Figure 5.10: Blebbing duration relative to duration of Ca2+ signals.

HEK293 cells were co-transfected with TN-XL and pLyn-CaRQ/nAChR-α4 (A) or Stim1-CaRQ/Orai1-CFP (B) and stimulated respectively. The traces show the cytoplasmic YFP/CFP intensity ratios with the duration of blebbing indicated by the grey bars at the top of the graph.

To quantify this apparent relationship, morphology changes were induced by ACh/nAChR-α4 or

engineered SOCE in the presence of the TN-XL Ca2+ biosensor (Figure 5.10). In these cells,

there was a single-peaked Ca2+ transient (ACh-induced Ca2+ transients usually have one or two

peaks) with a full width at half maximum duration of approximately 20 seconds. Blebbing

morphology appeared shortly after the Ca2+ transient expired and lasted for 13 minutes in this

figure. In engineered SOCE-stimulated cells, the CaCl2-induced Ca2+ transient had a full width

at half maximum duration of approximately 1 minute and the blebbing lasted 2 minutes.

Blebbing duration was measured from the time the first bleb appeared until the last bleb

retracted. Clearly, the duration of Ca2+ signals between these two experiments was not the only

66

difference: CaRQ was localized to different subcellular compartments, and the amplitude of the

Ca2+ transient was larger for engineered SOCE-induced than ACh/nAChR-α4-induced transients.

Nevertheless, this suggested that there may be a tunable relationship between the duration of the

Ca2+ transient and the duration of the blebbing morphology.

Co-expression of the Ca2+-buffering protein PAV reduced the duration of ACh/nAChR-α4-

induced blebbing morphology (Figure 5.11). Ca2+ buffering proteins such as PAV and calbindin

are thought to be used throughout the cell to adjust the kinetic and spatial characteristics of Ca2+

transients (109, 153-154). When PAV (fused to mRFP) was co-expressed with nAChR-α4 and

the TN-XL Ca2+ biosensor the characteristic Ca2+ peak after ACh stimulation tended to be

elongated by several seconds with an increased shoulder duration. In the cells for the FRET-

traces shown, the full width at half maximum duration was 30 seconds without PAV and 2.5

minutes with PAV. When cells co-expressing pLyn-CaRQ and nAChR-α4 were stimulated with

ACh, with or without PAV, there was a significant difference in the duration of blebbing in those

cells (Figure 5.11). The average duration of blebbing after ACh stimulation was significantly

decreased from 12.2 ± 5.0 minutes without PAV to 8.3 ± 3.4 minutes (n=12 cells, P=0.033).

These results indicate that the Ca2+-responsive system presented here is sensitive to the duration

of Ca2+ signal and that by tuning the duration of that signal, the duration of the morphology

output can be tuned. However, it should be noted that there may be some effect of PAV on

spatial buffering of the nAChR-α4 Ca2+ signal that was not detected in these measurements.

Figure 5.11: Effect of PAV on blebbing duration.

A, B. Representative ACh-induced Ca2+ transient in HEK293 cells expressing nAChR-α4 and the TN-XL biosensor with or without PAV, respectively. C: Average duration of blebbing after ACh stimulation in cells expressing pLyn-CaRQ, nAChR-α4 and PAV where indicated. The data is presented as the mean ± standard deviation (n=12 cells, * indicates P = 0.033).

67

5.2.5 Regulation of Cell Migration by CaRQ

Cells co-expressing pLyn-CaRQ and ChR2 increased wound closure compared to negative

controls (Figure 5.12). Wound assays are simple tests to assess the migratory potential of cells

under different chemical, physical and genetic conditions and have been used to assess the effect

of Rho proteins on cell migration previously (90-92). Wound assays were prepared using

confluent HEK293 cells co-expressing ChR2 and pLyn-CaRQ. A light sensitive Ca2+-mobilizing

module was chosen because light could be easily “flickered” over 24 hours using a blinking iPod

with a blue background, in contrast to the challenges of delivering chemicals in a similar manner.

Figure 5.12: Light-dependent migration by CaRQ using wound assays.

A, B: Wound width immediately after, and 24 hours after scraping, and percent wound closure for the indicated conditions. Data is the mean ± standard deviation, n=9 wounds. C: Images from wound closure assays immediately after, and 24 hours after scrape. Scale bar is 100 µm.

Wounds were consistently approximately 0.75 mm in width after scraping. RhoA(DP)-YFP cells

closed wounds quickly, with over 60% closure in 24 hours. Cells co-expressing pLyn-CaRQ and

ChR2, when illuminated overnight with blue light (1s/15s) showed approximately 55% wound

68

closure. Negative control conditions such as pLyn-CaRQ(T19N) or incubation with Y-27632

tended to result in wound closures of approximately 20-25%. Bright field images were used to

measure wound width. For each wound, three images were taken at random positions along the

wound. For each image, 5 lines were drawn perpendicularly to the wound edge to determine the

mean width. For each case of the pair-wise comparisons between the second column’s condition

and the conditions of the third to sixth columns, there was a significant difference in wound

closure (P<0.001, n=9 wounds). While these results show that ChR2 was able to generate Ca2+

signals to contribute to pLyn-CaRQ’s wound closure, the LOVS1K/Orai1 Ca2+-mobilizing

module was also able to activate pLyn-CaRQ to achieve a similar extent of wound closure.

Cells that migrated into the wound tended to move as clusters, often with only 1-2 transfected

cells per cluster of 5-10 cells. This suggests that blebbing-induced migration can direct cell

migration in groups. Cells migrating by blebs are known to form cell adhesion junctions with

neighbouring cells, which could facilitate coherent migration (81). Clustered cell migration is

known to contribute to development, for example the migrating border cell cluster in Drosophila

(7, 155), and the invasion of some metastatic cells (156-158). A more full discussion of this

phenomenon in relation to the above results follows in section 5.3.

Cell migration in response to chemical signals (cytokines, growth factors, etc) was not shown

with the wound closure assay, but could be demonstrated with other standard assays such as

transwell migration assays (71, 79, 89). Several growth factor receptors, such as EGFR (159)

and VEGFR2 (10), generate Ca2+ transients as part of their downstream signaling, and their

ligands may be useful stimulants with CaRQ. VEGF-A/VEGFR2-induced Ca2+ transients were

able to activate pLyn-CaRQ blebbing (Figure 5.13). The VEGFR2 gene was successfully

inserted into cassette and fused with YFP (Figure 5.13A) and VEGF-A elicited a Ca2+ transient

in HEK293 cells when VEGFR2-YFP is expressed (Figure 5.13B). The Ca2+ transient recorded

was similar in amplitude, shape and duration to those reported in the literature (10). The Ca2+

transient due to VEGF-A/VEGFR2 was similar to others used as Ca2+-mobilizing modules (such

as ATP and ACh), suggesting that VEGFR2 may also be used as a Ca2+-mobilizing module

where VEGF-A is the input chemical signal. VEGF-A stimulation resulted in bleb formation

when VEGFR2-YFP and pLyn-CaRQ-CFP are co-expressed (Figure 5.13C-D). Bleb formation

began 2-3 minutes after 10 ng/mL VEGF-A was added to Ca2+-PBS. Blebbing lasted for

approximately 5 minutes in most cells observed. Blebbing was abrogated by pre-incubation of

69

cells with CDZ and Y-27632. Similarly, stimulation of cells expressing the dominant negative

mutation, pLyn-CaRQ(T19N)-CFP with VEGFR2 or pLyn-CaRQ-CFP alone did not cause

blebbing. This shows that VEGFR2 is a Ca2+-mobilizing module capable of CaRQ activation.

Figure 5.13: VEGFR2 is a Ca2+-mobilizing mode that activated CaRQ.

A. Fluorescence image of VEGFR2-YFP in HEK293 cells. B. Ratio of YFP/CFP intensities of TNXL Ca2+ biosensor in cells expressing VEGFR2-YFP. C. CFP images of HEK293 cells co-expressing pLyn-CaRQ-CFP and VEGFR2-YFP stimulated with VEGF-A at 5 minutes. White arrow heads show blebs and scale bars in A and C are 10 µm. D. Percent of cells blebbing after stimulation in the indicated condition. Data is mean ± standard deviation, n=3 experiments.

Cells co-expressing pLyn-CaRQ and LOVS1K/Orai1, VEGFR2, or nAChR-α4 showed increased

migration compared to negative controls in response to their respective cues in transwell assays

(Figure 5.14). Migration was reported as the migration index, the ratio of migrated fluorescent

cells (basal side of the insert) to the total number of fluorescent cells seeded onto the insert.

70

Cells were counted by taking five random images of each well (in 24-well plate format) and

computing the average (see section 3.5.2 for more details).

The average migration index for pLyn-CaRQ cells co-expressing LOVS1K/Orai1 was 0.53 ±

0.02, for VEGFR2 it was 0.50 ± 0.10 and for nAChR-α4 it was 0.42 ± 0.05, when stimulated

with the appropriate conditions. For pLyn-CaRQ(T19N) the migration indices under those same

conditions were 0.07 ± 0.02, 0.07 ± 0.04 and 0.04 ± 0.04, and were similar for other controls.

These assays show that cell migration can be increased using CaRQ and three of the tested Ca2+-

mobilizing modules. The condition where VEGF-A or ACh were present in both the apical and

basal chambers showed that the chemical gradient was a significant contribution toward the

migration of the re-programmed cells. However, the transwell assay is not a model of migration

toward a point source. Rather, these experiments should be taken as a second line of evidence

that a variety of stimulants can be used to signal re-programmed cells to increase their migration.

Figure 5.14: Multi-signal migration of CaRQ cells using transwell assays.

A. Schematic cartoon of the three different setups used to stimulate CaRQ cells. B-D. Migration index of cells using the transwell assays for the indicated conditions. For all three cases, the Ca2+-mobilizing module was transfected in all cells (B, LOVS1K/Orai1; C, VEGFR2; D, nAChR-α4). The data is the mean ± standard deviation, n=3 independent experiments.

71

5.3 Discussion

CaRQ was designed to enable Ca2+/CaM control over RhoA. Using Ca2+ as a signal to control

intracellular proteins has some obvious advantages and drawbacks. Ca2+ is a desirable second

messenger to manipulate because it can be delivered to cells with spatiotemporal control (12, 29,

160), many technologies have been developed to visualize changes in intracellular Ca2+ (15-18,

121-122, 127), and for CaM in particular, there is a large library of structurally defined target

peptides on which designs could be based (14, 114, 117, 119). Ca2+ is not the only second

messenger that has been controlled in living cells. A variety of cell membrane-permeable,

photo-caged second messengers and small molecules exist (160). ChR2 has been engineered for

photoactivatable generation of cAMP and DAG/IP3 (28), and endogenous generation of cAMP

has been used to re-wire morphology (131). Perhaps the primary drawback of Ca2+ signaling is

its ubiquitous use in the cell. Experiments must be carefully designed to ensure that observed

effects are due to the engineered protein and are not “Ca2+ effects”. One objection may also be

that in order to ensure a particular Ca2+ stimulus has no adverse or secondary effects on the cell,

one would need to monitor a nearly endless number of proteins and pathways. However, this

blanket statement betrays the fidelity of Ca2+ sensitive proteins for Ca2+ signals of different

spatial patterns, amplitudes, frequencies and time-dependent shapes. A given Ca2+ signal only

has the potential to activate select Ca2+-dependent processes (8, 97).

In this work, the ubiquitous nature of Ca2+ can also be one of its greatest strengths: the extensive

library of proteins that exists to mobilize Ca2+ in different cells and by different stimuli means

that a number of signals can be used to control engineered systems, although, as noted above,

one would not expect all Ca2+ signals to be effective. Here, the work is driven by a specific

objective, and some off-target effects, depending on the eventual re-programming application,

could be acceptable trade-offs. As a tool to study Rho protein biology, Ca2+-based systems are

not the most appropriate, given existing alternatives (5-6). On balance, the benefits of Ca2+-

based engineered systems outweigh the potential for undesirable effects for this application.

Exploring Ca2+ signals and Ca2+-realted proteins as a broad design space can develop insights

into how Ca2+ signaling functions in nature. The ability to re-combine exogenous Ca2+-

mobilizing proteins such as VEGFR2 and nAChR-α4 with an engineered Ca2+-responsive protein

chimera shows that Ca2+ as a second messenger is capable of relaying signals in abnormal

72

environments and may contribute to its ubiquitous role throughout evolution, an idea that has

also been suggested elsewhere (161). Specific insights can also be gained into some of the

proteins used here. For example, the inability to activate Stim1-CaRQ with ChR2 excitation

suggests that the Ca2+ signals generated by Stim1-Orai1 oligomerization only have large

amplitudes (that is, sufficient to activate CaRQ) in a local area, consistent with the initial

characterization of LOVS1K/Orai1 (12).

The spatial sensitivity of CaRQ was not a phenomenon observed with CaM-RhoA(DP). The

cytoplasmic CaM-RhoA(DP) appeared to mediate Ca2+-dependent bleb retraction for stimuli

that, for CaRQ, only had an effect when CaRQ was localized to the plasma membrane. Making

direct comparisons between the overall efficiency of the two proteins is difficult because they are

based on different mechanisms and have different affinities and accessibility between CaM and

the intended CBP’s. As well, CaRQ and CaM-RhoA(DP) mediate morphology switches in

opposite directions. The cells used here do not bleb as their wild type morphology, so the overall

cytoskeletal arrangement may be biased against blebbing: for example myosin light chain

phosphatase is constitutively active, and only when ROCK is activated does the phosphatase stop

de-phosphorylating myosin light chain. Therefore, a relatively higher Ca2+ concentration, within

the range of CaM lobe’s differential affinities for Ca2+ (162), may be required to mediate bleb

formation than retraction with all else equal. Further, different CaM-CBP interactions, through

differential affinities and kinetics between Ca2+ and the four EF-hands in CaM, have been shown

to confer spatial sensitivity to CaM binding (13, 163). There are many potential factors that

could contribute to differential spatial sensitivity between CaM-based engineered proteins.

In addition to spatial sensitivity, CaRQ was activated by Ca2+ signals of different shapes,

amplitudes and duration. The extent and duration of morphology changes could be tuned by

spatial localization and, for the duration of ACh/nAChR-α4-induced blebbing, by co-expression

of the Ca2+-buffering protein PAV. In this case elongating the Ca2+ signal had the counter-

intuitive effect of shortening the observed duration of blebbing. The biochemical basis for this

phenomenon is unclear. The blebbing morphology arises due to phosphorylation downstream of

RhoA, so the effect of RhoA-ROCK activation can last longer than that RhoA-ROCK activity

itself. Buffering proteins also massage the spatial characteristic of a Ca2+ signal (109-110), so

some reduction in the amplitude of the PM-localized Ca2+ signal with cytoplasmic PAV

overexpression may have resulted in less CaRQ activity, relative to an un-buffered system.

73

However, as an overall observation, it appears that changing various parameters of Ca2+ signals

(such as their amplitude, shape, localization and buffering) is translated into changes in the

parameters of morphology change such as the duration and extent of blebbing. Furthermore, the

translation is often predictable and tunable. For example, by varying the amplitude of Ca2+

influx while holding signal shape and protein localization constant, the fraction of cells with

morphology change was controlled (Figure 5.5). Similarly, by changing the buffering of the Ca2+

signal while holding all other parameters constant, the duration of morphology change was

altered (Figure 5.11).

Repeated activation of CaRQ over 24 hours increased wound closure in a scratch wound assay,

demonstrating that inducing the blebbing morphology increased the ability of CaRQ-expressing

cells to migrate. Blebbing morphology and amoeboid-like cell migration have been observed in

a number of systems, such as D. Rerio germ cell development (81-82), metastatic cell invasion

(71, 74, 76), and some kinds of leukocyte migration (80). The blebbing morphology contributes

to migration through a cyclical process of increased cytoskeletal tension, bleb formation, bleb

growth and “tumbling”, and bleb retraction with transmembrane anchoring leading to forward

locomotion (see section 2.2.2). E-cadherin has been reported as a cell-cell transmembrane

anchor that translates tumbling to locomotion (81). The role of integrins in blebbing-based

migration is unclear: sometimes this mode is considered to be “integrin-independent”, but some

studies have shown that certain integrins such as β1 play a role in anchoring migrating, blebbing

cells to the substrate, albeit with reduced adhesion compared to non-blebbing cells (80, 164-165).

Since integrins, like cadherins, are linked to the cytoskeleton (166), they may serve an analogous

role in translating tumbling to locomotion. Local friction has also been suggested as a possible

way for blebbing cells to realize forward motion (70). In this model, a growing bleb becomes

lodged in an open space within the extracellular matrix. Non-specific attractive forces between

PM glycoproteins and transmembrane proteins and the matrix, or friction, anchors the cell in

place as the bleb retracts, thereby pulling the cell body forward. Clearly for this mechanism to

be practicable, the adhesion between the rest of the cell body and its substrate must be low. This

is most likely not a significant contribution to migration in the 2D model used here. In general

blebbing has not been widely studied in 2D environments, so many of the mechanisms discussed

here have been extrapolated from studies performed in 3D environments.

74

One feature of blebbing-based migration is that it was achieved by activation of dominant

positive RhoA, however overexpression of dominant positive Rho proteins are usually associated

with a reduction in cell migration (167-168). The salient feature of these experiments is that in

the given context, activation of CaRQ establishes the blebbing morphology, which is known to

increase cell motility (71-74, 76, 82). In mesenchymal migration, localized activation of

dominant positive Rho proteins can increase cell migration (5-6).

In wound assays, cells tended to migrate in clusters, with sometimes as little as 1-2 transfected

cells per cluster of 5-10 cells. Collective cell migration has been studied in the migration of cells

during development, physiological wound healing, and cancer (156-158, 169-171), and can

occur in a number of topologies, with cells moving as sheets, strands, or clusters. There are four

aspects of collective cell migration: cell-cell adhesion, supracellular polarity, force transduction,

and extracellular matrix remodeling.

• Coherent cluster migration is mediated by the same molecular mechanisms that adhere non-

motile cells together: tight and gap junctions, cadherins and Ig-superfamily cell adhesion

molecules (156). E-cadherin in particular has been implicated in several studies (157, 170-

171). During migration actomyosin tension at cell-cell junctions is reduced to prevent

dissociation: recently DDR1, which associates with E-cadherin, has been implicated in

reducing local tension by driving the localization of RhoE, a ROCK antagonist (157). The

prevalent role of E-cadherin in collective cell migration and blebbing-based migration (81)

suggests that it may be an important regulator of collective blebbing-based migration.

• The migrating cluster takes on a supracellular polarity with cells driving the migration

usually from the front or rear of the group (7, 155-156, 169-170). Cell morphology and gene

expression drive the polarity: physiologically, extracellular cues can induce this polarity such

as EGF, SDF or FGF (155, 169-170), depending on the context. Here, the polarity is induced

by the combination of CaRQ/Ca2+-mobilizing module expression and application of light or

chemical stimulants.

• Force transduction and actin regulation occur within individual cells in migrating clusters

similar to singly migrating cells, however there are additional communication mechanisms

between cells. In a scratch wound assay, migrating “pioneer” cells have been shown to pull

“follower” cells, 3-5 cell widths away, forward (169). Pioneer cells expressing FGFR were

75

stimulated by FGF and follower, FGFR- cells, which migrate randomly with FGF

stimulation, were apparently dragged along mechanically by VE-cadherin connections. The

ratio of 1 pioneer cell to approximately 3-5 follower cells is similar to what was noted for

CaRQ-induced migrating clusters. Other mechanisms of coherent force transduction appear

to exist, since collective migration in Drosophila border cells is dependent on JNK signaling,

through an unidentified autocrine or juxtacrine signaling factor (7, 155).

• ECM remodeling by matrix metalloproteinases is important for large clusters migrating in

vivo, or in invasion assays (158). In wound assays, cells are not presented with an

environment that needs to, or can be, remodeled and migrating clusters here can follow a

path of least resistance. In transwell assays, cells that had migrated past the filter were not

clustered but individual. Collectively migrating cells have been shown to dissociate when

presented with barriers insurmountable to the group (for example, by downregulating E-

cadherin) (158, 171).

The cell migration assays used here demonstrate that CaRQ-induced blebbing can increase the

migration of a certain population of cells, in this case HEK293 cells, but likely this is applicable

to several other epithelial-like cells, such as HeLa and CHO, in which blebbing was also induced

by CaRQ activation. In the wound assay, the light stimulus was applied from beneath the glass

substrate, so the light was projected perpendicularly to the plane of migration. Cell migration

into the wound was not directed by the illumination but rather by the fact that cells at the wound

edge were more likely to migrate into the open space than to regress into the cell monolayer. In

this case the stimulus provided a permissive signal that increased migratory potential (e.g., the

likelihood for cells to undergo a random walk) without directionality. In the transwell assays,

the light and chemical gradients were applied parallel to the direction of cell migration, however

the extent to which the stimuli served as directional cues rather than permissive signals is

unclear. Cell migration past the transwell filter requires a cell to squeeze through pores; the

thickness of the filter is approximately one cell width. The requirement for sustained migration

along a chemical gradient is therefore unclear. Experiments were the chemical gradient was

abolished by adding stimulant in the apical chamber reduced the migration index, suggesting that

the chemical gradient did contribute to the migration. Still, the transwell assay is not a model of

migration toward a point source. Rather, these experiments showed that a variety of stimulants

can be used to signal these re-programmed cells to increase their migratory potential.

76


In this chapter two CBP’s were fused with RhoA (one embedded within the structure of RhoA,

and one as an amino-terminal fusion) and CaM to create CaRQ. CaRQ regulated bleb formation

in response to several Ca2+ signals, and prolonged activation of CaRQ increased the tendency for

cells to migrate.

1. A Ca2+-activated RhoA chimera, CaRQ, was designed with an embedded IQp between

amino acids 49 and 50 and amino-terminal fusion of MLCKp and CaM.

2. On Ca2+ influx, CaRQ localization with RBD increased, and blebs were formed for

several minutes (in separate experiments). The Ca2+ EC50 for bleb formation was 27 µM.

The direction of the Ca2+ switch was changed by changing the embedded CBP.

3. CaRQ was activated by several different Ca2+-mobilizing modules, including ATP,

ACh/nAChR-α4, blue light/ChR2 or /LOVS1K, and VEGF-A/VEGFR2, however Ca2+

signals with different spatial patterns had different effects on CaRQ-activated blebbing.

4. The duration of a Ca2+ signal affected the duration of blebbing morphology that was

observed. Tuning the Ca2+ signal with PAV affected the duration of blebbing.

5. Prolonged activation of CaRQ by light resulted in increased wound closure in wound

assays, and activation by light, ACh and VEGF-A resulted in increased migration in

transwell migration assays.

In relation to the research objectives outlined in Chapter 1, the work in this chapter addresses

each of the three objectives. 1. A chimeric protein of CaM, RhoA and two CBP’s was created

using computational modeling and rational design that controlled cell morphology, bleb

formation in particular, in response to changes in intracellular Ca2+. Tuning the chimera or the

Ca2+ signal had predictable changes on the resultant morphology changes. 2. Control over bleb

formation was achieved by several exogenous signals including blue light, ACh and VEGF-A

using ChR2 or LOVS1K, nAChR-α4 and VEGFR2, respectively. 3. Prolonged activation of

CaRQ created a “permissive” signal that allowed blebbing cells to migrate into wounds or

through porous transwell filters.

77

6 RACer: A Ca2+-Sensitive Rac1 Chimera


Reprinted (adapted) with permission from ACS Synthetic Biology, E-publication ahead of print

April 28, 2012, DOI: 10.1021/sb3000172. Mills, E., Pham, E., Nagaraj S. and Truong, K.

“Engineered networks of natural and synthetic proteins to control cell migration,” © 2012,

American Chemical Society.


CaRQ is a Ca2+-responsive chimera that was combined with light- and chemical-sensing Ca2+-

mobilizing modules and was able increase cell migration in response to light and chemical cues.

CaRQ’s ability to increase motility is based on actomyosin contractility and blebbing, which is

used in several physiological and pathological contexts (72-74, 81-82). However, lamellipodia-

based, or mesenchymal, cell migration may be more appropriate in some circumstances such as

with highly adhesive cells or matrices (70) and may be used in a wider range of cell types (37)

than the epithelial-like set of cells used with CaRQ. Therefore, a compliment to CaRQ is desired

where migration is based on activation of Rac1, rather than RhoA, using actin polymerization

and lamellipodia formation. As with CaRQ, modular Ca2+ activation and migration in response

to a variety of exogenous stimuli are still objectives.

The five specific aims for the work in this chapter were drawn from the overall research aims:

1. Determine a suitable design for a Ca2+-activated Rac1 chimera.

2. Characterize the Rac1 chimera in terms of its overall Ca2+ affinity, response to Ca2+

transient-inducing chemicals, and characteristic morphology changes.

3. Investigate whether the Ca2+-mobilizing modules used to activate CaRQ can be used with

the Ca2+-activated Rac1 chimera.

4. Determine whether activation of the Ca2+-activated Rac1 chimera affects cell migration,

characterize the effect if there is one, and compare this to the amoeboid-like cell

migration observed with CaRQ.

5. Apply the design of the Ca2+-activated Rac1 chimera to the related cytoskeletal regulator

Cdc42.

78


Ca2+-activated Rac1 chimera (Figure 6.1).

Figure 6.1: Overview of main fusion proteins created in Chapter 6.

Schematic layout of the fusion proteins are shown on the left and their names as used in the text are given on the right. All of the Rac1 or Cdc42(47-188) fragments in the figure have the Q61L mutation. CFP-bearing mutants of RACer and RACer(T17N) were also created but have been left off the figure for simplicity.

6.2 Results

The experimental results of this chapter are described here. The rational design of the Rac1/CBP

chimera is described following from the design of CaRQ (section 6.2.1). The final designed

chimera, RACer, was characterized in a similar manner as CaRQ, by cellular pull-down assay

and by observation of changes in characteristic Rac1-dependent morphologies, in this case,

lamellipodia formation (section 6.2.2). A variety of Ca2+-mobilizing modules were used with

RACer including LOVS1K/Orai1 and nAChR-α4, and the morphology responses to these stimuli

were also characterized (section 6.2.3). As with CaRQ, prolonged, repeated activation of RACer

using various Ca2+-mbilizing domains was tested for the ability to regulate cell migration in

wound closure and transwell assays in response to light, VEGF-A and ACh (section 6.2.4).

Finally, the design of RACer was applied to a related Rho protein Cdc42. The ability of the

CaM-Cdc42 chimera to regulate characteristic Cdc42-dependent morphologies, in this case

filopodia, was assessed (section 6.2.5).

79

6.2.1 Design of RACer, a Ca2+-Sensitive Rac1 Chimera

A Ca2+-sensitive Rac1 chimera was created by applying the embedded IQp design of CaRQ to

Rac1 (Figure 6.2), although several other design strategies were also pursued (Appendices G and

H). The intended switching behaviour of CaM between IQp in the absence of Ca2+ toward

MLCKp on Ca2+ loading should function in a similar manner here. As well, the genetic and

structural similarity between RhoA and Rac1 should mean that the location of peptide insertion

site will function in a similar manner between the two GTPases. While there are some

differences between the surfaces of Rac1 and RhoA that are used to bind effector proteins

(comparing, for example the structures of Rac1/PKN, PDB 2RMK, (173) with RhoA-ROCK,

PDB 1S1C (174)), the locations of the switch regions in the two GTPases are the same, so the

peptide insertion point that was successful for designing a Ca2+-sensitive RhoA represents the

best starting point for this design. The YFP was moved to the amino terminus to allow the

CAAX motif to mediate membrane insertion, because carboxy-terminal YFP fusions have been

shown to limit Rac1/Cdc42 activity (175).

When the Rac1/CBP chimera, version 1, was created and expressed in cells, the fluorescence

localization corresponded mainly to the nucleus (Figure 6.2B). Since Rac1 contains a functional

NLS (176), the increased molecular mass of the chimera may have disturbed the balance

between nuclear localization and nuclear exclusion. The molecular mass of the Rac1/CBP

chimera would be greater than 65 kDa (by summing the masses of YFP, Rac1 and CaM),

whereas the nuclear pore cutoff is usually between 40 and 60 kDa in most cells (177). Rac1

localization by physiological regulation is not a required, so the chimera was constitutively

localized to the PM using the pLyn tag to create Rac1/CBP chimera, version 2. For this version

of the chimera many cells had lamellipodia before Ca2+ stimulation (Figure 6.2C). In other

reports of engineered Rac1, overexpressing the Q61L mutation was found to activate

endogenous Rac1, reducing the apparent dynamic range of the engineered switch (5). This was

because overexpressed GTP-bound Rac1 overwhelms the pool of cellular GAP’s (the role of a

GAP being to accelerate GTP hydrolysis), acting as a GAP sink and preventing GAP’s from

regulating endogenous GTPases. To correct this, specific mutations to reduce GAP-binding

were introduced (E91H, N92H), and these were also made here to reduce the chimera acting as a

GAP sink. In this version of the chimera, cells were less likely to show lamellipodia before

stimulation; this version was named RACer for Rac1 activated by Ca2+.

80

Designing a Ca2+-sensitive Rac1 pre-supposes that Rac1 is not Ca2+-sensitive already. There

have been reports that in some cell lines, isoforms of Ca2+-dependent kinases can activate Rac1

leading to indirect Ca2+-activation of Rac1 by phosphorylation of its GDI (178). However for

the cell lines used here stimulating cells with ionomycin, with or without Rac1 overexpression,

did not lead to any observable lamellipodia formation (see controls below in section 6.2.2.).

Figure 6.2: Design of RACer.

A. Schematic cartoon of RACer in the absence (left) and presence (right) of Ca2+. B-D. Iterations of the RACer design with only the Q61L mutation (B), constitutive membrane localization (C), and introduction of E91H, N92H mutations to reduce GAP sinking from overexpression of dominant positive Rac1. The scale bar is 10 µm.

6.2.2 Characterization of RACer

The RACer chimera had less Rac1 activity before stimulation with Ca2+ than Rac1(Q61L)

(Figure 6.3). Rac1 activity was measured in two ways – by counting the number of cells with

lamellipodia on cells expressing YFP, YFP-Rac1(Q61L), RACer and a dominant negative

RACer, RACer(T17N), and, in a different cell population, by computing the PC of colocalization

between these constructs and a probe containing the PBD (88) from p21-activated kinase, PBD-

mRFP-GST. As with CaRQ, mechanical extraction of RACer and PBD-mRFP-GST from E.

Coli was unsuccessful. Using both morphology and colocalization methods, RACer

81

demonstrated significantly different Rac1 activity than Rac1(Q61L) (P<0.001 by morphology

method and P=0.032 by PC method, n=3 and 10 experiments respectively, with at least 10 cells

considered in both cases). There appeared to be more Rac1-like activity of RACer at basal Ca2+

as measured by the PC method than by morphology. However since the primary motivation is to

regulate morphology and migration, a slightly higher PC colocalization did not merit further

modifications to RACer to reduce the colocalization.

Figure 6.3: Pre-stimulation effects of RACer.

A. Percentage of cells with lamellipodia for the indicated construct, and B average PC colocalization between PBD-mRFP-GST and the indicated construct before stimulation. For A and B, the data is the mean ± standard deviation n=10 cells. C. Representative YFP and RFP channel images showing the extent of colocalization between the indicated construct (YFP channel) and PBD-mRFP-GST (RFP channel). Scale bar in C is 10 µm.

Ca2+ stimulation by ionomycin of cells co-expressing RACer and PBD-mRFP-GST increased the

PC colocalization between the two signals, and for cells expressing RACer alone, Ca2+

stimulation resulted in the formation of lamellipodia (Figure 6.4 and Movies 6.1 and 6.2). Ca2+-

stimulation of RACer-expressing cells increased colocalization between the PBD-mRFP-GST

probe and RACer by 28% and RACer(T17N) by less than 5%. CFP was used as a volume

indicator but there were no noticeable changes in YFP-CFP correlation after stimulation.

82

Morphology changes were quantified by analyzing the changes in cell area upon Ca2+

stimulation, rather than counting the number of cells that developed lamellipodia. Relative

changes in cell area have also been used to report on the effect of engineered Rac1 chimeras (5).

Area changes were calculated by thresholding images in ImageJ and the image of a cell at time 0

was used as the baseline, or mask, to which subsequent images in that time-lapse experiment

were compared. To easily compare between experimental conditions, a binary judgment was

made on whether a cell had or did not have an area change. A cell was considered to have an

area change when the peak fold area change was at least twice the maximal fold area change

fluctuation in the preceding control period.

Ca2+-stimulation of RACer-expressing cells resulted in lamellipodia formation in 85% of cells

with an average peak area of 1.3-fold ± 0.06-fold basal (Figure 6.4B-G). Representative cells are

shown (RACer, Figure 6.4B and RACer(T17N), Figure 6.4D) with ionomycin stimulation after 5

minutes. For RACer the average area change plateaued at 1.25-fold ± 0.12-fold basal while there

was no significant change for RACer(T17N) (Figures 6.4C and 6.4E). The error bars become

large towards the later time points for the average analysis of RACer fold area change because in

some cells the morphology change continued for the duration of the imaging window, while for

others the morphology change ceased after 5-10 minutes and area returned to near baseline. This

was likely due to differences in duration of the ionomycin-induced Ca2+ transient between cells.

Cells expressing YFP or YFP-Rac1(Q61L) were insensitive to ionomycin Ca2+ stimulation, as

were cells treated with CDZ (Figures 6.4F and G). Y-27632 treatment had no effect on the

likelihood of morphology change or that maximal extent of area change. This was expected

because Y-27632 inhibits ROCK, a RhoA effector not implicated in mediating Rac1 effects.

Blebbistatin, a myosin ATPase inhibitor (179-180), appeared to have an effect on the likelihood

of area changes, but the difference was not significant; there was no effect on the peak area

change in those cells which did have an area change. Blebbistatin is a non-competitive myosin

ATPase inhibitor and therefore affects the ability of myosin to crosslink with actin. However,

there have been no reports of blebbistatin interfering with actin polymerization, and indeed

blebbistatin did not affect lamellipodia formation on activation of engineered PARac (5).

84

Figure 6.4: Effect of ionomycin on cells expressing RACer.

A. YFP and RFP channel images of cells expressing RACer or RACer(T17N) and PBD-mRFP-GST with calculated PC. B, C. Time-lapse images of RACer-expressing cells stimulated with ionomycin at 5 minutes and fold area change reported over time, respectively. D, E. Similar data shown for RACer(T17N). F, G. Percent of cells with area increase, and peak fold area change for the indicated control conditions. Data is the mean ± standard deviation. In C and E n=10 cells and in G and H, n=3 experiments with at least 10 cells. Scale bar is 10 µm in A, B and D.

The Ca2+ EC50 for RACer was 24 µM (Figure 6.5). This experiment was conducted in the same

manner as described above for the analogous experiments used to characterize CaRQ. The

percent of cells with an area increase was used as the output. This value, rather than the peak

fold area change, or the integral of the fold-area change versus time curve, was used because it

most closely resembles the measure used to characterize CaRQ. As a result, the Ca2+ EC50 value

for RACer was similar to the Ca2+ EC50 value for CaRQ of 27 µM. This result was not

surprising given that the CaM gene and CaM-binding peptides were not changed between CaRQ

and RACer, and only minimal changes were made to the fusion layout. While RhoA and Rac1

bind their effectors in different arrangements, differences in efficiency between the two chimeras

would result in changes to the maximal and minimal values of morphology changes, rather than

the Ca2+-sensitivity. The similarity of these values suggests that the Ca2+ EC50 of CaRQ and

RACer may be tunable by choice of CaM-binding peptide or mutations within the CaM gene.

Figure 6.5: The Ca2+ EC50 of RACer was 24 µM.

The sigmoid expression is given (top left), with the modeled EC50 value (top right) and the quality of fit indicated by r2. Data are the mean ± standard deviation, n=3 for each condition.

85

RACer mediated Ca2+-induced morphology changes in COS7, HeLa and RAW264.7 cells

(Figure 6.6). One motivation for developing RACer was that CaRQ only caused morphology

changes in a set of epithelial-like cell lines, whereas a Rac1 chimera should control morphology

in more cell lines (37). The ionomycin response was mostly consistent with earlier observations:

a small, but repeatable, area increase was observed in all three cell types. The extent of the area

increase was less than with HEK293, likely a result of the high protein expression levels known

to occur in HEK293 cells (181). In HeLa cells there was filopodia outgrowth in addition to

increase in total cell area, which is likely due to the crosstalk between Cdc42 and Rac1 pathways

(68). While the RACer-mediated Ca2+-induced changes in these other cell lines were not as

extensive as they were in HEK293, taken together, they show that RACer has a potentially broad

applicability to several cell types, which may be optimized on an application-dependent basis.

Figure 6.6: Effect of Ca2+ stimulation of RACer in various cell lines.

A. Summary of morphology changes for cells expressing RACer or RACer(T19N) in COS7 (B), RAW264.7 (C) and HeLa (D). In B, a fold area change versus time graph is also presented for representative COS7 cells expressing RACer (black line) and RACer(T17N) (grey line) stimulated with ionomycin after 5 minutes. Data is the mean ± standard deviation, n=3 experiments. White arrows highlight changes and white dashed lines show original cell outline. Scale bars are 10 µm and 5 µm in inset.

86

6.2.3 The Effect of Ca2+-mobilizing Modules on RACer Activation

Endogenous ATP receptors, nAChR-α4 and LOVS1K/Orai1 were each able to activate RACer

(Figure 6.7 and Movies 6.3 and 6.4), although with different durations of morphology response

and peak fold area changes. These three stimuli represent one of each of the main classes of

Ca2+-mobilizing modules used in this work: endogenous (ATP), exogenous (ACh/nAChR-α4)

and engineered or synthetic (blue light/LOVS1K/Orai1). With RACer the effect of spatial

localization of Ca2+ signal relative to the chimeric protein was not considered because Rac1-

dependent morphologies were only observed when Rac1, and therefore likely RACer, was

localized to the PM. ATP, ACh and blue light stimulation caused area increases in 61.1% ±

5.6%, 75.6% ± 12.3% and 56.1% ± 12.2% of cells, respectively, compared to 10.8% ± 5.6%,

16.7% ± 9.6% and 8.3% ± 7.3% of cells, respectively, for the T17N mutant. These differences

were each significant between RACer and RACer(T17N): P=0.003, 0.020 and 0.032 for ATP,

ACh/nAChR-α4 and blue light/LOVS1K, respectively. Data for ionomycin is reproduced from

the initial characterization (Figure 6.4) for comparison. These results show that the various Ca2+-

mobilizing domains used to activate CaRQ can also be used to mobilize Ca2+ to activate RACer.

However, there were noticeable differences in the duration of lamellipodia formation and the

maximal extent of the lamellipodia formation, and this appeared to be dependent on the

particular Ca2+-mobilizing domain used.

Cells co-expressing LOVS1K/Orai1 and RACer appeared to be somewhat less likely to form

new lamellipodia after illumination than with ionomycin stimulation, but the differences were

not significant (P=0.171). In cells that did form new lamellipodia during stimulation, the

magnitude of the peak cell area change was similar (1.30 ± 0.10 fold area change for blue

light/LOVS1K versus 1.29 ± 0.06 fold area change for ionomycin). The magnitude of area

change in the first 20-25 minutes was small, and in many cells was indistinguishable from signal

noise. However, the Ca2+ signals due to repeated LOVS1K activation are known to grow in time

over up to 1 hour when it was tested with other engineered Ca2+-sensitive proteins (12). With

continued illumination, the magnitude of area change increased, and in the cell shown, the area

increased almost linearly between 20 and 35 minutes after the onset of illumination. When

illumination ceased, the cell area increase ceased, then decreased and plateaued at approximately

30% greater than the initial cell area. When blue light illumination of LOVS1K ceases,

LOVS1K dissociates from Orai1 after 30-60 seconds (12), which is closely related to the thermal

87

relaxation of LOV2 excitation (34), which prevents further Ca2+ influx through Orai1. These

experiments suggest that for RACer and LOVS1K/Orai1, the duration of morphology change is

closely correlated to the duration of increased cellular Ca2+.

Figure 6.7: RACer can be activated by three distinct Ca2+-mobilizing modules.

A. Schematic diagram of how blue light activates RACer with the LOVS1K/Orai1 Ca2+-mobilziing module. B. Percentage of cells, co-expressing RACer (dark bars) or RACer(T17N) (light bars) and the relevant Ca2+-mobilizing module, with an area increase after stimulation with the indicated signal. C. Peak fold area change for cells with area increase from B. D. Images of

88

representative cells co-expressing RACer, LOVS1K and Orai1 and stimulated with flashing blue light for 30 minutes. E. Representative images of cells co-expressing RACer and nAChR-α4 stimulated with ACh at 5 minutes. F. Fold area change versus time graphs for the cell shown in D (left graph) and in E (right graph). Data is mean ± standard deviation, n=3 experiments (B and C). Scale bar is 10 µm. Dashed blue bar represents the duration of blue light illumination, and arrow shows when ACh was added.

Cells expressing RACer and stimulated with ATP or cells co-expressing RACer and nAChR-α4

and stimulated with ACh also appeared to be somewhat less likely to have an area increase than

cells stimulated with ionomycin, but the differences were not significant (P=0.055 and 0.933 for

ATP and ACh/nAChR-α4, respectively). In cells with an area increase, the magnitude of the area

increase was smaller than that with ionomycin (1.06 ± 0.01 for ATP and 1.12 ± 0.02 for ACh,

which were significantly different than the changes for ionomycin with P = 0.046 and 0.047,

respectively). Further, the duration of the area increase was shorter than with ionomycin or blue

light/LOVS1K: the full width at half maximum of the fold area change was approximately 1

minute for the representative ACh/nAChR-α4-stimulated cells shown here. The duration of

morphology change in these cells was close to the duration of the ACh/nAChR-α4-induced Ca2+

transient in these cells (see Figures 5.10 and 5.11). Taken together with the results from the blue

light/LOS1K experiments above, it appears that for RACer the duration of the morphology

change closely tracked the duration of the Ca2+ transient. This relationship is in contrast to the

behaviour of CaRQ where morphology changes often lasted for several minutes beyond the

expiration of the Ca2+ transient. This result will be discussed further below (section 6.3); it

shows that engineered Ca2+-dependent proteins, even where the proteins are structurally and

functionally similar, can have substantially different relationships with their input Ca2+ signals.

6.2.4 Regulation of Cell Migration by RACer

The role of RACer in cell migration was assayed using wound closure assays and transwell

migration assays in the same way as with CaRQ. The cellular morphology changes due to

RACer and CaRQ are different, and suggest different migration modes (mesenchymal and

amoeboid-like, respectively). These two migration modes are affected differently by several

factors including substrate adhesiveness or surface topology. For example, increased substrate

adhesiveness could impede amoeboid-like migration more than mesenchymal migration (70, 80),

therefore these parameters have not been actively altered in any way.

89

Flickering illumination of cells co-expressing RACer and LOVS1K/Orai1 over 24 hours resulted

in an average wound closure of 52.1% ± 4.6% (Figure 6.8). As with the previous wound assays

with CaRQ, the average wound width immediately after wounding was approximately 0.75 mm,

and there was a narrow range for this value showing that the initial wounds were created evenly.

Wound closure was measured using perpendicular lines drawn between wound edges as

described above (section 3.4.1). In all cases, except that of blebbistatin treatment, there was a

significant difference in the wound width at 0 and 24 hours (significant P values ranged from

0.004 for YFP-Rac1(Q61L) to 0.024 for RACer(T17N)). These differences were due to cell

proliferation and some background migration of cells at the wound edge.

Figure 6.8: Light-dependent migration by RACer using wound assays.

A, B: Wound width immediately after, and 24 hours after scraping, and percent wound closure for the indicated conditions. Data is the mean ± standard deviation, n=9 wounds. C: Images from wound closure assays immediately after, and 24 hours after scrape. Scale bar is 100 µm.

90

RACer wound assay control conditions were based on the control conditions used during the

initial characterization of RACer (Figure 6.8). Mostly the results of the wound assay control

experiments were consistent with those of the initial characterization. For illumination of

wounds co-expressing YFP-Rac1(Q61L), or RACer(T17N), and LOVS1K/Orai, the average

wound closures were 20.4% ± 3.5% and 17.9% ± 3.2%, which were significantly different than

RACer with LOVS1K/Orai1 (P=0.025 and 0.004, respectively). Similarly, lack of illumination

for RACer/LOVS1K/Orai1 co-expressing cells, or illumination of cells expressing RACer alone

resulted in average wound closure of 25.2% ± 3.7% and 19.4% ± 2.0% which were significantly

different than with illumination and co-expression (P=0.011 and 0.007, respectively).

For cells co-expressing RACer/LOVS1K/Orai1 and illuminated with blue light treated with the

ROCK inhibitor Y-27632 the average wound closure after 24 hours was 35.8% ± 3.8%. This

was the only condition which was not significantly different than the base condition of

RACer/LOVS1K/Orai1 co-expressing cells illuminated with blue light (P=0.052). The closeness

of this P value to the pre-set α value obscures the meaning of the result. Earlier experiments

showed that Y-27632 had no effect on lamellipodia formation (Figure 6.4), however migration

involves more cellular processes (rear cell contraction, focal adhesion formation, etc) than

lamellipodia formation alone, with which Y-27632 may interfere. For cells treated with

blebbistatin instead of Y-27632 the wound closure after 24 hours was 15.8% ± 6.0% which was

significantly different than without inhibitor (P=0.009). Again, where blebbistatin had little

effect on lamellipodia formation, it appeared to strongly inhibit wound closure, and therefore cell

migration, in these experiments.

Cell populations co-expressing RACer and an appropriate Ca2+-mobilizing module had higher

migration indices in transwell assays than control conditions (Figure 6.9). Transwell migration

assays were also used with RACer as a complimentary assay for the affect of blue light/LOVS1K

on cell migration, and to assess the affect of VEGF-A and ACh on RACer-mediated cell

migration. Cells co-expressing RACer and a Ca2+-mobilizing module had migration indices of

0.45 ± 0.03 (blue light and LOVS1K/Orai1), 0.35 ± 0.04 (VEGF-A and VEGFR2) and 0.21 ±

0.06 (ACh and nAChR-α4). For RACer(T17N) under the same conditions the migration indices

were 0.06 ± 0.01, 0.04 ± 0.02 and 0.04 ± 0.03, respectively. The differences between RACer and

RACer(T17N) were significant for blue light/LOVS1K/Orai1 (P=0.003) and VEGF-A/VEGFR2

(P=0.002) but not for ACh/nAChR-α4 (P=0.062).

91

Figure 6.9: Multi-signal migration of RACer cells using transwell assays.

Migration index of cells using the transwell assays for the indicated conditions. For all three cases, the Ca2+-mobilizing module was transfected in all cells (A, LOVS1K/Orai1; B, VEGFR2; C, nAChR-α4). The data is the mean ± standard deviation, n=3 independent experiments.

In the RACer transwell assays the results of the control experiments were largely consistent with

the results of the wound assay control conditions. As with the wound assay, blebbistatin

treatment significantly reduced the migration index compared to no inhibitor (P=0.004 and 0.005

for LOVS1K/Orai1 and VEGFR2, respectively). Unlike the wound assays, Y-27632 treatment

significantly reduced the migration index compared to no inhibitor (P=0.005 for both

LOVS1K/Orai1 and VEGFR2). The pattern of migration indices for transwell assays was

similar between RACer and CaRQ for LOVS1K/Orai1 and VEGFR2. While the magnitude of

the migration index was somewhat less for RACer than CaRQ the differences between

stimulated cells co-expressing RACer/LOVS1K/Orai1 or RACer/VEGFR2 and the control

conditions were all significant. However for cells co-expressing RACer/nAChR-α4, the

migration index for the stimulated condition was not significantly different than the control

conditions using RACer(T17N), pLyn-YFP, Y-27632 or blebbistatin treatment, suggesting that

nAChR-α4 is not a suitable Ca2+-mobilizing domain for regulating migration of RACer.

92

6.2.5 Design and Characterization of a Ca2+-sensitive Cdc42

The design of RACer was applied to the related Rho GTPase Cdc42 to create a Ca2+-sensitive

CaM-Cdc42 chimera (Figure 6.10 and Movies 6.5 and 6.6). Cdc42 plays a role in cell migration

by directing filopodia growth at the very leading edge of cells, therefore it may also be a useful

Rho GTPase to control toward the goal of re-programming cells with synthetic migration

regulation pathways. Cdc42, and its GEF’s, have been engineered for exogenous control

previously (6, 131-132). Given the structural similarity between Cdc42 and Rac1, the same

location as RACer, between amino acids 47 and 48, was chosen for embedding the IQp peptide.

Since Rac1 and Cdc42 are regulated by many of the same GEF’s and GAP’s, the E91H/N92H

mutations were also introduced to prevent the chimera from sinking cellular GAP’s.

The CaM-Cdc42 chimera was overexpressed in HEK293 cells with or without the presence of

the PBD-mRFP-GST probe. In cells without the PBD probe, the morphology was similar to wild

type HEK293 cells, and similar to RACer at basal Ca2+. There was some colocalization with the

PBD probe at basal Ca2+ (PC colocalization of 0.722 in a representative cell shown in the figure),

however this was also the case with RACer. There was no visible colocalization with the PBD

probe for the dominant negative mutant CaM-Cdc42(T17N) chimera, and the PC colocalization

was less, with a value of 0.458 in a representative cell.

When cells co-expressing the CaM-Cdc42 chimera and the PBD probe were stimulated with

ionomycin, there was an increase in colocalization between the probe and the PM-localized

chimera. In the cells shown, PC colocalization increased approximately 20% from 0.722 to

0.872, while for the CaM-Cdc42(T17N) chimera the PC colocalization increased 3.5% from

0.458 to 0.474. In cells without the PBD probe, ionomycin stimulation resulted in the

appearance of long, thing filopodia within 5-10 minutes after stimulation. In many cells this was

followed by the formation of sheet-like lamellipodia within 1-2 minutes from the onset of

filopodia. There was no apparent pattern to the formation of filopodia. In the cell shown there

are outgrowths on opposite ends of the cell. Overall, 78.5% ± 14.4% of cells expressing the

CaM-Cdc42 chimera stimulated with ionomycin developed filopodia and/or lamellipodia within

20 minutes after ionomycin stimulation. For cells expressing the CaM-Cdc42(T17N) chimera, or

the CaM-Cdc42 chimera pre-incubated with CDZ, ionomycin stimulation resulted in

morphology changes in 4.8% ± 4.7% and 5.6% ± 5.6% of cells, respectively. The likelihood of

93

morphology change were significantly different than without inhibitor and the Q61L mutant

(P=0.003 and 0.004, respectively). Pre-incubation with Y-27632 had no significant effect on the

likelihood of morphology change after ionomycin stimulation, 72.2% ± 2.8% of cells, compared

to the case without the inhibitor (P=0.613). Similarly, pre-incubation with blebbistatin had no

significant effect on the likelihood of morphology change after ionomycin stimulation, 55.6% ±

11.1% of cells, compared to no inhibitor (P=0.199).

Figure 6.10: Characterization of CaM-Cdc42 chimera.

A. Cells co-transfected with the CaM-Cdc42 chimera or its T17N mutant (YFP) and the PBD probe (RFP) before and after stimulation with ionomycin and the corresponding PCC values. B. Percentage of cells with filopodia or lamellipodia growth after stimulation with ionomycin for the indicated conditions (data is mean ± standard deviation, n=3 experiments with at least 10 cells studied per condition). C. A representative cell expressing the CaM-Cdc42 chimera stimulated with ionomycin at 5 minutes. White arrows indicate new filopodia/lamellipodia. Scale bars are 20 µm in A and 10 µm in C.

94

6.3 Discussion

RACer was activated by four different Ca2+-mobilizing stimuli representing a variety of Ca2+-

mobilizing modules. This shows that modular Ca2+ signaling leading to activation of engineered

proteins is not a phenomenon specific to RhoA chimeras such as CaM-RhoA(DP) or CaRQ, but

can be generalized to other engineered Ca2+-sensitive proteins. While the structural differences

between RhoA and Rac1 are small, modular activation of at least one other engineered Ca2+-

sensitive protein has been shown using ionomycin, LOVS1K and ChR2 (12, 124). Other aspects

of Ca2+ signaling such as spatial patterning, temporal responsiveness and frequency decoding

may also be applicable beyond the set of proteins developed here. The development of CaM-

RhoA(DP), CaRQ and RACer show that Ca2+ signaling, remembering its limitations, may be a

viable tool for other cell re-programming applications.

When RACer was activated by different Ca2+-mobilizing modules there was a qualitative

relationship between the duration of the Ca2+ signal and the duration of the cellular area increase:

shorter Ca2+ signals (those induced by ATP and ACh/nAChR-α4) resulted in shorter durations of

area increase and longer durations for longer Ca2+ signals (those induced by blue light/LOVS1K

or ionomycin). For CaRQ, the relationship between duration of Ca2+ signal and duration of

blebbing was less clear, but certainly in some cases the duration of blebbing lasted many (often

10-15) minutes longer than the Ca2+ signal. These differences can be explained by examining

how RhoA and Rac1 activate downstream proteins to result in cellular morphology changes.

• RhoA: Activation (natively by GTP loading, and here by Ca2+ loading) allows RhoA to bind

ROCK’s autoregulatory domain releasing autoinhibition and allowing ROCK to

phosphorylate myosin regulatory light chain on Ser19 (182) leading to ATPase activity and

cross-linking. The mechanism that translates phosphorylation into activity is not fully

known. One model suggests that phosphorylation releases a lever arm in the central helix of

myosin light chain (183-184) and this frees the two myosin heavy chain heads from blocking

each other (185). Once myosin regulatory light chain is phosphorylated, it maintains the

ability to activate myosin ATPase until it is de-phosphorylated by myosin light chain

phosphatase.

• Rac1: Activation allows Rac1 to interact with the WAVE regulatory complex (WRC). In a

recent study reporting the structure of WRC (58) Rac1 binding was proposed to release the

95

VCA region of WRC by either binding an associated protein called Sra1, which in the

absence of Rac1 binds the VCA, or by stabilizing WAVE so that the Sra1-VCA interaction is

abolished. Once the VCA region is released it is free to interact with Arp2/3 and other actin-

polymerizing factors. When Rac1 dissociates from WRC the VCA region is presumably re-

inhibited until such time as some other signal once again releases it.

• The main difference between the two cases is the requirement for Rho activation leading to

morphology changes. Once RhoA-ROCK phosphorylates its target to increase actomyosin

cross-linking and cytoskeletal tension, the effect of phosphorylation will continue after

RhoA-ROCK is de-activated, until myosin regulatory light chain is de-phosphorylated. For

Rac1-WRC, once Rac1 is de-activated, the VCA region of WAVE loses the ability to guide

actin polymerization. In this way, RhoA-ROCK may have some effect after it is turned off

whereas Rac1-WRC must be continuously activated to see a morphological response.

Changes in whole cell area were used as a way to quantify lamellipodia formation as a surrogate

for Rac1 activity. This method has been used previously (5) and was chosen here because it is

difficult to develop objective criteria for counting lamellipodia given there irregular size and

shape. The area analysis was not intended to be used as a way to distinguish between

lamellipodia and blebbing; applying the same area thresholding analysis to blebbing cells also

shows an area increase on morphology change (Appendix I shows area thresholding analysis

applied to the cells shown in Figure 5.7D). A close inspection shows that the fold area change

signal oscillates in synchrony with blebs as they form and retract. However, the method of

identifying blebs used in Chapter 5 was still the most appropriate because specific criteria can be

developed for identifying blebs by visual inspection.

The ability of RACer to control migration was assayed using the same methods as was done for

CaRQ: wound assays and transwell migration assays. Broadly speaking, the results were similar

for the two proteins, with two exceptions: the effect of inhibitors such as Y-27632 and

blebbistatin on RACer-expressing cells depended on the particular assay (whereas for CaRQ

most effects were consistent across assays), and the nAChR-α4 Ca2+-mobilizing module did not

effectively allow RACer-expressing cells to migrate in transwell assays.

For RACer, blebbistatin treatment did not have a significant effect on lamellipodia formation but

did inhibit migration in both assays. Blebbistatin is an inhibitor of myosin ATPase and

96

blebbistatin treatment prevents actin-myosin crosslinking from generating contractile forces

during cell migration (179-180). Lamellipodia formation is an actin polymerization-dependent

process utilizing the WAVE and Arp2/3 complexes. For a cell to migrate in the mesenchymal

mode both lamellipodia formation and actin-myosin contractility are required. Blebbistatin

therefore inhibits a process necessary for migration that is unnecessary for lamellipodia

formation. Blue light has been noted to inhibit blebbistatin, but at light intensities on the order of

1 mW/µm2 (186). Treatment with Y-27632, a ROCK inhibitor, did not inhibit lamellipodia

formation but did inhibit migration in the transwell assay. For cells to transit through the porous

filter they must “squeeze” through 8.0 µm diameter pores, and ROCK may be responsible for

generating this contraction force. The blebbistatin experiment suggests that actin-myosin

contractility is also important for migration in the wound assay, and whether Y-27632 also had a

significant effect on wound assays is unclear because the P value for that test was very close to

the pre-set α value of 0.05. Actomyosin contractility can be activated by other proteins such as

those that phosphorylate myosin light chain phosphatase (187), which may explain why ROCK

inhibition had only a partial effect on this mode of cell migration in the wound assay.

There was not a statistically significant effect of ACh/nAChR-α4-stimulation on RACer-

expressing cells in transwell migration assays. This may have represented a technical limitation

to the experiment whereby the small size of the ACh molecule allowed it rapidly diffuse through

the porous transwell filter abolishing the chemical gradient, however this effect was not noted in

similar transwell experiments with CaRQ-expressing cells. ACh (and ATP) stimulation of cells

co-expressing RACer and nAChR-α4 resulted in morphology change of short duration (less than

1 minute) and small magnitude (approximately 1.1 fold area change). The morphology change

induced by ACh/nAChR-α4-stimulation of RACer-expressing cells was likely insufficient to

affect the migration of those cells in a meaningful way. This suggests that not all Ca2+-

mobilizing domains will be appropriate for all engineering Ca2+-sensitive proteins; indeed this

could have been predicted by considering how natural proteins respond to different Ca2+ signals.

Increasing the amplitude or duration of the Ca2+ transient (for example using a Ca2+ buffer, with

repeated stimulation, or localization of RACer nearer the nAChR-α4 channel) may rescue the

ability of ACh/nAChR-α4 to induce migration in these cells.

A Ca2+-sensitive Cdc42 protein was created by applying the RACer design, and broadly speaking

the embedded IQp design, to Cdc42. Activation of CaRQ and RACer resulted in morphology

97

changes that closely resembled RhoA and Rac1 while activation of the CaM-Cdc42 chimera

yielded morphology changes reminiscent of Cdc42 and Rac1. This was likely because Cdc42

activation is known to activate Rac1, for example through Cdc42-PAK binding leading to α-PIX

phosphorylation and activation, α-PIX being a GEF for Rac1 (68). The high degree of structural

similarity between Rho protein family members suggested that embedding IQp at the same site

would confer a similar Ca2+-responsive characteristic between the chimeras. Applying that

argument and the design to other Rho family members such as RhoC, RhoE and RhoG, or Rac2

and Rac3, may expand on the functionality offered by the existing chimeras. There is also a

broad structural and functional similarity within the Ras superfamily. For example the dominant

positive mutants Q61L/Q63L in the Rho family are analogous to the Q63E or Q79L dominant

positive mutants in Rap1 and Rab5, respectively (188-189). Therefore, the structural similarity

within the entire Ras superfamily may mean that the Ca2+-control design can be applied to

control proteins regulating diverse processes such as vesicle trafficking (Rab and Arf),

nucleocytoplasmic transport (Ran) and cell adhesion (Rap).

The development of CaRQ and RACer is an important contribution to cell re-programming, as

defined in this work, because they show that control over cell migration can be rewired in a way

that is flexible and tunable.

• Control over cell migration could be applicable to many re-programming applications such as

guiding cells to populate a biomaterial scaffold or microfluidic device, or targeting cells to

sites of injury or disease in vivo. Other strategies have developed light-based control of cell

migration previously (5-6). CaRQ and RACer compliment those devices already reported in

the literature in several ways. One way is the tunable nature of Ca2+ control over CaRQ and

RACer: in the most extreme case, the direction of the morphology switch was reversed by

reversing the embedded and fused CBP’s, however this idea could be expanded by using

different CBP’s to alter the kinetics and Ca2+-sensitivity of morphology control.

• The ability to control cell migration with multiple signal inputs is a new feature not offered

by existing cell migration control strategies. Previous systems designed using specific

caging mechanisms (5) and protein-protein interactions (6) rely on specific wavelengths of

light to activate Rho GTPases. The strategy presented in this work shows how two modules

can be combined to achieve control by light or chemical signals. The use of exogenous

98

chemicals to control cell migration may make CaRQ and RACer more suitable for

controlling cell migration in vivo in large organisms with poor light penetration.

• In the migration assays used here, directional cues were provided by the physical

arrangement of the apparatuses. For wound assays, the open space of the wound provided a

directional cue by default. In transwell assays there was likely a combined contribution from

the chemical and light cues and the thinness of the porous filter. Demonstrating that CaRQ

and RACer can mediate directed migration toward a point source will be a goal for future

experiments (discussed below in section 8.1).


In this chapter a Ca2+-sensitive Rac1 chimera was created by applying the embedded IQp

insertion strategy of CaRQ to Rac1 to create RACer. Increasing the intracellular Ca2+ in cells

expressing RACer resulted in the formation of new lamellipodia indicative of Rac1 activation.

As with CaRQ, a variety Ca2+-mobilizing modules were used to increase intracellular Ca2+ using

different extracellular stimuli; long-term activation with these stimuli increased the migratory

potential of RACer-expressing cells.

1. A Ca2+-sensitive Rac1 chimera was created by embedding IQp at the same sites as were

used for CaRQ (between amino acids 47 and 48), with an amino terminal fusion of a

constitutive membrane-localization tag, YFP, CaM and MLCKp. Mutations were

introduced (E91H, N92H) to reduce GAP sinking.

2. Stimulation of RACer with ionomycin resulted in lamellipodia formation and cell area

increase. In cells co-expressing RACer and PBD-mRFP-GST, ionomycin increased the

PC colocalization between RACer and the PBD probe. The overall Ca2+ EC50 for

RACer-induced lamellipodia was 24 µM. RACer activation also resulted in morphology

changes in several non-epithelial cell lines including COS7 and RAW264.7.

3. Three classes of Ca2+-mobilizing modules (endogenous, exogenous and synthetic) were

used to activate RACer. Certain modules (ATP and ACh) resulted in area increases of a

small magnitude while LOVS1K/Orai1 caused large area increases. There was a

qualitatively proportional relationship between the duration of Ca2+ signal and the

duration of morphology change.

99

4. Repeated illumination of cells co-expressing RACer and LOVS1K/Orai1 showed

increase migration into wounds in wound scratch assays. Cells co-expressing RACer and

LOVS1K/Orai1 or VEGFR2 had increased migration indices when stimulated with light

or VEGF-A, relative to negative controls. Cells co-expressing RACer and nAChR-α4 did

not show significant changes in migration index when exposed to ACh.

5. The design of RACer was applied to Cdc42 to create a CaM-Cdc42 chimera that was

Ca2+ sensitive. Ionomycin stimulation of cells expressing the CaM-Cdc42 chimera

resulted in filopodia and lamellipodia formation.

In relation to the research objectives outlined in Chapter 1, the work in this chapter addresses

each of the three objectives. 1. Ca2+-sensitive Rac1 (RACer) and Cdc42 were created using the

chimeric design of CaRQ. Activation of RACer resulted in Rac1-distincitve morphologies and

similarly for the CaM-Cdc42 chimera, showing that the design is generalizable between Rho

proteins while maintaining protein-specific behaviours. 2. Control over lamellipodia formation

and cell area increase was achieved by several of the same Ca2+-mobilizing modules used with

CaRQ including ATP (endogenous), ACh (exogenous) and LOVS1K (synthetic). 3. Prolonged

activation of RACer increased wound closure in a wound scratch assay and migration indices in

a transwell migration assay. Not all Ca2+-mobilizing modules had equivalent effects on RACer-

mediated cell migration.

100

7 Summary and Conclusion

Ca2+-sensitive chimeras of RhoA, Rac1 and Cdc42 were created to control cell morphology and

migration (Figure 7.1). The results of this work were presented in three chapters (Chapters 4-6)

based on different strategies for engineering control over Rho proteins.

• Chapter 4: A CaM binding site was identified in RhoA, RhoACBP, and this knowledge was

used to create a tandem fusion of RhoA and CaM. The tandem fusion, CaM-RhoA(DP)-

YFP, mediated Ca2+-dependent bleb retraction in HEK293, HeLa and CHO cells in

responsive to different Ca2+-elevating chemicals such as ionomycin, UTP and ATP,

respectively. Bleb retraction was also achieved in response to ACh and blue light when cells

were co-expressing nAChR-α4 or ChR2, respectively.

• Chapter 5: The IQp CBP was embedded within the secondary structure of RhoA, and this

domain was fused with CaM-MLCKp to create a Ca2+-dependent chimera, CaRQ, that

regulated bleb formation. Changing the embedded CBP resulted in a chimera with different

behaviour. CaRQ was activated by different Ca2+-mobilizing domains including AChR-α4,

VEGFR2, LOVS1K/Orai1 and SOCE; however the spatial localization of CaRQ was

important suggesting that CaRQ has sensitivity to different Ca2+ signal patterns. Prolonged

activation of CaRQ by over 24 hours resulted in increased cell migration in both wound

scratch assays (light by LOVS1K) and transwell migration assays (light, ACh and VEGF-A).

• Chapter 6: The design of CaRQ was applied to Rac1 to create a Ca2+-sensitive Rac1

chimera, RACer. RACer mediated lamellipodia formation in response to Ca2+, and the

modularity of Ca2+ signaling with RACer was demonstrated with ACh/nAChR-α4 and blue

light/LOVS1K. There was a different relationship between the duration of Ca2+ signal and

morphology than was noticed for CaRQ, and these differences were likely due to the

mechanism of RhoA and Rac1 interaction with their effector proteins. As with CaRQ,

prolonged activation of RACer over 24 hours increased cell migration in wound and

transwell assays. The RACer design was also applied to Cdc42.

Three main objectives were set at the outset of this work. The objectives delineated specific,

logical steps toward the overall goal of engineering multiple-input control over cell migration.

• Objective 1: Several chimeras were created where Rho-associated morphologies were

switched by intracellular Ca2+. Chimeras were created for each of the three principal Rho

101

proteins, RhoA (CaM-RhoA(DP) and CaRQ), Rac1 (RACer) and Cdc42 (CaM-Cdc42

chimera), that controlled three morphological structures, blebs, lamellipodia and filopodia.

• Objective 2: Artificial Ca2+ signaling networks enabled control over cell morphology by a

variety of exogenous stimuli. Networks were created by combining a variety of Ca2+-

mobilizing domains (for example, nAChR-α4, ChR2, LOVS1K/Orai1, VEGFR2 and SOCE)

with the Ca2+-sensitive Rho chimeras list above. This resulted in control over cell

morphology by a variety of exogenous stimuli including ACh, blue light and VEGF-A. Not

all combinations behaved equally under the same conditions. Properties of Ca2+-signaling

such as spatial patterning, buffering and temporal responsiveness, were applied to improve or

tune the morphological response to different stimuli.

• Objective 3: Prolonged activation of artificial Ca2+ signaling networks resulted in increased

cell migration for the two chimeras tested, CaRQ and RACer. In both cases cells co-

expressing LOVS1K/Orai1 and illuminated over 24 hours migrated significantly in wound

scratch assays. In transwell migration assays stimulation of CaRQ (with blue light/LOVS1K,

VEGF-A/VEGFR2 and ACh/nAChR-α4) and RACer (with blue light/LOVS1K and VEGF-

A/VEGFR2) resulted in significant migration through porous filters.

Figure 7.1: Overview of selected artificial Ca2+ signaling networks developed here.

102

8 Future Directions

This chapter will describe several ways that the technology developed in the previous chapters

can be used in more complex applications, improved upon, and applied toward answering

biological questions in innovate ways. First, the use of CaRQ and RACer in more complex re-

programming applications will be discussed including potential challenges that will be

encountered. Second, further application of the principles of Ca2+ signaling to refine artificial

Ca2+ signaling networks will be considered. Third, ways to use the technology and insights

gained in this work to answer biological questions about Rho proteins will be discussed.

8.1 Further Development of Cell Re-programming

Further application of CaRQ and RACer as useful modules in re-programming cells will require

conclusive demonstrations of their ability to mediate migration towards a point or highly

localized source of chemical factors in increasingly complex environments. As discussed in

Chapters 5 and 6, the assays used here were able to assess the migratory potential of cells, that is,

their ability to engage in a random walk, with directional cues being given by the physical

arrangement of objects in the assays. A more localized source of chemical signal (for the

purpose of this discussion, assume the stimulus is VEGF-A) could be achieved using VEGF-A

coated beads (based on, for example, agarose, sepharose or a magnetic material for improved

handling), hydrogels with tunable rates of VEGF-A diffusion, or even VEGF-A expressing

tumour explants (190-191). In such an experiment re-programmed cells could be seeded nearby

in a continuous media and, using migration rate as a readout, the ability of artificial Ca2+

signaling networks to mediate migration toward the source could be verified.

Further experiments controlling the direction, duration and frequency of illumination may also

be used to investigate the behaviour of re-programmed cells. For example, the effect of

illumination using different colours of light, duty cycles, and specific animation routines (e.g. a

moving spot to guide cells) could be investigated with the iPod. Similarly, a neutral density filter

with a left-to-right intensity gradient could be used to determine whether a “light intensity

gradient” could control the direct of migration. A microcontroller, for example, the Arduino

Uno, may also be useful for further exploring the controllability of re-programmed cell migration

in vitro. The Arduino platform allows for simple “Sketches” to be written in a procedure-

oriented language and downloaded via a USB port to an integrated circuit board based on an

103

Atmega168 chip, which can then drive an analog or digital circuit. Pursuing an Arduino-based

project for manipulating re-programmed cells would have several key goals:

1. Develop, and code, dynamic light patterns: Dynamic light patterns (e.g. left-to-right

waves, concentric shrinking circles, moving spots, etc) can be coded in the Arduino

language using the core loop() and the delay(t) functions, where t is a number of

milliseconds. The complexity of the patterns will be limited because there are only 13

digital output pins available.

2. Create a useful array of LED’s: Creating LED circuits is trivial, however LED’s will

need to be small enough, and closely packed, to achieve good spatial resolution of light,

remembering that the area of a dish is ~10 cm2. The LED array will need to be wired so

that it can execute the various dynamic light patterns coded above.

3. Integrate the LED circuit, Arduino board and code: Functional integration of the three

components of the system should be straightforward, since there is much supporting

documentation online (www.arduino.cc).

4. Test the behaviour of re-programmed cells in response to dynamic light patterns:

Logistical challenges may occur during experimentation, such as whether the circuit

components can function in the incubator (high heat and humidity), and the physical

arrangement of the circuit board, LED array and cell dish. Eventually, more complex

experiments may be performed, for example, observing cells in the simultaneous

presence of a left-to-right light wave and a right-to-left growth factor gradient.

Notwithstanding the above in vitro experiments, the primary long-term goal of this work is to re-

program cells for in vivo applications; there are many challenges related to testing CaRQ and

RACer in in vivo models, three of which are presented below:

• Cell source and handling: The proper selection of a host cell population will be critical, and

will likely depend on the re-programming application. For re-programming to clear cells

within a tumour bulk, an immunological cell such as a T-cell or macrophage may be the most

appropriate; in one study, T-cells were removed from a host by apheresis, sorted, and

chimeric antigen receptors introduced by viral infection (2). iPS or direct transfer technology

may be alternatives since hematopoietic progenitor cells have been created in this manner

(192). However this field is still in its infancy, and the immunogenic potential of iPS cells in

104

vivo is unclear (193). The potential for in vivo re-programming should also be considered

(1). The transient transfection methods of this study will be replaced with stable, probably

viral-mediated gene transfer and cell sorting to ensure a homogenous population before cells

are returned to the host. Whether expansion of the population should be attempted ex vivo

should also be considered. Protocols will need to be developed to ensure an efficient and

safe handling of cells during ex vivo re-programming.

• Trafficking in vivo: Once implanted, cells will need to migrate toward the site of disease or

injury, or some other relevant site (for example the bone marrow for engineered T cells).

According to the thinking presented here, this will be mediated by an appropriate Ca2+-

mobilizing module which will be chosen based on the particular application. Migration

could be facilitated by appropriate choice of injection or implantation site. Strategies to

minimize host-mediated engineered cell clearance (194) and surmounting physiological

barriers (e.g. the blood brain barrier) (195-196) will be needed depending on the application.

• Available signals and biological noise: The complexity of the biological milieu in vivo may

present a substantial challenge. In large organisms light is of limited use and chemical cues

will be most appropriate. Migration cues should be chosen to avoid chemical signals present

in normal physiological operation that are not specific to disease or injury. The dynamic

range of a particular cue which cells will be able to sense may need to be tuned. The

chemical and genetic heterogeneity of a target should also be considered (197).

The engineered systems presented here to control cell migration will likely be only one of

several components in eventual re-programming applications. Other systems may be developed

such as triggered protein secretion, mechanisms for more sophisticated target cell recognition (2)

and gene transfer or membrane fusion with target cells (198-199). While designing these

systems is outside the scope of the current work, one could imagine how these functionalities

could be developed by borrowing and recombining necessary parts from other biological entities

and systems. For example, triggered protein secretion could be achieved by conditional gene

expression, or by manipulating GTPases that regulate exocytosis (200). Gene transfer via

membrane fusion could be achieved by expression of the VSV-G protein from vesicular

stomatitis virus: when the extracellular domain of VSV-G encounters an acidic

microenvironment, such as that found in the tumour bulk, conformational changes in the protein

allow it to bind glycoproteins and initiate membrane fusion (198). To ensure compatibility with

105

the cell migration control pathways developed here, the mechanism regulating other functional

modules must not overlap with the intermediate Ca2+ signal, unless the effects are desired to

occur at the same time. However, as in nature, and with a more sophisticated understanding of

Ca2+ signal decoding, multiple processes may be able to make use of the Ca2+ intermediate

provided that they are designed to interpret distinct spatiotemporal Ca2+ patterns.

8.2 Refining Artificial Ca2+ Signaling Networks

Modularity and spatial localization to subcellular compartments were the primary principles of

natural Ca2+ signaling networks that were used in this work to develop artificial Ca2+ signaling

networks. As these artificial networks are used in cells for progressively complex tasks the

sophistication of the artificial networks may also need to increase. Applying other principles of

natural Ca2+ signaling networks may allow the artificial networks developed here to have

increased functionalities and rescue module combinations that did not perform well initially,

such as ACh/nAChR-α4 and RACer in the transwell migration assays. Four principles in

particular will be considered: Ca2+ buffering, precise localization, chromatic multiplexing and

waveform decoding.

Ca2+ buffering: Ca2+ buffering proteins such as parvalbumin, calbindin and calsequestrin

sequester Ca2+ with differing capacities and affinities to augment physiological processes.

Calbindin for example shapes Ca2+ concentration microdomains at the cytoplasmic mouth of

Ca2+ channels during action potentials (201) while calsequestrin provides a ready source of Ca2+

for rapid muscle contraction (202). Buffering proteins may be useful if localized near a Ca2+-

mobilizing domain to both increase the local Ca2+ concentration for a downstream Ca2+-sensitive

protein and to restrict the area of the Ca2+ signal to minimize the Ca2+ signal seen by the rest of

the cell. Mutations within the Ca2+-binding EF-hands have been shown to affect the affinity and

kinetics of Ca2+ sequestration by these buffers, and this may also represent a way to tuning the

overall function the artificial Ca2+ network (202-203).

Precise localization: A more precise localization of Ca2+-sensitive proteins may improve their

ability to respond to local Ca2+ signals. While this strategy was applied in this work, localization

was considered at the compartment level. In experiments with Orai1 and Stim1-CaRQ, protein

complex-level localization was implicated in controlling the response to CaRQ to Ca2+ influx.

Indeed, Ca2+ concentration microdomains are often submicrometer in size (13, 108, 201) and

106

more precise localization of Ca2+-sensitive proteins may improve response while limiting the

effect of long-range or global Ca2+ signals on the engineered proteins. Precise channel-level

localization could be achieved by genetic fusion of the Ca2+-sensitive protein with a cytoplasmic

surface of a channel, either tandemly or embedded, as with fluorescent fusions of nAChR-α4 (9).

Fusion between Ca2+-sensitive proteins and other, natively channel-associated proteins may also

be sufficient to localize engineered Ca2+-sensitive proteins within the Ca2+ microdomain.

Chromatic multiplexing: Light-based control of biological systems has received a lot of

attention in the literature (5-6, 12, 58), and in this work because of its easy of use and precise

delivery. Blue light-sensitive proteins (LOV2 and ChR2) were used exclusively here to mobilize

Ca2+, but red light-gated VChR1 is a cation channel (27) that may also be a useful as a Ca2+-

mobilizing domain. The phycobilin family of chromophores have been used to create

red/infrared (6) and green/red (204) protein photoswitches which may be applied to Ca2+

signaling in the future. Using several wavelengths of light separately or simultaneously, or

chromatic multiplexing, will allow an expanded set of functionalities. To prevent Ca2+-sensitive

proteins being confused by other wavelengths of light which also generate the Ca2+ intermediate,

a mixed population of genetically distinct cells may be used, or genetic fusion of the Ca2+-

sensitive protein at the channel mouth may provide sufficiently distinct Ca2+ microdomains to

prevent confusion.

Waveform decoding: Native Ca2+-sensitive proteins are capable of responding to Ca2+

waveforms of specific shapes, amplitudes and frequencies while ignoring, or being less affected,

by others (8, 97, 107). This principle could be applied to artificial Ca2+ signaling networks to

enable two or more Ca2+-sensitive proteins to operate in the same cell, or perhaps for one Ca2+-

sensitive protein to have two or more effects. To do this, other native Ca2+-sensitive protein

domains may need to be considered, for example TnC (structurally similar to CaM but with its

own target peptides such as TnI), calcineurin (a Ca2+-dependent phosphatase) and PKC or

CaMKII (Ca2+-dependent kinases). For example, varying the frequency of Ca2+ oscillations

altered the amount of kinase activity in CaMKII (107). Since the frequency of Ca2+ oscillation

can be controlled with ChR2 and LOVS1K, this could allow CaMKII, and some activity

engineered downstream of it, to be controlled by the frequency of light delivery.

107

8.3 Studying Rho Proteins and Cell Migration

The technology developed and insights gained in this work can be used to answer questions

about Rho proteins and cell migration. Several outstanding questions regarding the role of Rho

proteins in cell migration will benefit from technologies that enable spatiotemporal activation,

transferability between Rho proteins and establishment of different cell morphologies. While

some experiments may be clouded by the use of the Ca2+ intermediate, some of these capabilities

are not currently possible with other methods. Other intermediate signals may also be possible,

as will be further discussed below. Several experimental approaches using the insights

developed in this work will be described below, toward answering three open questions:

1. What is the contribution of migration mode toward overall cancer cell invasiveness?

2. What is the role of lesser-studied Rho proteins, such as RhoC, RhoG and Rac2 in migration?

3. How do Rho proteins function in collective versus single cell migration?

Cancer cells are one example where cells are known to migrate with amoeboid-like and

mesenchymal modes (37, 48, 64, 69, 71, 74, 76, 82, 89, 157) and either collectively or singly

(156-158, 169-171). The invasive properties of cancer cells, such as migration speed and matrix

degradation have been shown to depend on DOCK1, a RhoGEF, by siRNA and mutational

studies (74, 86). CaRQ and RACer provide the unique capability of reversibly inducing either

migration mode for a controllable duration, and could be used to study the relationship between

migration mode and invasion. Since light can be applied in many assays to activate CaRQ and

RACer, the dependence of migration mode on substrate stiffness, substrate adhesion, chemical

milieu, MMP activity and the cellular genetic background can be tested using primary cell or

established model cell lines in a variety of in vitro assays. In terms of collective cell migration,

little is known about how specific Rho activities contribute to cells’ decisions to migration alone

or in groups. Rac1 is known to regulate cell-cell adhesion (205), and RACer localized to cell

junctions may be used provide new insights about Rac1’s role in coherent collective migration.

RhoE, a RhoA antagonist-like GTPase, has been shown to locally down-regulate ROCK activity

at cell-cell junctions (157), and localized CaRQ may also provide insight into the local and/or

global ROCK dependence of collective cell migration.

108

The above strategy could be applied to separate populations of cells expressing either CaRQ or

RACer for comparisons to be drawn between the two cases. However, many invasive cells have

been shown to exhibit both modes to some extent simultaneously. CaRQ and RACer could be

activated in the same cell using one of the multiplexing/decoding strategies outline above (see

section 8.2). For example in a cell with VChR1-localized CaRQ and LOVS1K/Orai1-localized

RACer, the relative duration or frequency of red and blue light illumination would lead to

differential activation of each migration mode, and the effects of this could be studied on the

same parameters described earlier. If Ca2+ signal decoding was insufficient to separate the two

light signals, a red light-activated CaRQ could be combined with blue light-activated PARac (5),

or blue light-activated CaRQ with red light-translocated Rac1/Cdc42 GEF’s (6).

The structural similarity between Rho proteins may allow for the embedded peptide strategy to

be applied beyond RhoA, Rac1 and Cdc42. Many of the other some 20 Rho proteins including

RhoC, RhoG, Rac2 and Rac3 (79, 206-208) have not been as well studied. The embedded

peptide strategy developed here could lead to a suite of real-time inducible Rho proteins which

would be studied independently, or using a multiplexing/decoding strategy, in combination with

other Rho proteins. While the Ca2+ intermediate presents a non-ideality in this situation, there is

currently no other strategy to achieve real-time activation of Rho proteins without introducing

additional point mutations, as was required when generalizing PARac to PACdc42 (5).

The identification of the insertion site in the structure of Rho may in itself be used to develop

inducible Rho proteins without relying on the Ca2+ intermediate. For example, one could

imagine embedding a modified PDZ-domain interacting peptide between amino acids 47/48

(49/50 for RhoA) such that PKA activation by forskolin treatment disrupts the interaction (131).

This could be analogous to CaM dissociating from IQp on Ca2+ loading. A second option may

be to use the embedded iFKBP strategy that has been applied to stabilize Src kinases on

rapamycin addition and FRB recruitment (209). In this case, rapamycin could stabilize the

insertion site allowing Rho protein activity to be restored. While both of these methods have the

drawback of being irreversible, they could allow real-time activation with potential applicability

across the entire Rho family without using the Ca2+ intermediate. These new tools could later be

combined with pre-existing ones (such as CaRQ/RACer or photo-activatable proteins developed

by others) to develop a suite of inducible Rho proteins responsive to different exogenous stimuli

that could be activated within the same cell without the need for signal decoding.

109

9 References 1. Breitbach CJ, et al. (2011) Intravenous delivery of a multi-mechanistic cancer-targeted

oncolytic poxvirus in humans. Nature 477(7362):99-102. 2. Kalos M, et al. (2011) T cells with chimeric antigen receptors have potent antitumor

effects and can establish memory in patients with advanced Leukemia. Sci Transl Med 3(95):95ra73.

3. Perez EE, et al. (2008) Establishment of HIV-1 resistance in CD4+ T cells by genome editing using zinc-finger nucleases. Nat Biotechnol 26(7):808-816.

4. Shimizu S, et al. (2010) A highly efficient short hairpin RNA potently down-regulates CCR5 expression in systemic lymphoid organs in the hu-BLT mouse model. Blood 115(8):1534-1544.

5. Wu YI, et al. (2009) A genetically encoded photoactivatable Rac controls the motility of living cells. Nature 461(7260):104-108.

6. Levskaya A, Weiner OD, Lim WA, & Voigt CA (2009) Spatiotemporal control of cell signalling using a light-switchable protein interaction. Nature 461(7266):997-1001.

7. Wang X, He L, Wu YI, Hahn KM, & Montell DJ (2010) Light-mediated activation reveals a key role for Rac in collective guidance of cell movement in vivo. Nat Cell Biol 12(6):591-597.

8. Berridge MJ, Lipp P, & Bootman MD (2000) The versatility and universality of calcium signalling. Nat Rev Mol Cell Biol 1(1):11-21.

9. Nashmi R, et al. (2003) Assembly of alpha4beta2 nicotinic acetylcholine receptors assessed with functional fluorescently labeled subunits: effects of localization, trafficking, and nicotine-induced upregulation in clonal mammalian cells and in cultured midbrain neurons. J Neurosci 23(37):11554-11567.

10. Dawson NS, Zawieja DC, Wu MH, & Granger HJ (2006) Signaling pathways mediating VEGF165-induced calcium transients and membrane depolarization in human endothelial cells. FASEB J 20(7):991-993.

11. Petreanu L, Huber D, Sobczyk A, & Svoboda K (2007) Channelrhodopsin-2-assisted circuit mapping of long-range callosal projections. Nat Neurosci 10(5):663-668.

12. Pham E, Mills E, & Truong K (2011) A synthetic photoactivated protein to generate local or global Ca(2+) signals. Chem Biol 18(7):880-890.

13. Dick IE, et al. (2008) A modular switch for spatial Ca2+ selectivity in the calmodulin regulation of CaV channels. Nature 451(7180):830-834.

14. Crivici A & Ikura M (1995) Molecular and structural basis of target recognition by calmodulin. Annu Rev Biophys Biomol Struct 24:85-116.

15. Miyawaki A, et al. (1997) Fluorescent indicators for Ca2+ based on green fluorescent proteins and calmodulin. Nature 388(6645):882-887.

16. Nakai J, Ohkura M, & Imoto K (2001) A high signal-to-noise Ca(2+) probe composed of a single green fluorescent protein. Nat Biotechnol 19(2):137-141.

17. Truong K, et al. (2001) FRET-based in vivo Ca2+ imaging by a new calmodulin-GFP fusion molecule. Nat Struct Biol 8(12):1069-1073.

18. Zhao Y, et al. (2011) An expanded palette of genetically encoded Ca(2) indicators. Science 333(6051):1888-1891.

110

19. Kolishetti N, et al. (2010) Engineering of self-assembled nanoparticle platform for precisely controlled combination drug therapy. Proc Natl Acad Sci U S A 107(42):17939-17944.

20. Shi J, Votruba AR, Farokhzad OC, & Langer R (2010) Nanotechnology in drug delivery and tissue engineering: from discovery to applications. Nano Lett 10(9):3223-3230.

21. Takahashi K & Yamanaka S (2006) Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126(4):663-676.

22. Ambasudhan R, et al. (2011) Direct reprogramming of adult human fibroblasts to functional neurons under defined conditions. Cell Stem Cell 9(2):113-118.

23. Fischer A, Hacein-Bey-Abina S, & Cavazzana-Calvo M (2010) 20 years of gene therapy for SCID. Nat Immunol 11(6):457-460.

24. Fenno L, Yizhar O, & Deisseroth K (2011) The development and application of optogenetics. Annu Rev Neurosci 34:389-412.

25. Strickland D, Moffat K, & Sosnick TR (2008) Light-activated DNA binding in a designed allosteric protein. Proc Natl Acad Sci U S A 105(31):10709-10714.

26. Strickland D, et al. (2010) Rationally improving LOV domain-based photoswitches. Nat Methods 7(8):623-626.

27. Zhang F, et al. (2008) Red-shifted optogenetic excitation: a tool for fast neural control derived from Volvox carteri. Nat Neurosci 11(6):631-633.

28. Airan RD, Thompson KR, Fenno LE, Bernstein H, & Deisseroth K (2009) Temporally precise in vivo control of intracellular signalling. Nature 458(7241):1025-1029.

29. Kleinlogel S, et al. (2011) Ultra light-sensitive and fast neuronal activation with the Ca(2)+-permeable channelrhodopsin CatCh. Nat Neurosci 14(4):513-518.

30. Knopfel T, et al. (2010) Toward the second generation of optogenetic tools. J Neurosci 30(45):14998-15004.

31. Harper SM, Christie JM, & Gardner KH (2004) Disruption of the LOV-Jalpha helix interaction activates phototropin kinase activity. Biochemistry 43(51):16184-16192.

32. Harper SM, Neil LC, & Gardner KH (2003) Structural basis of a phototropin light switch. Science 301(5639):1541-1544.

33. Kay CW, et al. (2003) Blue light perception in plants. Detection and characterization of a light-induced neutral flavin radical in a C450A mutant of phototropin. J Biol Chem 278(13):10973-10982.

34. Corchnoy SB, et al. (2003) Intramolecular proton transfers and structural changes during the photocycle of the LOV2 domain of phototropin 1. J Biol Chem 278(2):724-731.

35. Hewavitharana T, Deng X, Soboloff J, & Gill DL (2007) Role of STIM and Orai proteins in the store-operated calcium signaling pathway. Cell Calcium 42(2):173-182.

36. Nagel G, et al. (2003) Channelrhodopsin-2, a directly light-gated cation-selective membrane channel. Proc Natl Acad Sci U S A 100(24):13940-13945.

37. Hall A (1998) Rho GTPases and the actin cytoskeleton. Science 279(5350):509-514. 38. Jaffe AB & Hall A (2005) Rho GTPases: biochemistry and biology. Annu Rev Cell Dev

Biol 21:247-269. 39. Ridley AJ (2001) Rho GTPases and cell migration. J Cell Sci 114(Pt 15):2713-2722. 40. Hakoshima T, Shimizu T, & Maesaki R (2003) Structural basis of the Rho GTPase

signaling. J Biochem 134(3):327-331.

111

41. Tkachenko E, et al. (2011) Protein kinase A governs a RhoA-RhoGDI protrusion-retraction pacemaker in migrating cells. Nat Cell Biol 13(6):660-667.

42. Ding F, Yin Z, & Wang HR (2011) Ubiquitination in Rho signaling. Curr Top Med Chem 11(23):2879-2887.

43. Fang WB, Brantley-Sieders DM, Hwang Y, Ham AJ, & Chen J (2008) Identification and functional analysis of phosphorylated tyrosine residues within EphA2 receptor tyrosine kinase. J Biol Chem 283(23):16017-16026.

44. Blomquist A, et al. (2000) Identification and characterization of a novel Rho-specific guanine nucleotide exchange factor. Biochem J 352 Pt 2:319-325.

45. Taya S, et al. (2001) Direct interaction of insulin-like growth factor-1 receptor with leukemia-associated RhoGEF. J Cell Biol 155(5):809-820.

46. Garcia-Mata R, Boulter E, & Burridge K (2011) The 'invisible hand': regulation of RHO GTPases by RHOGDIs. Nat Rev Mol Cell Biol 12(8):493-504.

47. Tcherkezian J & Lamarche-Vane N (2007) Current knowledge of the large RhoGAP family of proteins. Biol Cell 99(2):67-86.

48. Worthylake RA, Lemoine S, Watson JM, & Burridge K (2001) RhoA is required for monocyte tail retraction during transendothelial migration. J Cell Biol 154(1):147-160.

49. Blumenstein L & Ahmadian MR (2004) Models of the cooperative mechanism for Rho effector recognition: implications for RhoA-mediated effector activation. J Biol Chem 279(51):53419-53426.

50. Maekawa M, et al. (1999) Signaling from Rho to the actin cytoskeleton through protein kinases ROCK and LIM-kinase. Science 285(5429):895-898.

51. Watanabe N, et al. (1997) p140mDia, a mammalian homolog of Drosophila diaphanous, is a target protein for Rho small GTPase and is a ligand for profilin. EMBO J 16(11):3044-3056.

52. Debidda M, Wang L, Zang H, Poli V, & Zheng Y (2005) A role of STAT3 in Rho GTPase-regulated cell migration and proliferation. J Biol Chem 280(17):17275-17285.

53. Hodge JC, Bub J, Kaul S, Kajdacsy-Balla A, & Lindholm PF (2003) Requirement of RhoA activity for increased nuclear factor kappaB activity and PC-3 human prostate cancer cell invasion. Cancer Res 63(6):1359-1364.

54. Gotoh Y, et al. (2004) Protein kinase PKN1 associates with TRAF2 and is involved in TRAF2-NF-kappaB signaling pathway. Biochem Biophys Res Commun 314(3):688-694.

55. Buchanan FG, Elliot CM, Gibbs M, & Exton JH (2000) Translocation of the Rac1 guanine nucleotide exchange factor Tiam1 induced by platelet-derived growth factor and lysophosphatidic acid. J Biol Chem 275(13):9742-9748.

56. Gakidis MA, et al. (2004) Vav GEFs are required for beta2 integrin-dependent functions of neutrophils. J Cell Biol 166(2):273-282.

57. Hamelers IH, et al. (2005) The Rac activator Tiam1 is required for (alpha)3(beta)1-mediated laminin-5 deposition, cell spreading, and cell migration. J Cell Biol 171(5):871-881.

58. Chen Z, et al. (2010) Structure and control of the actin regulatory WAVE complex. Nature 468(7323):533-538.

112

59. Cheng L, Mahon GM, Kostenko EV, & Whitehead IP (2004) Pleckstrin homology domain-mediated activation of the rho-specific guanine nucleotide exchange factor Dbs by Rac1. J Biol Chem 279(13):12786-12793.

60. Klinger MB, Guilbault B, & Kay RJ (2004) The RhoA- and CDC42-specific exchange factor Dbs promotes expansion of immature thymocytes and deletion of double-positive and single-positive thymocytes. Eur J Immunol 34(3):806-816.

61. Nimnual AS, Taylor LJ, & Bar-Sagi D (2003) Redox-dependent downregulation of Rho by Rac. Nat Cell Biol 5(3):236-241.

62. Vidal C, Geny B, Melle J, Jandrot-Perrus M, & Fontenay-Roupie M (2002) Cdc42/Rac1-dependent activation of the p21-activated kinase (PAK) regulates human platelet lamellipodia spreading: implication of the cortical-actin binding protein cortactin. Blood 100(13):4462-4469.

63. Tang Y, et al. (1998) A role for Pak protein kinases in Schwann cell transformation. Proc Natl Acad Sci U S A 95(9):5139-5144.

64. Ward ME, Wu JY, & Rao Y (2004) Visualization of spatially and temporally regulated N-WASP activity during cytoskeletal reorganization in living cells. Proc Natl Acad Sci U S A 101(4):970-974.

65. Takenawa T & Miki H (2001) WASP and WAVE family proteins: key molecules for rapid rearrangement of cortical actin filaments and cell movement. J Cell Sci 114(Pt 10):1801-1809.

66. Takenawa T & Suetsugu S (2007) The WASP-WAVE protein network: connecting the membrane to the cytoskeleton. Nat Rev Mol Cell Biol 8(1):37-48.

67. Schirenbeck A, Bretschneider T, Arasada R, Schleicher M, & Faix J (2005) The Diaphanous-related formin dDia2 is required for the formation and maintenance of filopodia. Nat Cell Biol 7(6):619-625.

68. Baird D, Feng Q, & Cerione RA (2005) The Cool-2/alpha-Pix protein mediates a Cdc42-Rac signaling cascade. Curr Biol 15(1):1-10.

69. Pertz O, Hodgson L, Klemke RL, & Hahn KM (2006) Spatiotemporal dynamics of RhoA activity in migrating cells. Nature 440(7087):1069-1072.

70. Renkawitz J & Sixt M (2010) Mechanisms of force generation and force transmission during interstitial leukocyte migration. EMBO Rep 11(10):744-750.

71. Torka R, Thuma F, Herzog V, & Kirfel G (2006) ROCK signaling mediates the adoption of different modes of migration and invasion in human mammary epithelial tumor cells. Exp Cell Res 312(19):3857-3871.

72. Charras G & Paluch E (2008) Blebs lead the way: how to migrate without lamellipodia. Nat Rev Mol Cell Biol 9(9):730-736.

73. Fackler OT & Grosse R (2008) Cell motility through plasma membrane blebbing. J Cell Biol 181(6):879-884.

74. Sahai E & Marshall CJ (2003) Differing modes of tumour cell invasion have distinct requirements for Rho/ROCK signalling and extracellular proteolysis. Nat Cell Biol 5(8):711-719.

75. Tournaviti S, et al. (2007) SH4-domain-induced plasma membrane dynamization promotes bleb-associated cell motility. J Cell Sci 120(Pt 21):3820-3829.

76. Rosel D, et al. (2008) Up-regulation of Rho/ROCK signaling in sarcoma cells drives invasion and increased generation of protrusive forces. Mol Cancer Res 6(9):1410-1420.

113

77. Albiges-Rizo C, Destaing O, Fourcade B, Planus E, & Block MR (2009) Actin machinery and mechanosensitivity in invadopodia, podosomes and focal adhesions. J Cell Sci 122(Pt 17):3037-3049.

78. Machacek M, et al. (2009) Coordination of Rho GTPase activities during cell protrusion. Nature 461(7260):99-103.

79. Vega FM, Fruhwirth G, Ng T, & Ridley AJ (2011) RhoA and RhoC have distinct roles in migration and invasion by acting through different targets. J Cell Biol 193(4):655-665.

80. Lammermann T, et al. (2008) Rapid leukocyte migration by integrin-independent flowing and squeezing. Nature 453(7191):51-55.

81. Kardash E, et al. (2010) A role for Rho GTPases and cell-cell adhesion in single-cell motility in vivo. Nat Cell Biol 12(1):47-53; sup pp 41-11.

82. Blaser H, et al. (2006) Migration of zebrafish primordial germ cells: a role for myosin contraction and cytoplasmic flow. Dev Cell 11(5):613-627.

83. Tinevez JY, et al. (2009) Role of cortical tension in bleb growth. Proc Natl Acad Sci U S A 106(44):18581-18586.

84. Charras GT, Hu CK, Coughlin M, & Mitchison TJ (2006) Reassembly of contractile actin cortex in cell blebs. J Cell Biol 175(3):477-490.

85. Asnaghi L, et al. (2010) E-cadherin negatively regulates neoplastic growth in non-small cell lung cancer: role of Rho GTPases. Oncogene 29(19):2760-2771.

86. Sanz-Moreno V, et al. (2008) Rac activation and inactivation control plasticity of tumor cell movement. Cell 135(3):510-523.

87. Itoh RE, et al. (2002) Activation of rac and cdc42 video imaged by fluorescent resonance energy transfer-based single-molecule probes in the membrane of living cells. Mol Cell Biol 22(18):6582-6591.

88. Kraynov VS, et al. (2000) Localized Rac activation dynamics visualized in living cells. Science 290(5490):333-337.

89. Worthylake RA & Burridge K (2003) RhoA and ROCK promote migration by limiting membrane protrusions. J Biol Chem 278(15):13578-13584.

90. Desai LP, Aryal AM, Ceacareanu B, Hassid A, & Waters CM (2004) RhoA and Rac1 are both required for efficient wound closure of airway epithelial cells. Am J Physiol Lung Cell Mol Physiol 287(6):L1134-1144.

91. Hirakawa M, et al. (2007) Protein kinase A inhibits lysophosphatidic acid-induced migration of airway smooth muscle cells. J Pharmacol Exp Ther 321(3):1102-1108.

92. Liang CC, Park AY, & Guan JL (2007) In vitro scratch assay: a convenient and inexpensive method for analysis of cell migration in vitro. Nat Protoc 2(2):329-333.

93. Ridley AJ, Paterson HF, Johnston CL, Diekmann D, & Hall A (1992) The small GTP-binding protein rac regulates growth factor-induced membrane ruffling. Cell 70(3):401-410.

94. Schwartz MA, Meredith JE, & Kiosses WB (1998) An activated Rac mutant functions as a dominant negative for membrane ruffling. Oncogene 17(5):625-629.

95. Ren XD, Kiosses WB, & Schwartz MA (1999) Regulation of the small GTP-binding protein Rho by cell adhesion and the cytoskeleton. EMBO J 18(3):578-585.

114

96. Burbelo PD, Drechsel D, & Hall A (1995) A conserved binding motif defines numerous candidate target proteins for both Cdc42 and Rac GTPases. J Biol Chem 270(49):29071-29074.

97. Berridge MJ, Bootman MD, & Roderick HL (2003) Calcium signalling: dynamics, homeostasis and remodelling. Nat Rev Mol Cell Biol 4(7):517-529.

98. Bootman MD, et al. (2001) Calcium signalling--an overview. Semin Cell Dev Biol 12(1):3-10.

99. Ralevic V & Burnstock G (1998) Receptors for purines and pyrimidines. Pharmacol Rev 50(3):413-492.

100. Maurice DH, Waldo GL, Morris AJ, Nicholas RA, & Harden TK (1993) Identification of G alpha 11 as the phospholipase C-activating G-protein of turkey erythrocytes. Biochem J 290 ( Pt 3):765-770.

101. Berridge MJ & Irvine RF (1989) Inositol phosphates and cell signalling. Nature 341(6239):197-205.

102. Huang GN, et al. (2006) STIM1 carboxyl-terminus activates native SOC, I(crac) and TRPC1 channels. Nat Cell Biol 8(9):1003-1010.

103. Morgan AJ & Jacob R (1994) Ionomycin enhances Ca2+ influx by stimulating store-regulated cation entry and not by a direct action at the plasma membrane. Biochem J 300 ( Pt 3):665-672.

104. Ruch C, Skiniotis G, Steinmetz MO, Walz T, & Ballmer-Hofer K (2007) Structure of a VEGF-VEGF receptor complex determined by electron microscopy. Nat Struct Mol Biol 14(3):249-250.

105. Bruegmann T, et al. (2010) Optogenetic control of heart muscle in vitro and in vivo. Nat Methods 7(11):897-900.

106. Perrin BJ & Huttenlocher A (2002) Calpain. Int J Biochem Cell Biol 34(7):722-725. 107. De Koninck P & Schulman H (1998) Sensitivity of CaM kinase II to the frequency of

Ca2+ oscillations. Science 279(5348):227-230. 108. Wei C, et al. (2009) Calcium flickers steer cell migration. Nature 457(7231):901-905. 109. Schwaller B, Meyer M, & Schiffmann S (2002) 'New' functions for 'old' proteins: the role

of the calcium-binding proteins calbindin D-28k, calretinin and parvalbumin, in cerebellar physiology. Studies with knockout mice. Cerebellum 1(4):241-258.

110. Berridge MJ (2006) Calcium microdomains: organization and function. Cell Calcium 40(5-6):405-412.

111. Watras J, Benevolensky D, & Childs C (1989) Calcium release from aortic sarcoplasmic reticulum. J Mol Cell Cardiol 21 Suppl 1:125-130.

112. Teramoto T & Iwasaki K (2006) Intestinal calcium waves coordinate a behavioral motor program in C. elegans. Cell Calcium 40(3):319-327.

113. Olwin BB & Storm DR (1985) Calcium binding to complexes of calmodulin and calmodulin binding proteins. Biochemistry 24(27):8081-8086.

114. Yap KL, et al. (2000) Calmodulin target database. J Struct Funct Genomics 1(1):8-14. 115. Yamniuk AP & Vogel HJ (2004) Calmodulin's flexibility allows for promiscuity in its

interactions with target proteins and peptides. Mol Biotechnol 27(1):33-57. 116. Bahler M & Rhoads A (2002) Calmodulin signaling via the IQ motif. FEBS Lett

513(1):107-113.

115

117. Lukas TJ, Burgess WH, Prendergast FG, Lau W, & Watterson DM (1986) Calmodulin binding domains: characterization of a phosphorylation and calmodulin binding site from myosin light chain kinase. Biochemistry 25(6):1458-1464.

118. Osawa M, et al. (1999) A novel target recognition revealed by calmodulin in complex with Ca2+-calmodulin-dependent kinase kinase. Nat Struct Biol 6(9):819-824.

119. Trybus KM, et al. (2007) Effect of calcium on calmodulin bound to the IQ motifs of myosin V. J Biol Chem 282(32):23316-23325.

120. Parekh AB, Foguet M, Lubbert H, & Stuhmer W (1993) Ca2+ oscillations and Ca2+ influx in Xenopus oocytes expressing a novel 5-hydroxytryptamine receptor. J Physiol 469:653-671.

121. Gee KR, et al. (2000) Chemical and physiological characterization of fluo-4 Ca(2+)-indicator dyes. Cell Calcium 27(2):97-106.

122. Kao JP & Tsien RY (1988) Ca2+ binding kinetics of fura-2 and azo-1 from temperature-jump relaxation measurements. Biophys J 53(4):635-639.

123. Knauer SK, et al. (2005) Translocation biosensors to study signal-specific nucleo-cytoplasmic transport, protease activity and protein-protein interactions. Traffic 6(7):594-606.

124. Mills E & Truong K (2010) Engineering Ca2+/calmodulin-mediated modulation of protein translocation by overlapping binding and signaling peptide sequences. Cell Calcium 47(4):369-377.

125. Oancea E, Teruel MN, Quest AF, & Meyer T (1998) Green fluorescent protein (GFP)-tagged cysteine-rich domains from protein kinase C as fluorescent indicators for diacylglycerol signaling in living cells. J Cell Biol 140(3):485-498.

126. Muto A, et al. (2011) Genetic visualization with an improved GCaMP calcium indicator reveals spatiotemporal activation of the spinal motor neurons in zebrafish. Proc Natl Acad Sci U S A 108(13):5425-5430.

127. Mank M, et al. (2006) A FRET-based calcium biosensor with fast signal kinetics and high fluorescence change. Biophys J 90(5):1790-1796.

128. Truong K, Khorchid A, & Ikura M (2003) A fluorescent cassette-based strategy for engineering multiple domain fusion proteins. BMC Biotechnol 3:8.

129. Bolte S & Cordelieres FP (2006) A guided tour into subcellular colocalization analysis in light microscopy. J Microsc 224(Pt 3):213-232.

130. Mills E, Pham E, & Truong K (2010) Structure based design of a Ca2+-sensitive RhoA protein that controls cell morphology. Cell Calcium 48(4):195-201.

131. Yeh BJ, Rutigliano RJ, Deb A, Bar-Sagi D, & Lim WA (2007) Rewiring cellular morphology pathways with synthetic guanine nucleotide exchange factors. Nature 447(7144):596-600.

132. Leung DW, Otomo C, Chory J, & Rosen MK (2008) Genetically encoded photoswitching of actin assembly through the Cdc42-WASP-Arp2/3 complex pathway. Proc Natl Acad Sci U S A 105(35):12797-12802.

133. Ihara K, et al. (1998) Crystal structure of human RhoA in a dominantly active form complexed with a GTP analogue. J Biol Chem 273(16):9656-9666.

134. Meador WE, Means AR, & Quiocho FA (1993) Modulation of calmodulin plasticity in molecular recognition on the basis of x-ray structures. Science 262(5140):1718-1721.

116

135. Meador WE, Means AR, & Quiocho FA (1992) Target enzyme recognition by calmodulin: 2.4 A structure of a calmodulin-peptide complex. Science 257(5074):1251-1255.

136. Pham E, Chiang J, Li I, Shum W, & Truong K (2007) A computational tool for designing FRET protein biosensors by rigid-body sampling of their conformational space. Structure 15(5):515-523.

137. Reid DG, et al. (1990) A proton nuclear magnetic resonance and molecular modeling study of calmidazolium (R24571) binding to calmodulin and skeletal muscle troponin C. J Biol Chem 265(17):9744-9753.

138. Coleman ML, et al. (2001) Membrane blebbing during apoptosis results from caspase-mediated activation of ROCK I. Nat Cell Biol 3(4):339-345.

139. Kimura K, et al. (1996) Regulation of myosin phosphatase by Rho and Rho-associated kinase (Rho-kinase). Science 273(5272):245-248.

140. Elliott MR, et al. (2009) Nucleotides released by apoptotic cells act as a find-me signal to promote phagocytic clearance. Nature 461(7261):282-286.

141. Mills E & Truong K (2011) Ca(2+)-mediated synthetic biosystems offer protein design versatility, signal specificity, and pathway rewiring. Chem Biol 18(12):1611-1619.

142. Fiser A, Do RK, & Sali A (2000) Modeling of loops in protein structures. Protein Sci 9(9):1753-1773.

143. Ikura M, et al. (1992) Solution structure of a calmodulin-target peptide complex by multidimensional NMR. Science 256(5057):632-638.

144. Houdusse A, et al. (2006) Crystal structure of apo-calmodulin bound to the first two IQ motifs of myosin V reveals essential recognition features. Proc Natl Acad Sci U S A 103(51):19326-19331.

145. Sali A & Blundell TL (1993) Comparative protein modelling by satisfaction of spatial restraints. J Mol Biol 234(3):779-815.

146. Correia SS, et al. (2008) Motor protein-dependent transport of AMPA receptors into spines during long-term potentiation. Nat Neurosci 11(4):457-466.

147. Robinson DN, Cavet G, Warrick HM, & Spudich JA (2002) Quantitation of the distribution and flux of myosin-II during cytokinesis. BMC Cell Biol 3:4.

148. Jurado LA, Chockalingam PS, & Jarrett HW (1999) Apocalmodulin. Physiol Rev 79(3):661-682.

149. Palmer AE, et al. (2006) Ca2+ indicators based on computationally redesigned calmodulin-peptide pairs. Chem Biol 13(5):521-530.

150. Bardet PL, et al. (2008) A fluorescent reporter of caspase activity for live imaging. Proc Natl Acad Sci U S A 105(37):13901-13905.

151. Zhou Y, et al. (2010) STIM1 gates the store-operated calcium channel ORAI1 in vitro. Nat Struct Mol Biol 17(1):112-116.

152. Xu P, et al. (2006) Aggregation of STIM1 underneath the plasma membrane induces clustering of Orai1. Biochem Biophys Res Commun 350(4):969-976.

153. Bucurenciu I, Kulik A, Schwaller B, Frotscher M, & Jonas P (2008) Nanodomain coupling between Ca2+ channels and Ca2+ sensors promotes fast and efficient transmitter release at a cortical GABAergic synapse. Neuron 57(4):536-545.

117

154. Jacob SN, et al. (2005) Signaling microdomains regulate inositol 1,4,5-trisphosphate-mediated intracellular calcium transients in cultured neurons. J Neurosci 25(11):2853-2864.

155. Llense F & Martin-Blanco E (2008) JNK signaling controls border cell cluster integrity and collective cell migration. Curr Biol 18(7):538-544.

156. Friedl P & Gilmour D (2009) Collective cell migration in morphogenesis, regeneration and cancer. Nat Rev Mol Cell Biol 10(7):445-457.

157. Hidalgo-Carcedo C, et al. (2011) Collective cell migration requires suppression of actomyosin at cell-cell contacts mediated by DDR1 and the cell polarity regulators Par3 and Par6. Nat Cell Biol 13(1):49-58.

158. Wolf K, et al. (2007) Multi-step pericellular proteolysis controls the transition from individual to collective cancer cell invasion. Nat Cell Biol 9(8):893-904.

159. Tinhofer I, et al. (1996) Differential Ca2+ signaling induced by activation of the epidermal growth factor and nerve growth factor receptors. J Biol Chem 271(48):30505-30509.

160. Ellis-Davies GC (2007) Caged compounds: photorelease technology for control of cellular chemistry and physiology. Nat Methods 4(8):619-628.

161. Collins SR & Meyer T (2011) Evolutionary origins of STIM1 and STIM2 within ancient Ca2+ signaling systems. Trends Cell Biol 21(4):202-211.

162. Vetter SW & Leclerc E (2003) Novel aspects of calmodulin target recognition and activation. Eur J Biochem 270(3):404-414.

163. Tadross MR, Dick IE, & Yue DT (2008) Mechanism of local and global Ca2+ sensing by calmodulin in complex with a Ca2+ channel. Cell 133(7):1228-1240.

164. Carragher NO, et al. (2006) Calpain 2 and Src dependence distinguishes mesenchymal and amoeboid modes of tumour cell invasion: a link to integrin function. Oncogene 25(42):5726-5740.

165. Yoshida K & Soldati T (2006) Dissection of amoeboid movement into two mechanically distinct modes. J Cell Sci 119(Pt 18):3833-3844.

166. Hynes RO (2002) Integrins: bidirectional, allosteric signaling machines. Cell 110(6):673-687.

167. Panopoulos A, Howell M, Fotedar R, & Margolis RL (2011) Glioblastoma motility occurs in the absence of actin polymer. Mol Biol Cell 22(13):2212-2220.

168. Xu J, et al. (2003) Divergent signals and cytoskeletal assemblies regulate self-organizing polarity in neutrophils. Cell 114(2):201-214.

169. Vitorino P & Meyer T (2008) Modular control of endothelial sheet migration. Genes Dev 22(23):3268-3281.

170. Ewald AJ, Brenot A, Duong M, Chan BS, & Werb Z (2008) Collective epithelial migration and cell rearrangements drive mammary branching morphogenesis. Dev Cell 14(4):570-581.

171. Hegerfeldt Y, Tusch M, Brocker EB, & Friedl P (2002) Collective cell movement in primary melanoma explants: plasticity of cell-cell interaction, beta1-integrin function, and migration strategies. Cancer Res 62(7):2125-2130.

172. Mills E, Pham, E., Nagaraj, S. and Truong K. (2012) Engineered networks of natural and synthetic proteins to control cell migration. ACS Synth Biol. Accepted, in press.

118

173. Modha R, et al. (2008) The Rac1 polybasic region is required for interaction with its effector PRK1. J Biol Chem 283(3):1492-1500.

174. Dvorsky R, Blumenstein L, Vetter IR, & Ahmadian MR (2004) Structural insights into the interaction of ROCKI with the switch regions of RhoA. J Biol Chem 279(8):7098-7104.

175. Inoue T, Heo WD, Grimley JS, Wandless TJ, & Meyer T (2005) An inducible translocation strategy to rapidly activate and inhibit small GTPase signaling pathways. Nat Methods 2(6):415-418.

176. Sandrock K, Bielek H, Schradi K, Schmidt G, & Klugbauer N (2010) The nuclear import of the small GTPase Rac1 is mediated by the direct interaction with karyopherin alpha2. Traffic 11(2):198-209.

177. Grunwald D, Singer RH, & Rout M (2011) Nuclear export dynamics of RNA-protein complexes. Nature 475(7356):333-341.

178. Price LS, et al. (2003) Calcium signaling regulates translocation and activation of Rac. J Biol Chem 278(41):39413-39421.

179. Kovacs M, Toth J, Hetenyi C, Malnasi-Csizmadia A, & Sellers JR (2004) Mechanism of blebbistatin inhibition of myosin II. J Biol Chem 279(34):35557-35563.

180. Straight AF, et al. (2003) Dissecting temporal and spatial control of cytokinesis with a myosin II Inhibitor. Science 299(5613):1743-1747.

181. Thomas P & Smart TG (2005) HEK293 cell line: a vehicle for the expression of recombinant proteins. J Pharmacol Toxicol Methods 51(3):187-200.

182. Amano M, et al. (1996) Phosphorylation and activation of myosin by Rho-associated kinase (Rho-kinase). J Biol Chem 271(34):20246-20249.

183. Baumann BA, Hambly BD, Hideg K, & Fajer PG (2001) The regulatory domain of the myosin head behaves as a rigid lever. Biochemistry 40(26):7868-7873.

184. Ikebe M, et al. (1998) A hinge at the central helix of the regulatory light chain of myosin is critical for phosphorylation-dependent regulation of smooth muscle myosin motor activity. J Biol Chem 273(28):17702-17707.

185. Baumann BA, et al. (2012) Phosphorylated smooth muscle heavy meromyosin shows an open conformation linked to activation. J Mol Biol 415(2):274-287.

186. Sakamoto T, Limouze J, Combs CA, Straight AF, & Sellers JR (2005) Blebbistatin, a myosin II inhibitor, is photoinactivated by blue light. Biochemistry 44(2):584-588.

187. Muranyi A, et al. (2002) Phosphorylation of the myosin phosphatase target subunit by integrin-linked kinase. Biochem J 366(Pt 1):211-216.

188. Yoshida H, Okumura N, Kitagishi Y, Shirafuji N, & Matsuda S (2010) Rab5(Q79L) interacts with the carboxyl terminus of RUFY3. Int J Biol Sci 6(2):187-189.

189. Dao VT, Dupuy AG, Gavet O, Caron E, & de Gunzburg J (2009) Dynamic changes in Rap1 activity are required for cell retraction and spreading during mitosis. J Cell Sci 122(Pt 16):2996-3004.

190. Norton LW, Tegnell E, Toporek SS, & Reichert WM (2005) In vitro characterization of vascular endothelial growth factor and dexamethasone releasing hydrogels for implantable probe coatings. Biomaterials 26(16):3285-3297.

191. Tufro A (2000) VEGF spatially directs angiogenesis during metanephric development in vitro. Dev Biol 227(2):558-566.

119

192. Szabo E, et al. (2010) Direct conversion of human fibroblasts to multilineage blood progenitors. Nature 468(7323):521-526.

193. Zhao T, Zhang ZN, Rong Z, & Xu Y (2011) Immunogenicity of induced pluripotent stem cells. Nature 474(7350):212-215.

194. Hu CM, Zhang L, Aryal S, Cheung C, & Fang RH (2011) Erythrocyte membrane-camouflaged polymeric nanoparticles as a biomimetic delivery platform. Proc Natl Acad Sci U S A 108(27):10980-10985.

195. Fazakas C, et al. (2011) Transmigration of melanoma cells through the blood-brain barrier: role of endothelial tight junctions and melanoma-released serine proteases. PLoS One 6(6):e20758.

196. Cayrol R, et al. (2008) Activated leukocyte cell adhesion molecule promotes leukocyte trafficking into the central nervous system. Nat Immunol 9(2):137-145.

197. Dalerba P, et al. (2011) Single-cell dissection of transcriptional heterogeneity in human colon tumors. Nat Biotechnol 29(12):1120-1127.

198. Salsman J, Top D, Barry C, & Duncan R (2008) A virus-encoded cell-cell fusion machine dependent on surrogate adhesins. PLoS Pathog 4(3):e1000016.

199. Mangeot PE, et al. (2011) Protein transfer into human cells by VSV-G-induced nanovesicles. Mol Ther 19(9):1656-1666.

200. Collins RN (2003) Rab and ARF GTPase regulation of exocytosis. Mol Membr Biol 20(2):105-115.

201. Muller A, et al. (2005) Endogenous Ca2+ buffer concentration and Ca2+ microdomains in hippocampal neurons. J Neurosci 25(3):558-565.

202. Kim E, et al. (2007) Characterization of human cardiac calsequestrin and its deleterious mutants. J Mol Biol 373(4):1047-1057.

203. Pauls TL, Durussel I, Berchtold MW, & Cox JA (1994) Inactivation of individual Ca(2+)-binding sites in the paired EF-hand sites of parvalbumin reveals asymmetrical metal-binding properties. Biochemistry 33(34):10393-10400.

204. Tabor JJ, Levskaya A, & Voigt CA (2011) Multichromatic control of gene expression in Escherichia coli. J Mol Biol 405(2):315-324.

205. Takaishi K, Sasaki T, Kotani H, Nishioka H, & Takai Y (1997) Regulation of cell-cell adhesion by rac and rho small G proteins in MDCK cells. J Cell Biol 139(4):1047-1059.

206. Ambruso DR, et al. (2000) Human neutrophil immunodeficiency syndrome is associated with an inhibitory Rac2 mutation. Proc Natl Acad Sci U S A 97(9):4654-4659.

207. Knaus UG, Heyworth PG, Evans T, Curnutte JT, & Bokoch GM (1991) Regulation of phagocyte oxygen radical production by the GTP-binding protein Rac 2. Science 254(5037):1512-1515.

208. Samson T, Welch C, Monaghan-Benson E, Hahn KM, & Burridge K (2010) Endogenous RhoG is rapidly activated after epidermal growth factor stimulation through multiple guanine-nucleotide exchange factors. Mol Biol Cell 21(9):1629-1642.

209. Karginov AV, Ding F, Kota P, Dokholyan NV, & Hahn KM (2010) Engineered allosteric activation of kinases in living cells. Nat Biotechnol 28(7):743-747.

120

Appendices

These appendices contain data on gene and protein sequences, a legend for supplemental movies

and additional experimental results that may be valuable to the interested reader.

Appendix Page

Appendix A: Gene and Protein Sequences 121

Appendix B: List of Oligonucleotide Primers and Sequences 127

Appendix C: Supplemental Movie Legends 128

Appendix D: Additional Control Experiments for CaM-RhoA(DP) 130

Appendix E: Supplemental Colocalization Analysis of RBD/PBD probes 132

Appendix F: Behaviour of CaRQ in HeLa and CHO Cells 133

Appendix G: Mutations in Rac1 and Cdc42 to Introduce a CaM Binding Site 135

Appendix H: Ca2+ Control Designs for Rac1 and Cdc42 with PBD 137

Appendix I: Area Thresholding Analysis Applied to Blebbing Cells 139

121

Appendix A: Gene and Protein Sequences

Nucleotide and amino acid sequences for the main proteins created and domains used in this

work are given below in the standard one letter code. Sequences are not given for proteins that

were used as developed elsewhere (e.g. LOVS1K, nAChR-α4, ChR2, etc) or were modified

simply by adding a fluorescent protein (e.g. VEGFR2). The sequences of fluorescent protein

tags have not been included.

Amino acid sequence legend: CaM Rho MLCKp IQp

CaM-RhoA(DP)

Nucleotide Sequence:

CCATGGGCCTGACTAGTGAGCAGATTGCAGAGTTCAAAGAAGCCTTCTCATTATTCGACAAGGATGGGGACGGCACCA

TCACCACAAAGGAACTTGGCACCGTTATGAGGTCGCTTGGACAAAACCCAACGGAAGCAGAATTGCAGGATATGATCA

ATGAAGTCGATGCTGATGGCAATGGAACGATTTACTTTCCTGAATTTCTTACTATGATGGCTAGAAAAATGAAGGACA

CAGACAGCGAAGAGGAAATCCGAGAAGCATTCCGTGTTTTTGACAAGGATGGGAACGGCTACATCAGCGCTGCTGAAT

TACGTCACGTCATGACAAACCTCGGGGAGAAGTTAACAGATGAAGAAGTTGATGAAATGATAAGGGAAGCAGATATCG

ATGGTGATGGCCAAGTAAACTATGAAGAGTTTGTACAAATGATGACAGCAAAGGCTAGTGCTGCCATCCGGAAGAAAC

TGGTGATTGTTGGTGATGGAGCCTGTGGAAAGACATGCTTGCTCATAGTCTTCAGCAAGGACCAGTTCCCAGAGGTGT

ATGTGCCCACAGTGTTTGAGAACTATGTGGCAGATATCGAGGTGGATGGAAAGCAGGTAGAGTTGGCTTTGTGGGACA

CAGCTGGGCTGGAAGATTATGATCGCCTGAGGCCCCTCTCCTACCCAGATACCGATGTTATACTGATGTGTTTTTCCA

TCGACAGCCCTGATAGTTTAGAAAACATCCCAGAAAAGTGGACCCCAGAAGTCAAGCATTTCTGTCCCAACGTGCCCA

TCATCCTGGTTGGGAATAAGAAGGATCTTCGGAATGATGAGCACACAAGGCGGGAGCTAGCCAAGATGAAGCAGGAGC

CGGTGAAACCTGAAGAAGGCAGAGATATGGCAAACAGGATTGGCGCTTTTGGGTACATGGAGTGTTCAGCAAAGACCA

AAGATGGAGTGAGAGAGGTTTTTGAAATGGCTACGAGAGCTGCTCTGCAAGCTAGATCTCCCGGGGCTAGC

Amino Acid Sequence:

MGLTSEQIAEFKEAFSLFDKDGDGTITTKELGTVMRSLGQNPTEAELQDMINEVDADGNGTIYF

PEFLTMMARKMKDTDSEEEIREAFRVFDKDGNGYISAAELRHVMTNLGEKLTDEEVDEMIREAD

IDGDGQVNYEEFVQMMTAKASAAIRKKLVIVGDGACGKTCLLIVFSKDQFPEVYVPTVFENYVA

DIEVDGKQVELALWDTAGLEDYDRLRPLSYPDTDVILMCFSIDSPDSLENIPEKWTPEVKHFCP

NVPIILVGNKKDLRNDEHTRRELAKMKQEPVKPEEGRDMANRIGAFGYMECSAKTKDGVREVFE

MATRAALQARSPGAS

122

CaRQ

Nucleotide Sequence:

CCATGGGCCTGACTAGTGAGCAGATTGCAGAGTTCAAAGAAGCCTTCTCATTATTCGACAAGGATGGGGACGGCACCA

TCACCACAAAGGAACTTGGCACCGTTATGAGGTCGCTTGGACAAAACCCAACGGAAGCAGAATTGCAGGATATGATCA

ATGAAGTCGATGCTGATGGCAATGGAACGATTTACTTTCCTGAATTTCTTACTATGATGGCTAGAAAAATGAAGGACA

CAGACAGCGAAGAGGAAATCCGAGAAGCATTCCGTGTTTTTGACAAGGATGGGAACGGCTACATCAGCGCTGCTGAAT

TACGTCACGTCATGACAAACCTCGGGGAGAAGTTAACAGATGAAGAAGTTGATGAAATGATAAGGGAAGCAGATATCG

ATGGTGATGGCCAAGTAAACTATGAAGAGTTTGTACAAATGATGACAGCAAAGGCTAGTAAGAGGCGCTGGAAGAAAA

ACTTCATTGCCGTCAGCGCTGCCAACCGGTACAAGAAGATCTCCAGCTCCGGGGCACTGGCTGCCATCCGGAAGAAAC

TGGTGATTGTTGGTGATGGAGCCTGTGGAAAGACATGCTTGCTCATAGTCTTCAGCAAGGACCAGTTCCCAGAGGTGT

ATGTGCCCACAGTGTTTGAGAACTATGTGGCAGATATCGAGGTGGATGCTAGTGGATCCGCTATTACTGTCCAAAGAT

ATGTCAGAGGAATCCAAGCTAGAGCTTATGCTAGATTCCTCGCTAGTTATGATCGCCTGAGGCCCCTCTCCTACCCAG

ATACCGATGTTATACTGATGTGTTTTTCCATCGACAGCCCTGATAGTTTAGAAAACATCCCAGAAAAGTGGACCCCAG

AAGTCAAGCATTTCTGTCCCAACGTGCCCATCATCCTGGTTGGGAATAAGAAGGATCTTCGGAATGATGAGCACACAA

GGCGGGAGCTAGCCAAGATGAAGCAGGAGCCGGTGAAACCTGAAGAAGGCAGAGATATGGCAAACAGGATTGGCGCTT

TTGGGTACATGGAGTGTTCAGCAAAGACCAAAGATGGAGTGAGAGAGGTTTTTGAAATGGCTACGAGAGCTGCTCTGC

AAGCTAGATCTCCCGGGGCTAGC

Amino Acid Sequence:

MGLTSEQIAEFKEAFSLFDKDGDGTITTKELGTVMRSLGQNPTEAELQDMINEVDADGNGTIYF

PEFLTMMARKMKDTDSEEEIREAFRVFDKDGNGYISAAELRHVMTNLGEKLTDEEVDEMIREAD

IDGDGQVNYEEFVQMMTAKASKRRWKKNFIAVSAANRYKKISSSGALAAIRKKLVIVGDGACGK

TCLLIVFSKDQFPEVYVPTVFENYVADIEVDASGSAITVQRYVRGIQARAYARFLASYDRLRPL

SYPDTDVILMCFSIDSPDSLENIPEKWTPEVKHFCPNVPIILVGNKKDLRNDEHTRRELAKMKQ

EPVKPEEGRDMANRIGAFGYMECSAKTKDGVREVFEMATRAALQARSPGAS

RACer (Rac1/IQ2p core)

Nucleotide sequence:

CCATGGGCCTGACTAGTGGATCCCAGGCCATCAAGTGTGTGGTGGTGGGAGACGGAGCTGTAGGTAAAACTTGCCTAC

TGATCAGTTACACAACCAATGCATTTCCTGGAGAATATATCCCTACTGTCTTTGACAATTATTCTGCCAATGTTATGG

TAGATGCTAGTGGATCCGCTATTACTGTCCAAAGATATGTCAGAGGAATCCAAGCTAGAGCTTATGCTAGATTCCTCG

CTAGTGGAAAACCGGTGAATCTGGGCTTATGGGATACAGCTGGACTAGAAGATTATGACAGATTACGCCCCCTATCCT

ATCCGCAAACAGATGTGTTCTTAATTTGCTTTTCCCTTGTGAGTCCTGCATCATTTCATCATGTCCGTGCAAAGTGGT

ATCCTGAGGTGCGGCACCACTGTCCCAACACTCCCATCATCCTAGTGGGAACTAAACTTGATCTTAGGGATGATAAAG

ACACGATCGAGAAACTGAAGGAGAAGAAGCTGACTCCCATCACCTATCCGCAGGGTCTAGCTATGGCTAAGGAGATTG

123

GTGCTGTAAAATACCTGGAGTGCTCGGCGCTCACACAGCGAGGCCTCAAGACAGTGTTTGACGAAGCGATCCGAGCAG

TCCTCTGCCCGCCTCCCGTGAAGAAGAGGAAGAGAAAATGCCTGCTGGCTAGC

Amino acid sequence:

MGLTSGSQAIKCVVVGDGAVGKTCLLISYTTNAFPGEYIPTVFDNYSANVMVDASGSAITVQRY

VRGIQARAYARFLASGKPVNLGLWDTAGLEDYDRLRPLSYPQTDVFLICFSLVSPASFHHVRAK

WYPEVRHHCPNTPIILVGTKLDLRDDKDTIEKLKEKKLTPITYPQGLAMAKEIGAVKYLECSAL

TQRGLKTVFDEAIRAVLCPPPVKKRKRKCLLAS

CaM-Cdc42 Chimera (Cdc42/IQ2p core)


CCATGGGGACTAGTATTAAGTGTGTTGTTGTGGGCGATGGTGCTGTTGGTAAAACATGTCTCCTGATATCCTACACAA

CAAACAAATTTCCATCGGAATATGTACCGACTGTTTTTGACAACTATGCAGTCACAGTTATGATTGGTGCTAGTGGAT

CCGCTATTACTGTCCAAAGATATGTCAGAGGAATCCAAGCTAGAGCTTATGCTAGATTCCTCGCTAGTGGAGAACCAT

ATACTCTTGGACTTTTTGATACTGCAGGGCTAGAGGATTATGACAGATTACGACCGCTGAGTTATCCACAAACAGATG

TATTTCTAGTCTGTTTTTCAGTGGTCTCTCCATCTTCATTTCATCACGTGAAAGAAAAGTGGGTGCCTGAGATAACTC

ACCACTGTCCAAAGACTCCTTTCTTGCTTGTTGGGACTCAAATTGATCTCAGAGATGACCCCTCTACTATTGAGAAAC

TTGCCAAGAACAAACAGAAGCCTATCACTCCAGAGACTGCTGATAAGCTGGCCCGTGACCTGAAGGCTGTCAAGTATG

TGGAGTGTTCTGCACTTACACAGAAAGGCCTAAAGAATGTATTTGACGAAGCAATATTGGCTGCCCTGGAGCCTCCAG

AACCGAAGAAGAGCCGCGCTAGC


MGTSIKCVVVGDGAVGKTCLLISYTTNKFPSEYVPTVFDNYAVTVMIGASGSAITVQRYVRGIQ

ARAYARFLASGEPYTLGLFDTAGLEDYDRLRPLSYPQTDVFLVCFSVVSPSSFHHVKEKWVPEI

THHCPKTPFLLVGTQIDLRDDPSTIEKLAKNKQKPITPETADKLARDLKAVKYVECSALTQKGL

KNVFDEAILAALEPPEPKKSRAS

pLyn


CCATGGGCTGCATCAAGAGCAAGGGCAAGGACAGCGCCACTAGT


MGCIKSKGKDSATS

124

Stim1


GGATCCGATGTATGCGTCCGTCTTGCCCTGTGGCTCCTCTGGGGACTCCTCCTGCACCAGGGCCAGAGCCTCAGCCAT

AGTCACAGTGAGAAGGCGACAGGAACCAGCTCGGGGGCCAACTCTGAGGAGTCCACTGCAGCAGAGTTTTGCCGAATT

GACAAGCCCCTGTGTCACAGTGAGGATGAGAAACTCAGCTTCGAGGCAGTCCGTAACATCCACAAACTGATGGACGAT

GATGCCAATGGTGATGTGGATGTGGAAGAAAGTGATGAGTTCCTGAGGGAAGACCTCAATTACCATGACCCAACAGTG

AAACACAGCACCTTCCATGGTGAGGATAAGCTCATCAGCGTGGAGGACCTGTGGAAGGCATGGAAGTCATCAGAAGTA

TACAATTGGACCGTGGATGAGGTGGTACAGTGGCTGATCACATATGTGGAGCTGCCTCAGTATGAGGAGACCTTCCGG

AAGCTGCAGCTCAGTGGCCATGCCATGCCAAGGCTGGCTGTCACCAACACCACCATGACAGGGACTGTGCTGAAGATG

ACAGACCGGAGTCATCGGCAGAAGCTGCAGCTGAAGGCTCTGGATACAGTGCTCTTTGGGCCTCCTCTCTTGACTCGC

CATAATCACCTCAAGGACTTCATGCTGGTGGTGTCTATCGTTATTGGTGTGGGCGGCTGCTGGTTTGCCTATATCCAG

AACCGTTACTCCAAGGAGCACATGAAGAAGATGATGAAGGACTTGGAGGGGTTACACCGAGCTGAGCAGAGTCTGCAT

GACCTTCAGGAAAGGCTGCACAAGGCCCAGGAGGAGCACCGCACAGTGGAGGTGGAGAAGGTCCATCTGGAAAAGAAG

CTGCGCGATGAGATCAACCTTGCTAAGCAGGAAGCCCAGCGGCTGAAGGAGCTGCGGGAGGGTACTGAGAATGAGCGG

AGCCGCCAAAAATATGCTGAGGAGGAGTTGGAGCAGGTTCGGGAGGCCTTGAGGAAAGCAGAGAAGGAGCTAGAATCT

CACAGCTCATGGTATGCTCCAGAGGCCCTTCAGAAGTGGCTGCAGCTGACACATGAGGTGGAGGTGCAATATTACAAC

ATCAAGAAGCAAAATGCTGAGAAGCAGCTGCTGGTGGCCAAGGAGGGGGCTGAGAAGATAAAAAAGAAGAGAAACACA

CTCTTTGGCACCTTCCACGTGGCCCACAGCTCTTCCCTGGATGATGTAGATCATAAAATTCTAACAGCTAAGCAAGCA

CTGAGCGAGGTGACAGCAGCATTGCGGGAGCGCCTGCACCGCTGGCAACAGATCGAGATCCTCTGTGGCTTCCAGATT

GTCAACAACCCTGGCATCCACTCACTGGTGGCTGCCCTCAACATAGACCCCAGCTGGATGGGCAGTACACGCCCCAAC

CCTGCTCACTTCATCATGACTGACGACGTGGATGACATGGATGAGGAGATTGTGTCTCCCTTGTCCATGCAGTCCCCT

AGCCTGCAGAGCAGTGTTCGGCAGCGCCTGACGGAGCCACAGCATGGCCTGGGATCTCAGAGGGATTTGACCCATTCC

GATTCGGAGTCCTCCCTCCACATGAGTGACCGCCAGCGTGTGGCCCCCAAACCTCCTCAGATGAGCCGTGCTGCAGAC

GAGGCTCTCAATGCCATGACTTCCAATGGCAGCCACCGGCTGATCGAGGGGGTCCACCCAGGGTCTCTGGTGGAGAAA

CTGCCTGACAGCCCTGCCCTGGCCAAGAAGGCATTACTGGCGCTGAACCATGGGCTGGACAAGGCCCACAGCCTGATG

GAGCTGAGCCCCTCAGCCCCACCTGGTGGCTCTCCACATTTGGATTCTTCCCGTTCTCACAGCCCCAGCTCCCCAGAC

CCAGACACACCATCTCCAGTTGGGGACAGCCGAGCCCTGCAAGCCAGCCGAAACACACGCATTCCCCACCTGGCTGGC

AAGAAGGCTGTGGCTGAGGAGGATAATGGCTCTATTGGCGAGGAAACAGACTCCAGCCCAGGCCGGAAGAAGTTTCCT

CTCAAAATCTTTAAGAAGCCTCTTAAGAAGGCTAGC


GSDVCVRLALWLLWGLLLHQGQSLSHSHSEKATGTSSGANSEESTAAEFCRIDKPLCHSEDEKL

SFEAVRNIHKLMDDDANGDVDVEESDEFLREDLNYHDPTVKHSTFHGEDKLISVEDLWKAWKSS

EVYNWTVDEVVQWLITYVELPQYEETFRKLQLSGHAMPRLAVTNTTMTGTVLKMTDRSHRQKLQ

LKALDTVLFGPPLLTRHNHLKDFMLVVSIVIGVGGCWFAYIQNRYSKEHMKKMMKDLEGLHRAE

QSLHDLQERLHKAQEEHRTVEVEKVHLEKKLRDEINLAKQEAQRLKELREGTENERSRQKYAEE

ELEQVREALRKAEKELESHSSWYAPEALQKWLQLTHEVEVQYYNIKKQNAEKQLLVAKEGAEKI

KKKRNTLFGTFHVAHSSSLDDVDHKILTAKQALSEVTAALRERLHRWQQIEILCGFQIVNNPGI

125

HSLVAALNIDPSWMGSTRPNPAHFIMTDDVDDMDEEIVSPLSMQSPSLQSSVRQRLTEPQHGLG

SQRDLTHSDSESSLHMSDRQRVAPKPPQMSRAADEALNAMTSNGSHRLIEGVHPGSLVEKLPDS

PALAKKALLALNHGLDKAHSLMELSPSAPPGGSPHLDSSRSHSPSSPDPDTPSPVGDSRALQAS

RNTRIPHLAGKKAVAEEDNGSIGEETDSSPGRKKFPLKIFKKPLKKAS

Parvalbumin


TCGATGACAGACTTGCTCAGCGCTGAGGACATCAAGAAGGCGATAGGAGCCTTTACTGCTGCAGACTCCTTCGACCA

CAAAAAGTTCTTCCAGATGGTGGGCCTGAAGAAAAAGAGTGCGGATGATGTGAAGAAGGTGTTCCACATTCTGGACA

AAGACAAAAGTGGCTTCATTGAGGAGGATGAGCTGGGGTCCATTCTGAAGGGCTTCTCCTCAGATGCCAGAGACTTG

TCTGCTAAGGAAACAAAGACGCTGATGGCTGCTGGAGACAAGGACGGGGACGGCAAGATTGGGGTTGAAGAATTCTC

CACTCTGGTGGCCGAAAGC


SMTDLLSAEDIKKAIGAFTAADSFDHKKFFQMVGLKKKSADDVKKVFHILDKDKSGFIEEDELG

SILKGFSSDARDLSAKETKTLMAAGDKDGDGKIGVEEFSTLVAES

Rho Binding Domain (RBD)


ACAGAGATGCAGGACAGATTGCACATCCTGGAGGACCTGAATATGCTCTACATTCGGCAGATGGCACTCAGCCTGGA

GGACACGGAGTTGCAGAGGAAGCTAGACCATGAGATCCGGATGAGGGAAGGGGCCTGTAAGCTGCTGGCAGCCTGCT

CCCAGCGAGAGCAGGCTCTGGAGGCCACCAAGAGCCTGCTAGTGTGCAACAGCCGCATCCTCAGCTACATGGGCGAG

CTGCAGCGGCGCAAGGAGGCGCAGGTGCTGGGGAAGACAAGC


ILEDLNMLYIRQMALSLEDTELQRKLDHEIRMREGACKLLAACSQREQALEATKSLLVCNSRIL

SYMGELQRRKEAQVLGKTS

126

p21 Binding Domain (PBD)


TAAGAAGAAAGAGAAAGAGCGGCCAGAGATTTCTCTCCCTTCAGATTTTGAACACACAATTCATGTCGGTTTTGATG

CTGTCACAGGGGAGTTTACGGGAATGCCAGAGCAGTGGGCCCGCTTGCTTCAGACATCAAATATCACTAAGTCGGAG

CAGAAGAAAAACCCGCAGGCTGTTCTGGATGTGTTGGAGTTTTACAACTCGAAGAAGACATCCAACAGCCAGAAATA

CATGAGCTTTACAGATAAGTCA


KKKEKERPEISLPSDFEHTIHVGFDAVTGEFTGMPEQWARLLQTSNITKSEQKKNPQAVLDLEF

YNSKKTSNSQKYMSFTDKS

127

Appendix B: List of Oligonucleotide Primers and Sequences

Oligonucleotide primers used to create the embedded RhoA, Rac1 and Cdc42 fragments,

dominant negative mutants and anti-GAP sinking mutants are given here.

Primers used for point mutations and chimera construction. F1 and F2 refer to the two fragments of each GTPases (N-term and C-term, respectively). DN is that dominant negative T17N (T19N for RhoA) point mutation and AntiGAP is the E91H, N92H point mutation. Primer Name Oligonucleotide Sequence

RhoA_F1_53 CATGCCATGGGGACTAGTGCTGCCATCCGGAAGAAACTGG

RhoA_F1_35 CGGCTAGCATCCACCTCGATATCTGCCACATA

RhoA_F2_53 CATGCCATGGGGACTAGTGGAAAGCAGGTAGAGTTGGCTTTG

RhoA_F2_35 GAAGATCTTCTAGCTTGCAGAGCAGCTCTCG

Rac1_F1_53 CGCGGATCCCAGGCCATCAAGTGTGTGGTGG

Rac1_F1_35 CGGCTAGCATCTACCATAACATTGGCAGAATAATTGTC

Rac1_F2_53 CATGCCATGGGGACTAGTGGAAAACCGGTGAATCTGGGCTTA

Rac1_F2_35 CGGCTAGCCAGCAGGCATTTTCTCTTCCTCTT

Cdc42_F1_53 CATGCCATGGGGACTAGTATTAAGTGTGTTGTTGTGGGCGAT

Cdc42_F1_35 CGGCTAGCACCATTCATAACTGTGACTGCATAG

Cdc42_F2_53 CATGCCATGGGGACTAGTGGAGAACCATATACTCTTGGACTTTT

Cdc42_F2_35 CGGCTAGCGCGGCTCTTCTTCGGTTCTGGA

RhoA_DN_53 CCTGTGGAAAGAACTGCTTGCTC

RhoA_DN_35 GAGCAAGCAGTTCTTTCCACAGG

Rac1_DN_53 GCTGTAGGTAAAAACTGCCTACTGATC

Rac1_DN_35 GATCAGTAGGCAGTTTTTACCTACAGC

Rac1_AntiGAP_53 CCTGCATCATTTCATCATGTCCGTGCAAAG

Rac1_AntiGAP_35 CTTTGCACGGACATGATGAAATGATGCAGG

Cdc42_DN_53 GCTGTTGGTAAAAACTGTCTCCTGATA

Cdc42_DN_35 TATCAGGAGACAGTTTTTACCAACAGC

Cdc42_AntiGAP_53 CCATCTTCATTTCATCACGTGAAAGAAAAG

Cdc42_AntiGAP_35 CTTTTCTTTCACGTGATGAAATGAAGATGG

128

Appendix C: Supplemental Movie Legends

Below is a brief description of the experimental conditions, scale and timescale of the

supplemental movies presented in this work. Movies can be downloaded from the following

URL: http://individual.utoronto.ca/emills, or through the Supporting Online Information at each

article’s respective publication website.

Movies 4.1-4.3:

Movies from Chapter 4 were created using Windows Movie Maker and are saved in .wmv

format. Yellow arrows have been placed in the frames to direct the viewer toward regions of bleb

retraction. Images were captured with 60x objective and the square edge of the frame represents

167 µm. In these movies 1s playback = 1 min elapsed time. ATP/UTP/Ionomycin were added

after 1 minute.

- Movie 4.1: CHO cells were expressing CaM-RhoA(DP) and stimulated with ATP. See also Figure 4.6D-G. - Movie 4.2: HeLa cells were expressing CaM-RhoA(DP) and stimulated with UTP. See also Figure 4.6H-K. - Movie 4.3: HEK293 cells were eexpressing CaM-RhoA(DP) and stimulated with ionomycin. See also Figure 4.6L-O.

Movies 5.1-5.6:

Movies from Chapter 5 were created using MetaMorph Advanced, are saved in .avi format and

are falsely coloured for the viewer. Images were captured with the 40x objective and the square

edge of the frame represents 250 µm. Each movie represents a 25 minute experiment. All cells

are HEK293.

- Movie 5.1: Cells were expressing CaRQ and stimulated with ionomycin after a 5 minute control period (2 s in playback). See also Figure 5.4D. - Movie 5.2: Cells were expressing CaRM and stimulated with ionomycin after 5 minutes. See also Figure 5.4E. - Movie 5.3: Cells were expressing YFP and stimulated with ionomycin after 5 minutes. See also Figure 5.4C.

129

- Movie 5.4: Cells were co-expressing pLyn-CaRQ and nAChR-α4 and stimulated with ACh after 5 minutes. See also Figure 5.7D. - Movie 5.5: Cells were co-expressing CaRQ and nAChR-α4 and stimulated with ACh after 5 minutes. See also Figure 5.7A. - Movie 5.6: Cells were co-expressing Orai1-CFP and Stim1-CaRQ (CFP channel is shown because it best outlines the cell). Tg was added after 5 minutes and CaCl2 was added 5 minutes later (10 minutes since the start of the experiment). See also Figure 5.9C.

Movies 6.1-6.6:

Movies from Chapter 6 were created using MetaMorph Advanced, saved in .avi format and were

captured using the 60x objective. For cells stimulated with ionomycin the movie corresponds to

a 20 minute experiment. All cells shown in movies were HEK293.

- Movie 6.1: Cells were expressing RACer and stimulated with ionomycin after a 5 minute control period (corresponds to 2 s in playback). See also Figure 6.4B. - Movie 6.2: Cells were expressing RACer(T17N) and stimulated with ionomycin after 5 minutes. See also Figure 6.4D. - Movie 6.3: Cells were co-expressing RACer/LOVS1K/Orai1 and were illuminated with CFP-excitation light (300 ms per 10 s) for the first 30 minutes of the experiment. The entire movie corresponds to 50 minutes of elapsed time. See also Figure 6.7B. - Movie 6.4: Cells were co-expressing RACer(T17N)/LOVS1K/Orai1 and were illuminated with CFP-excitation light (300 ms per 10 s) for the duration. The movie corresponds to 30 minutes of elapsed time. - Movie 6.5: Cells were expressing the CaM-Cdc42 chimera and were stimulated with ionomycin after 5 minutes. See also Figure 6.10D. - Movie 6.6: Cells were expressing the CaM-Cdc42(T17N) chimera and were stimulated with ionomycin after 5 minutes.

130

Appendix D: Additional Control Experiments for CaM-RhoA(DP)

Characterization and control experiments for the CaM-RhoA(DP)-YFP fusion protein were

carried out in each of the three cell lines used in Chapter 4 including HeLa, CHO and HEK293

cells. The chart given in Figure 4.7 shows the results of those experiments in CHO cells. In this

appendix, the analogous data is presented for experiments in HeLa (Figure D.1) and HEK293

(Figure D.2) cells.

Figure D.1: Bleb retraction in HeLa cells. The number of blebs was counted before (black bars) and 20 minutes after stimulation (grey bars); the percentage change was calculated (white bars). All data are the mean of at least 10 cells over at least 3 independent experiments and the error bars are the standard deviation. * denotes P = 0.003 and ** denotes P < 0.001 by paired t-test and *** denotes P < 0.001 by unpaired t-test. Note Venus = YFP.

131

Figure D.2: Bleb retraction in HEK293 cells. The number of blebs was counted before (black bars) and 20 minutes after stimulation (grey bars); the percentage change was calculated (white bars). All data are the mean of at least 10 cells over at least 3 independent experiments and the error bars are the standard deviation. * denotes P < 0.001 for paired t-test (top) and unpaired t-test (bottom). Note Venus = YFP.

132

Appendix E: Supplemental Colocalization Analysis of RBD/PBD Probes

Colocalization between Ca2+-sensitive Rho chimeras and RBD or PBD-based probes was used as

a supplemental assay, together with morphology changes, to verify the activity of the Ca2+-

sensitive chimeras. Throughout, background-subtracted PC was used in a qualitative way to

provide a binary-type response to the question of whether there was a change in probe-chimera

colocalization. Below, two other methods of colocalization analysis are presented using the

JACoP ImageJ plugin (129), which generates Van Steenel’s cross-correlation function (CCF)

and PC with Costes’s thresholding. In both cases, the magnitude of the percentage change

before and after Ca2+ stimulus, compared to the background subtracted PC, is less. However, in

all cases the relative magnitude of changes when comparing the T19N/T17N mutants with the

indicated chimera is preserved (that is, changes for the DN mutant chimeras are less than

changes for the DP chimeras in all cases, regardless of analysis method).

Table E.1: Comparison of colocalization analysis for Ca2+-sensitive chimeras. Data presented is for the images displayed in the relevant section of the document, with RBD or PBD co-transfection as appropriate. The time indicated is relative to ionomycin addition.

133

Appendix F: Behaviour of CaRQ in HeLa and CHO Cells

Characterization and control experiments of CaRQ and some related constructs were performed

in HeLa and CHO cells in addition to the characterization in HEK293 cells. The data for

HEK293 cells is shown in Figure 5.4. Below (next page) pre- and post-stimulation data is

provided extensively for HeLa cells and anecdotally for CHO cells (Figure F.1).

134

Figure F.1: Demonstration of CaRQ in HeLa and CHO cells. A. Percent of HeLa cells expressing indicated construct that demonstrated a blebbing morphology before any stimulation. B. Percent of morphologically normal HeLa cells blebbing after ionomycin stimulation under the indicated conditions (transfected construct + inhibitor). C-F. Representative cells stimulated with ionomycin: YFP in HeLa (C), CaRQ in HeLa (D), pLynCaRQ in HeLa (E) and CaRQ in CHO (F). White arrows indicate blebs. Graphed data is the mean ± standard deviation of 10 experiments and at least 100 cells (A) and 3 experiments and at least 10 cells (B). Scale bars in C-F are 25 µm, and 12 µm in insets.

135

Appendix G: Mutations in Rac1 and Cdc42 to Introduce a CaM Binding Site

The bioinformatic analysis applied to RhoA that identified a potential CBP did not identify

CBP’s in the analogous regions of Rac1 and Cdc42. However, the mechanism of Ca2+-switching

RhoA thought to be responsible for CaM-RhoA(DP)’s behaviour may be applicable to Rac1 and

Cdc42 if a CBP could be engineered into them at that site. The hydrophobic anchors Val170,

Ala174 and Leu179 are conserved between the three GTPases (Val168, Ala172 and Leu177 for

Rac1 and Cdc42). CaM binding relies on key hydrophobic and electrostatic interactions (14).

Point mutations that reduce the peptide’s negative charge (such as Glu � Gln) and increase its

α–helical propensity (such as Pro � Ala) may improve the ability of the peptide to bind CaM.

A minimum number of point mutations were designed into Rac1 and Cdc42 (to create mRac1

and mCdc42, respectively) using the principles described above (Figure G.1A). The peptides

were fused with YFP to facilitate CaM-pulldown assay. Fluorescence intensities were

normalized by visual inspection to provide a rough estimation of the relative CaM binding ability

of each peptide. Surprisingly there was already some CaM-pulldown activity of the wild type

peptides, which was improved somewhat in each case (Figure G.1B). When the mutations were

introduced into the full length proteins, fused with YFP, there appeared to be increased CaM

association for the Rac1 mutant but no noticeable difference for the Cdc42 mutant, compared to

wild type GTPases. Both mutant GTPases were expressed in live cells to compare the protein

localization and cell morphology with the wild type GTPases to see if the mutations disrupted

characteristic activity. Morphologically, cells appeared to be similar to those expressing wild

type GTPases, however stimulation with ionomycin, either with or without CaM fusion, had no

apparent effect on causing lamellipodia or filopodia to retract.

These results showed that rationally designed point mutations can be used to increase CaM

binding to the carboxy-terminal tail region of Rho GTPases, and in some cases, to the full length

GTPase. However these mutations were insufficient to create a Ca2+-sensitive switch with

notable cell morphology effects. It is possible that in the absence of Rac1/Cdc42 activity, the

cell’s mechanisms for reversing their characteristic structures act on a timescale beyond the

imaging window that was used in these experiments. Further mutations may result in

sufficiently increased Ca2+-CaM affinity such that a morphology effect is noticeable, but the

extent to which these mutations disrupt native GTPase activity should be considered. These

136

experiments also provided further support for the argument of embedded peptides with Rho

proteins of known behaviour, rather than attempting to make mutations to the native sequence to

bring about some desired behaviour.

Figure G.1: Muations to Rac1 and Cdc42 to promote CaM binding. A. Carboxy-terminal regions of RhoA, Rac1 and Cdc42 with putative hydrophobic anchors shown in bold. Point mutations introduced to mutant Rac1 (mRac1) and mCdc42 are shown in red. B. CaM-pulldown assay for the indicated peptide fusions with YFP. Fluorescence intensity of the solution was normalized by visual inspection prior to pulldown to provide an estimate of relative Ca2+-CaM affinity.

137

Appendix H: Ca2+ Control Designs for Rac1 and Cdc42 with PBD

Incorporating an inhibitor into the Ca2+ control paradigm for Rho GTPases has the potential to

increase the size of the design space for Ca2+-controlled GTPases. PBD from PAK binds both

Rac1 and Cdc42 (96). A method of controlling the presentation of PBD to Rac1 or Cdc42 may

be sufficient to control the activation of the GTPase without the need for extensive changes to its

sequence or structure. Two broad categories of control were considered: intermolecular control

strategies and intramolecular control strategies.

Intermolecluar control strategies focused on co-expressing Rac1 or Cdc42 and PBD where either

the GTPase or the inhibitor were modified to be Ca2+ sensitive (Figure H.1). For example, by

flanking PBD with CaM and MLCKp, presumably in a low Ca2+ environment Rac1 or Cdc42

and PBD would associate, disrupting Rac1 or Cdc42-dependent morphology. On Ca2+ loading,

CaM-MLCKp association may cause a sufficient conformational change in the PBD fusion to

disrupt the GTPase-PBD interaction and allow the GTPase to interact with its native effectors.

Several combinations and arrangements were created. The GTPase was constitutively localized

to the PM so that translocation could serve as a readout for changes in interaction.

Figure H.1: Fusion domain layout of intermolecular control strategies. In the top portion of the figure, layouts are shown where unmodified Rac1 was combined with modified PBD constructs. In the bottom portion layouts are shown for the converse strategy. Rac1 is shown, but the designs were also tried for Cdc42.

138

PBD fusions with Rac1 or Cdc42 were also tried as an intramolecular Ca2+-control strategy

(Figure H.2). The idea supporting this strategy was the difference in flexibility of the CBP when

bound to Ca2+-CaM or unbound. In the absence of Ca2+-CaM, many CBP’s are unstructured (14,

115), meaning that they are extremely flexible with many degrees of freedom. In the presence of

Ca2+-CaM, CBP’s associate with Ca2+-CaM in known structures suggesting that much of this

freedom is lost. Therefore in the absence of Ca2+ the PBD-CaM-MLCKp-YFP-Rac1 fusion may

be sufficiently flexible for PBD to bind Rac1 and in the presence of Ca2+, sufficiently inflexible

to prevent PBD from associating with Rac1.

Figure H.2: Fusion domain layout and cartoon for intramolecular control strategies. One potential domain layout is given, and below, a cartoon showing how the fusion may behave on Ca2+ loading. Domain colours in the cartoon match those in the domain layout diagram.

Experiments performed with both intermolecular and intramolecular Ca2+-control strategies were

generally unsuccessful in inducing Ca2+-dependent morphology changes in COS7 cells.

However, these experiments were performed before many of the more sophisticated Ca2+-

mobilizing modules (such as LOVS1K/Orai1) were available and so Ca2+ could only be

delivered by ionomycin which does not allow much control over the spatiotemporal delivery

characteristics. Modifications of these strategies using the principles outlined in section 8.2 may

rescue their intended function. However, given the development of RACer and the CaM-Cdc42

chimera, further pursuing these strategies is unwarranted at present.

139

Appendix I: Area Thresholding Analysis Applied to Blebbing Cells

The area thresholdhing analysis that was applied to RACer and control cells in Chapter 6 was

used as a way to quantify lamellipodia formation (5), and not for the purposes of distinguishing

lamellipodia from blebbing. In wild type morphology cells that developed the blebbing

morphology the cell area, as calculated using the same method in Chapter 6, also increased

(Figure I.1). Here, the cells shown in Figure 5.7D were analyzed using that method. The time

course in the graph below has three phases: control, bleb development and dynamic blebbing. In

the first phase (0-5 minutes), area changed by less than .05-fold. In the second phase (5-15

minutes) the area increased nearly 0.15-fold while blebs developed. In the third phase of

dynamic blebbing (15-25 minutes) the area change oscillates with peak to trough amplitude

variation of approximately 0.5-fold. This oscillation appeared to be common in other blebbing

cells that were analyzed by this method, although “natural” fluctuations in area (such as that seen

during the control period) may also contribute to this pattern. If this pattern were a general

feature of blebbing cells then it may form an alternative to the method of bleb identification used

here, which was visual inspection based on the shape, fluorescence and time scale of the

protrusions.

Figure I.1: Area thresholding of HEK293 cells co-expressing CaRQ and nAChR-α4 stimulated by ACh. The cells analyzed (shown in Figure 5.7D) were thresholded in the same manner as was done for RACer cells. Cells were stimulated with ionomycin after a 5 minute control period, as indicated. In these cells, blebbing began at approximately 10 minutes and continued for the duration.

engineering synthetic control over rho gtpases using ca ... · engineering synthetic control over...

Documents