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THE MOLECULAR PATHOGENESIS OF NOONAN SYNDROME-ASSOCIATED RAF1 MUTATIONS by Xue Wu A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Department of Medical Biophysics University of Toronto © Copyright by Xue Wu 2013

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Page 1: THE MOLECULAR PATHOGENESIS OF NOONAN SYNDROME-ASSOCIATED ... · 1.2 Noonan syndrome and the RASopathies 13 1.2.1 Noonan syndrome 13 1.2.2 RASopathies: disorders clinically related

THE MOLECULAR PATHOGENESIS OF NOONAN

SYNDROME-ASSOCIATED RAF1 MUTATIONS

by

Xue Wu

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Department of Medical Biophysics

University of Toronto

© Copyright by Xue Wu 2013

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THE MOLECULAR PATHOGENESIS OF NOONAN SYNDROME-

ASSOCIATED RAF1 MUTATIONS

Xue Wu

Doctor of Philosophy

Department of Medical Biophysics

University of Toronto

2013

ABSTRACT

Noonan syndrome (NS) is one of several autosomal dominant “RASopathies” caused by

mutations in components of the RAS-RAF-MEK-ERK MAPK pathway. Germ line mutations in

RAF1 (encoding the serine-threonine kinase for MEK1/2) account for ~3-5% of NS, and unlike

other NS alleles, RAF1 mutations that confer increased kinase activity are highly associated with

hypertrophic cardiomyopathy (HCM). Notably, some NS-associated RAF1 mutations show

normal or decreased kinase activity. To explore the pathogenesis of such mutations, I generated

“knock-in” mice that express kinase-activating (L613V) or -impaired (D486N) Raf1 mutants,

respectively. Knock-in mice expressing the kinase-activated allele Raf1L613V developed typical

NS features (short stature, facial dysmorphia, haematological abnormalities), as well as HCM.

As expected, agonist-evoked Mek/Erk activation was enhanced in multiple cell types expressing

Raf1L613V. Moreover, postnatal Mek inhibition normalized the growth, facial, and cardiac defects

in L613V/+ mice, showing that enhanced Mek/Erk activation by Raf1 mutant is critical for

evoking NS phenotypes. D486N/+ female mice exhibited a mild growth defect. Male and female

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D486N/D486N mice developed concentric cardiac hypertrophy and incompletely penetrant, but

severe, growth defects. Remarkably, Mek/Erk activation was enhanced in Raf1D486N-expressing

cells compared with controls. In both mouse and human cells, RAF1D486N, as well as other

kinase-impaired RAF1 mutants, show increased heterodimerization with BRAF, which is

necessary and sufficient to promote increased MEK/ERK activation. Furthermore, kinase-

activating RAF1 mutants also require heterodimerization to enhance MEK/ERK activation. Our

results suggest that increased heterodimerization ability is the common pathogenic mechanism

for NS-associated RAF1 mutations.

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ACKNOWLEDGMENTS

Foremost, I would like to thank my supervisor Dr. Benjamin Neel for all your guidance,

continuous support and encouragement during my Ph.D study. You have been a tremendous

mentor for me with your immense knowledge, inspiration, enthusiasm, and criticism. I also thank

you for the critical reading of this thesis.

I am very grateful to Dr. Toshiyuki Araki as a co-supervisor for my research project.

Thank you for sharing your knowledge, good discussions and friendship.

I thank the Department of Medical Biophysics at University of Toronto for giving me the

opportunity to study here. I greatly appreciate the insightful discussions and comments from all

my supervisory committee members, Dr. Peter Backx, Dr. Vuk Stambolic and Dr. Benjamin

Alman. Your advice on both research, as well as on my career, have been invaluable.

This research project would not have been possible without the help of many people.

Special thanks to Dr. Peter Backx and Dr. Jeremy Simpson as important collaborators on the

cardiology studies of my mouse model. Dr. Jeremy Simpson performed all the echocardiography,

invasive hemodynamic analysis and transverse aortic constriction experiments. Our collaboration

has continued after he established his own lab at the University of Guelph. Thanks to Dr.

Kyoung-Han Kim for preparing the neonatal cardiomyocytes for me, and for your patience in

teaching me this technique. I also thank Dr. Tara Paton at The Centre for Applied Genomics

(TCAG) at SickKids for helping me with the mouse genotyping array and Dr. Pingzhao Hu in the

same facility for providing statistical analysis support. I would also like to thank Dr. Jason

Moffat (University of Toronto) for providing lentiviral shRNA vectors and Dr. Bruce Gelb (Mt

Sinai Hospital, NY) for provding the human RAF1 cDNA construct used in my research.

I am thankful to all of the past and present members in the Neel lab, Jocelyn Stewart, Rob

Karish, Shengqing Gu, Xiannan Wang, Peter Bayliss, Angel Sing, Cathy Iorio, Gordon Chan,

Ziqiang Yang, Richard Marcotte, Dong Hu, Robert Banh, Yang Xu, Lingyan Jiang, Anderson

Chang, Erica Tiberia, Kwan Ho Tang, Paulina Cybulska, Richard Chan and many others. The

group has been a source of friendships, as well as good advice and collaboration. All of them

made my study in this lab a cherished memory. Thank you Shengqing Gu for your help of

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analyzing the hematological defects in my mice, and for your biostatistics support. I am honored

to have had the opportunity to supervise two talented and hard-working summer students, Jenny

Hong and Connie Yin. I am grateful for all their help in many aspects of my project. I also would

like to thank our previous lab member, Dr. Nirusha Thavarajah, for synthesizing the MEK

inhibitor. I thank Peter Bayliss, and Angel Sing for their technical support, and our lab manager,

Cathy Iorio, for making the lab organized and functional.

The Frederick Banting and Charles Best Canada Graduate Scholarship from Canadian

Institutes of Health Research (CIHR) is greatly appreciated. This research also was supported by

grants from the CIHR, the National Institutes of Health (NIH) and the Heart and Stroke

Foundation of Ontario (H&SFO).

Finally, I dedicate this thesis to my parents, Feng Yu and Shouzhi Wu, my husband,

Kevin Hu, and my beloved daughter, Claire Hu, for their constant support and unconditional love.

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TABLE OF CONTENTS

ABSTRACT II

ACKNOWLEDGMENTS IV

TABLE OF CONTENTS VI

LIST OF FIGURES X

LIST OF TABLES XIV

LIST OF ABBREVIATIONS XV

CHAPTER 1 INTRODUCTION 1

1.1 The RAS-RAF-MEK-ERK MAPK pathway 2

1.1.1 MAPK pathways 2

1.1.2 The RAS-RAF-MEK-ERK MAPK pathway 5

1.2 Noonan syndrome and the RASopathies 13

1.2.1 Noonan syndrome 13

1.2.2 RASopathies: disorders clinically related to Noonan syndrome 21

1.2.3 Noonan syndrome mouse models 25

1.3 Function and regulation of RAF1 26

1.3.1 Structure of RAF family kinases 27

1.3.2 Regulation of RAF1 29

1.3.3 MEK-independent functions of RAF1 39

1.4 Hypertrophic cardiomyopathy and the RAS/ERK pathway 41

1.4.1 Cardiac hypertrophy and hypertrophic cardiomyopathy 41

1.4.2 Signaling pathways involved in cardiac hypertrophy 45

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1.5 Rationale and hypothesis 50

CHAPTER 2 MATERIALS AND METHODS 52

2.1 Mice 53

2.1.1 Generation of Raf1L613V knock-in mice 53

2.1.2 Generation of Raf1D486N knock-in mice 55

2.2 Cell culture 57

2.2.1 Mouse embryonic stem (ES) cells 57

2.2.2 Mouse embryonic fibroblasts (MEFs) 57

2.2.3 Neonatal cardiomyocytes and cardiac fibroblasts 58

2.2.4 Flp-In T-REx 293 cell lines 58

2.3 Generation of Flp-In T-REx 293 expression cell lines 59

2.4 Inducible RAF1/BRAF heterodimerization 59

2.5 Lentivirus production and transduction 59

2.6 Retrovirus production and transduction 60

2.7 Biochemical analysis 60

2.8 Body size analysis and morphometry 61

2.9 Histology and immunohistochemistry 61

2.10 BrdU incorporation assays 62

2.11 Hematopoietic analysis 62

2.12 Echocardiographic and hemodynamic measurements 62

2.13 Transverse aortic constriction (TAC) 63

2.14 MEK inhibitor treatment 63

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2.15 Quantitative real time RT-PCR 64

2.16 Statistics 64

CHAPTER 3 MEK-ERK PATHWAY MODULATION AMELIORATES DISEASE

PHENOTYPES IN A MOUSE MODEL OF NOONAN SYNDROME ASSOCIATED

WITH THE RAF1L613V MUTATION 65

3.1 Abstract 66

3.2 Background 66

3.3 Results 70

3.3.1 Generation of L613V/+ mice 70

3.3.2 L613V/+ mice show multiple NS phenotypes 72

3.3.3 L613V/+ mice show cardiac hypertrophy with chamber dilatation 72

3.3.4 Enhanced hypertrophic response and functional decompensation in L613V/+ hearts

following pressure overload

81

3.3.5 The Raf1L613V mutant increases Mek and Erk activation in response to multiple stimuli

85

3.3.6 MEK inhibitor treatment normalizes NS phenotypes in L613V/+ mice 95

3.4 Discussion 102

CHAPTER 4 INCREASED BRAF HETERODIMERIZATION IS THE COMMON

PATHOGENIC MECHANISM FOR NOONAN SYNDROME-ASSOCIATED RAF1

MUTANTS 106

4.1 Abstract 107

4.2 Background 107

4.3 Results 109

4.3.1 Generation of Raf1D486N mice 109

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4.3.2 Phenotypes of D486N/+ and D486N/D486N mice 112

4.3.3 Raf1D486N expression increases Mek/Erk activation in response to multiple stimuli 113

4.3.4 Quantitative differences in effects of NS-associated Raf1 D486N and L613V mutants

on Mek/Erk activation 117

4.3.5 Kinase-impaired Raf1 mutants enhance Mek/Erk activation by promoting

heterodimerization with Braf

124

4.3.6 Heterodimerization with BRAF is required for RAF1D486N to enhance MEK/ERK

activation 127

4.4 Discussion 138

CHAPTER 5 SUMMARY AND FUTURE DIRECTIONS 143

5.1 Summary and Key Findings 144

5.2 Future Directions 145

5.2.1 Optimizing therapy for NS-associated HCM 145

5.2.2 Downstream target(s) of the RAS/ERK signaling in NS and HCM 150

5.2.3 Cell(s)-of-origin for Raf1 mutant-induced HCM 153

5.2.4 The roles of specific Erk isoforms in Raf1-induced HCM 158

5.2.5 Genetic modifiers in D486N/D486N mice 163

5.3 Concluding Remarks 169

References 170

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LIST OF FIGURES

Figure 1-1. Schematic overview of MAPK pathways. ................................................................... 3 

Figure 1-2. Schematic representation of the structure of RAS-RAF-MEK-ERK MAPK pathway.

......................................................................................................................................................... 6 

Figure 1-3. Downstream targets of ERK signaling. ...................................................................... 11 

Figure 1-4. Facial dysmorphism in Noonan syndrome. ................................................................ 15 

Figure 1-5. RASopathies and the RAS-RAF-MEK-ERK MAPK pathway. ................................ 18 

Figure 1-6. RAF1 domain structure and location of residues altered in NS. ................................ 20 

Figure 1-7. Common structure of RAF proteins. .......................................................................... 28 

Figure 1-8. Overview of RAF1 activation/deactivation. .............................................................. 31 

Figure 1-9. Regulatory phosphorylation sites of RAF proteins. ................................................... 32 

Figure 1-10. Allosteric mechanism for activation of the kinase domain of RAF1. ...................... 36 

Figure 1-11. Scaffolding proteins in RAF-MEK-ERK signaling. ................................................ 37 

Figure 1-12. MEK-independent RAF1 signaling pathways. ........................................................ 40 

Figure 1-13. Types of cardiac hypertrophy. .................................................................................. 43 

Figure 1-14. Signaling pathways involved in cardiac hypertrophy. ............................................. 47 

Figure 3-1.Generation of inducible Raf1L613V knock-in mice. ..................................................... 71 

Figure 3-2. Short stature in L613V/+ mice. .................................................................................. 73 

Figure 3-3. L613V/+ mice have facial dysmorphia. ..................................................................... 74 

Figure 3-4. L613V/+ mice have hematological defects. ............................................................... 75 

Figure 3-5. L613V/+ mice show cardiac hypertrophy with normal cardiomyocyte proliferation

and valve development. ................................................................................................................ 77 

Figure 3-6. L613V/+ mice show cardiac hypertrophy with chamber dilatation. .......................... 79 

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Figure 3-7. L613V/+ mice show a shift from alpha-Mhc to beta-Mhc expression in hearts. ...... 83 

Figure 3-8. Abnormal response of L613V/+ mice to pressure overload. ..................................... 84 

Figure 3-9. Severe perivascular fibrosis and infarct in L613V/+ mice after TAC. ...................... 86 

Figure 3-10. Echocardiographic parameters in WT and L613V/+ mice following pressure

overload. ........................................................................................................................................ 87 

Figure 3-11. Hemodynamic parameters in WT and L613V/+ mice following pressure overload.

....................................................................................................................................................... 88 

Figure 3-12. Raf1L613V mutant causes increased Mek and Erk activation in multiple cell types. 90 

Figure 3-13. Raf1L613V mutant increases Mek and Erk activation in cardiomyocytes. ................. 91 

Figure 3-14. Raf1L613V mutant increases Mek and Erk activation in cardiac fibroblasts. ............ 92 

Figure 3-15. Enhanced Mek and Erk activation in L613V/+ hearts after pressure overload. ...... 93 

Figure 3-16. Other signaling pathways are unaffected in neonatal cardiac myocytes and

fibroblasts. ..................................................................................................................................... 94 

Figure 3-17. MEK inhibitor treatment rescues growth defect and cardiac hypertrophy in L613V/+

mice. .............................................................................................................................................. 97 

Figure 3-18. Normalized cardiac morphology and function after MEK inhibitor treatment. ....... 98 

Figure 3-19. MEK inhibitor treatment normalizes cardiac function in L613V/+ mice. ............... 99 

Figure 3-20. Early post-natal MEK inhibitor treatment rescues facial dysmorphia in L613V/+

mice. ............................................................................................................................................ 101 

Figure 4-1. Generation of Raf1D486N knock-in mice. .................................................................. 111 

Figure 4-2. Phenotypes of D486N/+ and D486N/D486N mice. ................................................. 115 

Figure 4-3. Additional phenotyping of D486N/+ and D486N/D486N mice. ............................. 116 

Figure 4-4. Raf1D486N mutant increases Mek and Erk activation. ............................................... 119 

Figure 4-5. Mek/Erk activation in WT and D486N/D486N neonatal cardiomyocytes. ............. 120 

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Figure 4-6. Mek/Erk activation in WT and D486N/D486N neonatal cardiac fibroblasts. ......... 121 

Figure 4-7. NS-associated Raf1 mutants differentially activate Mek/Erk. ................................. 123 

Figure 4-8. Severity of cardiac phenotype in D486N/D486N and L613V/+ mice correlates with

Mek/Erk activation. ..................................................................................................................... 126 

Figure 4-9. RAF1D486N forms more heterodimers with BRAF. .................................................. 129 

Figure 4-10. Kinase-impaired RAF1 mutants associated with NS enhance MEK/ERK activation

and form more RAF1/BRAF heterodimers. ................................................................................ 130 

Figure 4-11. BRAF is required for RAF1D486N to enhance MEK/ERK activation. .................... 131 

Figure 4-12. Heterodimerization with BRAF is required for RAF1D486N mutant to enhance

MEK/ERK activation. ................................................................................................................. 135 

Figure 4-13. Induced RAF1/BRAF heterodimerization restores activity of RAF1R401H/D486N

mutant to enhance MEK/ERK activation. .................................................................................. 136 

Figure 4-14. RAF1L613V mutant enhances MEK/ERK activation via RAF1/BRAF heterodimer

formation. .................................................................................................................................... 137 

Figure 5-1. Lower dose MEK inhibitor treatment rescues growth defect, but not cardiac

hypertrophy and chamber dilatation in L613V/+ mice. .............................................................. 148 

Figure 5-2. Lower dose MEK inhibitor treatment partially normalizes cardiac function in

L613V/+ mice. ............................................................................................................................ 149 

Figure 5-3. Increased pS6 (S235/236) in cardiomyocytes and cardiac fibroblasts from L613V/+

mice. ............................................................................................................................................ 151 

Figure 5-4. Rapamycin treatment does not rescue growth defect and cardiac hypertrophy in

L613V/+ mice. ............................................................................................................................ 152 

Figure 5-5. Cardiomycyte-specific expression of Raf1L613V does not cause significant cardiac

hypertrophy. ................................................................................................................................ 154 

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Figure 5-6. Cardiomycyte-specific expression of Raf1L613V causes mild increase in cardiac

function. ...................................................................................................................................... 156 

Figure 5-7. Cardiomycyte-specific expression of Raf1L613V causes mild increase in cardiac

contractility. ................................................................................................................................ 157 

Figure 5-8. Reducing Erk1 levels in L613V/+ mice does not rescue the cardiac hypertrophy and

chamber dilatation. ...................................................................................................................... 159 

Figure 5-9. Reducing Erk1 levels in L613V/+ mice does not rescue the cardiac function. ....... 160 

Figure 5-10. Reducing Erk1 levels in L613V/+ mice rescues the cardiac contractility. ............ 161 

Figure 5-11. Reducing Erk2 levels in L613V/+ mice does not rescue the cardiac hypertrophy. 162 

Figure 5-12. Distributions of the body weight data for all the 56 samples. ................................ 164 

Figure 5-13. Genome-wide LOD score plot of the quantitative trait. ......................................... 165 

Figure 5-14. Genome-wide LOD score plot of the binary trait. ................................................. 166 

Figure 5-15. LOD score plot of the binary trait on Chromosome 8. .......................................... 167 

Figure 5-16. Boxplots of body weight for mice with different genotypes. ................................. 168 

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LIST OF TABLES

Table 1-1. Clinical features of Noonan syndrome. ....................................................................... 14 

Table 2-1. PCR primers for generating the Raf1L613V mice. ......................................................... 54 

Table 2-2. PCR primers for generating the Raf1D486N mice.......................................................... 56 

Table 3-1. Echocardiographic parameters in WT and L613V/+ mice. ......................................... 80 

Table 3-2. Additional hemodynamic parameters of hearts from 4 month-old mice. .................... 82 

Table 4-1. Comparison of cardiac phenotypes in D486N/D486N and L613V/+ mice. ............. 122 

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LIST OF ABBREVIATIONS

ADHD attention deficit-hyperactive disorder

ALL acute lymphoblastic leukemia

AML acute myeloid leukemia

AMPK adenosine monophosphate-activated protein kinase

AN anal-nasal

Ang-II angiotensin-II

ANP atrial natriuretic peptide

AP1 activating protein-1

AS aortic stenosis

ASK1 apoptosis signal-regulating kinase 1

ATF1 activating transcription factor 1

BNP B-type natriuretic peptide

CaMK calmodulin-dependent kinase

CDK cyclin-dependent kinase

CFCS cardio-facio-cutaneous syndrome

CHD congenital heart defects

CK2 casein kinase 2

CNK connector enhancer of KSR

CO cardiac output

COT Cancer Osaka Thyroid oncogene

CR conserved region

CRD cysteine-rich domain

CREB cAMP response element binding protein

CS Costello syndrome

CSK C-terminal SRC kinase

DAG diacylglycerol

DMEM Dulbecco’s modified Eagle’s medium

DUSP dual specificity phosphatase

ECM extracellular matrix

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EDC epidermal differentiation cluster

EDV end-diastolic volume

EF ejection fraction

elF-4E eukaryotic initiation factor-4E

ELK-1 Ets-like gene 1

ERK extracellular signal-regulated kinase

ES embryonic stem

ESV end-systolic volume

ET-1 endothelin-1

FACS fluorescence activated cell sorting

FBS fetal bovine serum

FIAU 1-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)-5 iodouracil

FS fractional shortening

GAP GTPase activating protein

GATA4 GATA binding protein 4

GC-A guanylyl cyclase-A

GEF guanine nucleotide exchange factor

GH growth hormone

GHD growth hormone deficiency

G-loop glycine-rich loop

Gp130 glycoprotein-130

GPCR G-protein coupled receptor

GRB2 growth factor receptor-bound protein 2

GSK3β glycogen synthase kinase-3β

HCM hypertrophic cardiomyopathy

HDAC histone deacetylase

IEG immediate-early gene

IGF-I insulin-like growth factor I

IKK inhibitor of NF-κB kinase

IL-6 interleukin-6

IP intraperitoneally

IQGAP IQ motif-containing GTPase-activating protein

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IκB inhibitor of NF-κB

JMML juvenile myelomonocytic leukemia

JNK c-Jun N-terminal kinase

KSR1 kinase suppressor of Ras 1

LAH loose anagen hair

LAMP2 lysosome-associated membrane protein-2-α-galactosidase

LD linkage disequilibrium

LEOPARD Lentigines, ECG conduction abnormalities, Ocular hypertelorism,

Pulmonic stenosis, Abnormal genitalia, Retardation of growth, and

sensorineural Deafness

LPA lysophosphatidic acid

LS LEOPARD syndrome

LV left ventricle

LVH left ventricular hypertrophy

LVIDd left ventricular internal end-diastolic dimention

LVIDs left ventricular internal end-systolic dimention

LVPWd left ventricular diastolic posterior wall thickness

MAP mean arterial pressure

MAPK mitogen-activated protein kinase

MAPKK mitogen-activated protein kinase kinase

MAPKKK mitogen-activated protein kinase kinase kinase

MEF mouse embryonic fibroblast

MEF2 myocyte enhancer factor 2

MEK1 MAPK/ERK kinase 1

MI myocardial infarction

MKP MAP kinase phosphatase

MLP muscle LIM-domain protein

MNK MAPK-interacting kinase

MORG1 MAPK organizer 1

MHC myosin heavy chain

MP1 MEK partner-1

MPNST malignant peripheral nerve sheath tumors

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MSK mitogen- and stress-activated protein kinase

MST2 mammalian STE20-like protein kinase 2

MSV murine sarcoma virus

mTOR mammalian target of rapamycin

NEAA non-essential amino acid

NF1 neurofibromatosis type 1

NFAT nuclear factor of activated T cells

NFLS NF1-like syndrome

NF-κB nuclear factor-κB

NIK NF-κB-inducing kinase

NS Noonan syndrome

NS/LAH Noonan-like syndrome with loose anagen hair

OPG optic pathway gliomas

PDK phosphoinositide-dependent kinase

PH plekstrin homology

PHB prohibitin

PI3K phosphoinositide-3 kinase

PKC protein kinase C

PKCθ protein kinase C theta

PLA2 phospholipase A2

PLC phospholipase C

PP2A protein phosphatase 2A

PP5 protein phosphatase 5

PS pulmonic stenosis

PTB phosphotyrosine binding

PTK protein-tyrosine kinase

PTP protein-tyrosine phosphatase

RAF rapid accelerated fibrosarcoma

RASSF1A Ras association domain-containing protein 1A

RBD Ras binding domain

REM Ras exchanger motif

RKIP Raf kinase inhibitor protein

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ROS reactive oxygen species

RSK ribosomal S6 kinase

RTK receptor tyrosine kinase

RV right ventricle

SAP1 SRF accessory protein 1

SAPK stress-activated protein kinase

SEF similar expression to FGF

SERCA2a sarcoplasmic reticulum Ca2+-ATPase

SH2 Src homology 2

SH3 Src homology 3

SHPS-1 Shp Substrate-1

SOS son of sevenless

SPRED1 Sprouty-related EVH1 domain-containing protein 1

SRE serum response element

SRF serum response factor

SPRY SPROUTY

SV stroke volume

TAC transverse aortic constriction

TAK TGFβ-activated kinase

TCF ternary complex factor

TGFβ transforming growth factor-β

TNF-α tumor necrosis factor-α

TPL2 tumor progression locus 2

UBF upstream binding factor

VDAC voltage-dependent anion channel

WGA wheat germ agglutinin

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Chapter 1

Introduction

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1.1 The RAS-RAF-MEK-ERK MAPK pathway

1.1.1 MAPK pathways

Mitogen-activated protein kinase (MAPK) pathways are evolutionarily conserved kinase

modules that link extracellular signals to the machinery that controls fundamental cellular

processes, such as proliferation, differentiation, migration and apoptosis (1). MAPK pathways

are comprised of a three-tier kinase module in which a MAPK is activated upon phosphorylation

by a mitogen-activated protein kinase kinase (MAPKK) which, in turn, is activated when

phosphorylated by a mitogen-activated protein kinase kinase kinase (MAPKKK) (Figure 1-1).

To date, three main families of MAPKs have been characterized in mammals, which are

grouped by their structures and functions: extracellular signal-regulated kinase (ERK), c-Jun N-

terminal kinase (JNK, also known as stress-activated protein kinase or SAPK)1/2/3 and the p38

isoforms α/β/γ (ERK6)/δ (2-5). Members of the ERK family can be divided further into four

subgroups: the ERK1/2 group, comprising classic MAPKs that consist primarily of a kinase

domain, and the three large MAPK groups, ERK3/4, ERK5 and ERK7/8, containing enzymes

that consist of both a kinase domain and a C-terminal domain ranging in size from 60 kDa to

greater than 100 kDa (6-9). The C-terminal regions function as protein interaction domains that

regulate kinase localization (10), activation (9, 11), and transcriptional activity (7).

All MAPKs contain a TXY (Thr-X-Tyr) motif within their activation loops. The

phosphorylation of both the threonine and the tyrosine within the activation loop is necessary and

sufficient for their activation. The majority of ERK family members contain a TEY (Thr-Glu-Tyr)

activation motif (12), except for ERK3/4, which possesses an SEG (Ser-Glu-Gly) sequence (6).

The p38 family has a TGY (Thr-Gly-Tyr) activation motif (13), whereas JNK family members

contain TPY (Thr-Pro-Tyr) in their activation loop (14).

The ERK pathway is the best studied mammalian MAPK pathway and is activated by

numerous extracellular signals (Figure 1-1). In the ERK1/2 MAPK module, ERK1/2 are

activated upon phosphorylation by MAPK/ERK kinase1/2 (MEK1/2), which is itself activated

when phosphorylated by Rapid Accelerated Fibrosarcoma (RAF) family proteins. The details of

this signaling cascade will be discussed later. Although RAF isoforms are the primary

MAPKKKs in the ERK1/2 module, MAP3K8, also called TPL2 (tumor progression locus 2) or

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Figure 1-1. Schematic overview of MAPK pathways.

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COT (Cancer Osaka Thyroid oncogene) also can act as a MAPKKK that regulates the ERK1/2,

ERK5, JNK and p38 pathways in a cell type- and stimulus-specific manner (15). MEKK1 (16)

and MOS (17) are two additional ERK1/2 MAPKKKs utilized in more restricted cell type- and

stimulation-specific situations. The MAPKK for ERK5 is MEK5, which is activated upon

phosphorylation by MEKK2/3 or TPL2 (15, 18). The MAPKKs and MAPKKKs for ERK3/4 and

ERK7/8 remain unknown.

Many MAPK pathways participate in stress signaling. Stress-activated MAPK cascades

contain a large number of MAPKKKs, probably because stresses come in many forms (Figure

1-1). JNK family kinases can be activated by cytokines, UV radiation, oxidative stress, growth

factor deprivation and DNA-damaging agents (19). JNK activation requires dual phosphorylation

on the TPY motif, which is catalyzed by MEK4 or MEK7. MEK4/7 are themselves

phosphorylated and activated by a wide range of MAPKKKs, including MEKK1–4, DLK,

MLK2/3, TPL-2, TAO1/2, TAK1 and ASK1/2. Like ERKs, JNKs translocate from the cytoplasm

to the nucleus following activation, and phosphorylate transcription factors (e.g., c-JUN, ATF-2

and STAT3) (20). Active JNK also functions in the cytoplasm, although relatively little is known

about cytoplasmic JNK substrates.

p38 MAPKs are strongly activated by physical and chemical stresses, such as high

osmolarity, oxidative stress, hypoxia and UV irradiation, and pro-inflammatory cytokines, such

as endotoxin, TNF-α, and IL-1 (21, 22). In addition to its role in stress responses, the p38

pathway also helps regulate apoptosis, cell cycle progression, growth and differentiation, which

reflects its activation by a broad range of extracellular stimuli, including growth factors (e.g.,

FGF, IGF-I, PDGF and nerve growth factor) and hormones. The conserved TGY

phosphorylation motif of p38 isoforms is phosphorylated by MEK3 or MEK6, which themselves

are activated by various MAPKKKs shared with JNK, including TAK1, ASK1/2, DLK,

MEKK4, TAO1/2/3 and MLK2/3. In some instances, p38 can be activated by MEK4 (23). The

downstream targets of the active p38 MAPKs include a wide array of cytoplasmic (e.g., Caspase-

3/6, Caspase 8, and MNK1/2) and nuclear (e.g., ATF2, p53, ELK1 and STAT1) targets (22).

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1.1.2 The RAS-RAF-MEK-ERK MAPK pathway

The RAS-RAF-MEK-ERK MAPK signal transduction cascade (hereafter, the RAS/ERK

pathway) functions in many cellular processes, including proliferation, differentiation, survival,

cell adhesion, migration and metabolism (Figure 1-2) (24). The pathway is initiated by the

activation of one of three small guanosine triphosphatases (GTPases) KRAS, NRAS or HRAS,

which are stimulated by guanine-nucleotide exchange factors (GEFs), such as son of sevenless

(SOS) (25). Active RAS proteins interact with a wide range of effectors and stimulate

downstream signaling components, including RAF kinases, phosphoinositide-3 kinase (PI3K),

RAL guanine nucleotide dissociation stimulator (RALGDS), MEKK1, RAS

interaction/interference 1 (RIN1), ALL-1 fusion partner in chromosome 6 (AF-6), phospholipase

C epsilon, and novel RAS effector (NORE)/mammalian STE20-like protein kinase (MST) (26,

27). In the RAS/ERK pathway, RAS recruits RAF proteins to the cell membrane, where they are

activated and subsequently form complexes with MEK1/2 and ERK1/2, aided by scaffolds, such

as KSR. Activated RAF proteins phosphorylate MEK1/2, which in turn phosphorylate ERK1/2.

ERKs phosphorylate cytosolic substrates and also translocate to the nucleus to stimulate diverse

gene expression programs by phosphorylating several transcription factors (26, 28).

Deregulation of the RAS/ERK pathway is associated with various pathologies, most

notably cancer (29, 30), but also developmental abnormalities (31, 32). Oncogenic mutations in

human KRAS occur in about 90% of pancreatic, 40% of colorectal, and 30% of biliary cancers,

whereas NRAS mutations occur in approximately 15-20% of melanomas. Overall, RAS genes are

activated in about 30% of all human cancers (26, 33, 34). Activating mutations in BRAF occur in

approximately 7% of all human cancers, including 27-70% of malignant melanomas, 36-53% of

papillary thyroid cancers, 30% of ovarian cancers, and 5-22% of colorectal cancers (35, 36). The

RASopathies are a group of developmental syndromes caused by germline mutations in genes

that alter the RAS/ERK pathway (31, 32), and will be discussed later in detail.

The conversion of inactive guanine diphosphate (GDP)-bound form to active guanine

triphosphate (GTP)-bound form of RAS is promoted by the action of several receptor tyrosine

kinases (RTKs), including those of the epidermal growth factor receptor (EGFR) family, the

insulin-like growth factor receptor (IGFR), the vascular endothelial growth factor receptor

(VEGFR) family, and many others (37). Membrane-spanning cell surface RTKs are endowed

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Figure 1-2. Schematic representation of the structure of RAS-RAF-MEK-ERK MAPK

pathway.

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with intrinsic tyrosine kinase activity. All RTKs contain an extracellular ligand-binding domain,

a single hydrophobic transmembrane helix, and a cytoplasmic region that contains a conserved

protein-tyrosine-kinase domain (TKD) plus additional C-terminal and juxamembrane regulatory

regions that are subject to autophosphorylation and phosphorylation by other kinases. Most

RTKs are monomers at the cell membrane, with ligand binding or ectopic over-expression

resulting in receptor dimerization and tyrosine “autophosphorylation” in trans, although some

RTKs exist as oligomers; e.g., the insulin receptor and IGFR, which form disulfide-linked dimers

in the absence of activating ligand. Whether the “inactive” state is monomeric or oligomeric,

activation of the RTKs still requires the bound ligand to stabilize the individual receptor

molecules in an “active” dimer or oligomer. In some RTKs, such as the insulin receptor, KIT,

and TIE2, trans-phosphorylation of tyrosines within the activation loop, the juxtamembrane

segment, and/or the C-terminal region disrupts their cis-autoinhibitory interaction with TKD, and

promotes receptor activation (38-40). However, the EGFR family and RET are exceptions. Their

activation does not require trans-phosphorylation of their activation loop or elsewhere. Instead,

dimerization of these receptors promotes conformational changes and thereby allosteric

activation of their TKDs (41, 42).

Following RTK activation, additional tyrosines are autophosphorylated in other parts of

the receptor cytoplasmic region. The resulting phospho-tyrosines provide a mechanism for the

recognition and assembly of signaling complexes, functioning as binding sites for a variety of

cytoplasmic signaling molecules containing Src homology 2 (SH2) domain (43) and/or

phosphotyrosine binding (PTB) (44) domains. One such signaling protein is growth factor

receptor-bound protein 2 (GRB2). GRB2, a cytosolic adaptor, contains a central SH2 domain,

flanked by two Src homology 3 (SH3) domains, which allow constitutive association with the

proline-rich regions of SOS (45). For example, phospho-tyrosine 1068 of the activated EGFR is

a binding site for the SH2 domain of GRB2 (46), either directly or through the assistance of

another SH2 adaptor, SHC (47). Activated EGFR phosphorylates SHC on Tyr317, which

promotes the interaction between SHC and GRB2 (48). The recruitment of GRB2 from the

cytoplasm to the plasma membrane brings SOS near membrane-bound RAS, where SOS

enhances GDP release and GTP binding to RAS, converting this GTPase into its active

conformation. It is well accepted that the protein-tyrosine phosphatase SHP2 (encoded by

PTPN11) has a positive effect on RAS/ERK activation, but the mechanisms involved have

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remained elusive (49). Several mechanisms have been proposed, one of which is that SHP2 may

act as an adaptor protein leading to the recruitment of the GRB2/SOS complex to the cell

membrane (50-53). Alternatively, SHP2 can dephosphorylate several targets, which, when

dephosphorylated, will promote ERK activation. One possible target is SPROUTY (SPRY),

which, when phosphorylated, purportedly sequesters GRB2/SOS in the cytoplasm (54-56). SHP2

also can dephosphorylate RAS-GAP binding site borne by RTKs or GAB1, and thereby exclude

GAPs from signaling complexes and promote RAS activation (57, 58). Several studies suggest

that SHP2 could act upstream of the SRC family kinases through dephosphorylation of SRC-

regulatory proteins (59-61).

In addition to RTKs, signals that activate G-protein coupled receptors (GPCRs), such as

lysophosphatidic acid (LPA), angiotensin II and beta-adrenergic agonists, can activate the

RAS/ERK pathway (62, 63). Integrins, which are integral membrane proteins that mediate

cellular adhesion to the extracellular matrix (ECM) and to other cells, also can lead to the

activation of the RAS/ERK cascade (64).

RAS family members belong to a large family of small GTPases that bind and hydrolyze

GTP. First discovered as transforming oncogenes of murine sarcoma viruses, three highly related

21 kDa mammalian proteins, Harvey-RAS (HRAS), Kirsten-RAS (KRAS), and Neuroblastoma-

RAS (NRAS) have been identified (65). RAS family members are anchored to the cytoplasmic

face of the plasma membrane, and such anchoring is essential for their biological function.

Membrane-targeting of RAS is achieved through lipid-anchors by post-translational lipid

modification of its C-terminal Cys residues (66). Unlike KRAS, NRAS and HRAS also signal

from endomembranes and cycle between the plasma membrane and Golgi apparatus depending

on their palmitoylation status (67). The localization to the inner leaflet brings RAS into close

proximity with RAS-GEFs, catalyzing the exchange of GDP for the more abundant GTP. GTP

loading alters RAS conformation, allowing it to interact with a number of downstream effectors

(68). Within the ERK signaling cascade, active RAS functions as an adaptor that binds to

effector RAF kinases with high affinity, causing their translocation to the cell membrane, where

RAF activation takes place. The detailed mechanism of RAF activation will be discussed later.

Active RAS-GTP is converted to the inactive RAS-GDP when its intrinsic RAS-GTPase activity

is stimulated by GTPase activating proteins (GAPs) (69). The balance between GEF and GAP

activity determines the guanine nucleotide status of RAS, thereby regulating RAS activity.

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The RAF family kinases, including ARAF, BRAF and RAF1/CRAF in vertebrates,

catalyze the phosphorylation and activation of the dual-specificity protein kinases, MEK1 and

MEK2 (also known as MKK1 and MKK2) (70). RAF family activation of MEK1/2 occurs

through phosphorylation of two serine residues, at positions 218 and 222 in the activation loop of

MEK1 (S222 and S226 on MEK2). Different RAF isoforms are not equal in their ability to

activate MEK: ARAF appears to be a poor MEK activator (71); BRAF displays a higher affinity

for MEK1 and MEK2 than RAF1, and is more efficient in phosphorylating MEKs (71-74).

MEK1/2 catalyze the phosphorylation of threonine and tyrosine residues in the activation

segment of ERK1/2, their only known physiological substrates (75). Several regulatory

phosphorylation sites on MEK outside the activation loop either positively or negatively regulate

the MAPKK. Phosphorylation of Ser298 by p21-activated kinase-1 (PAK1), downstream of the

small G-protein RAC, enhances the association of MEK with ERK, and results in RAF-

independent activation of MEK1 (76, 77). Conversely, in vivo phosphorylation by an unknown

kinase at Ser212, a site conserved in all MAPKKs, sharply decreases MEK1 activity (78). MEK1

and MEK2 can form heterodimers subject to negative feedback by ERK-catalyzed

phosphorylation of MEK1 on T292, which is absent on MEK2, facilitating the

dephosphorylation of S218/222 in the MEK1 activation loop (79).

The MAP kinases ERK1 and ERK2, also known as MAPK3 and MAPK1, are 44- and

42-kDa protein serine/threonine kinases, respectively (80). Initially isolated and cloned as

kinases activated in response to insulin and nerve growth factor (NGF) (81, 82), ERK1 and

ERK2 are expressed ubiquitously, with ERK2 levels generally higher than ERK1 (83). All

known cellular stimuli of the ERK1/2 pathway lead to the parallel activation of ERK1 and ERK2

(83, 84). Knocking-down ERK1 and/or ERK2 by RNA interference indicates that both ERK1

and ERK2 are positive regulators of cell proliferation and immediate-early gene (IEG)

transcription, and the positive role of ERK1 on cell proliferation is uncovered only when the

ERK2 level is markedly reduced and becomes limiting (83). Gene ablation studies in mice

demonstrate that either ERK may at least partially compensate for the other's loss, although some

evidence has been provided for differential functions of ERK1 and ERK2. The Erk1 gene is

dispensable for the development of mice, but ablation of the Erk2 gene is embryonic lethal (85,

86). Erk2-null mice fail to form mesoderm, although embryonic stem (ES) cell proliferation is

unaffected (86). The development of embryonic trophoblast and placental vasculature is severely

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impaired in Erk2-deficient mice, which leads to embryonic lethality (87, 88). Taken together,

these studies suggest that Erk1 is unable to compensate for Erk2 deficiency in vivo. Erk1-

deficient mice are viable, fertile, and of normal size with minor defects, such as impaired

terminal differentiation of T lymphocytes (85), and decreased adiposity with fewer adipocytes

(89), suggesting a specific role of Erk1 in thymocyte development and adipogenesis.

Dual threonine and tyrosine phosphorylations activate both ERKs, at Thr202/Tyr204 for

human ERK1 and Thr185/Tyr187 for human ERK2. Unlike MEK, significant ERK activation

requires phosphorylation at both sites, with tyrosine phosphorylation preceding that of threonine

(90). The ERKs are proline-directed protein kinases, phosphorylating Pro-neighboring Ser or Thr

residues. Docking sites present on physiological substrates confer additional specificity (91).

These docking interactions, through non-catalytic regions on ERK, team with scaffold proteins to

ensure signaling fidelity and enzymatic efficiency both to and from the MAPK. Unlike the RAF

kinases and MEK1/2, which have narrow substrate specificity, ERK1 and ERK2 have more than

175 documented cytoplasmic and nuclear substrates (92) (Figure 1-3).

Cytosolic substrates for ERK include several pathway components involved in negative

feedback regulation (Figure 1-3). Negative feedback by ERK has been proposed to occur through

direct phosphorylation of the EGFR at Thr669, which inhibits EGFR kinase activity (93, 94).

Multiple residues on SOS are phosphorylated by ERK following growth factor stimulation (95,

96). SOS phosphorylation destabilizes the SOS-GRB2 complex, eliminating SOS recruitment to

the plasma membrane and interfering with RAS activation of the ERK pathway. The RAF family

kinases also are substrates of activated ERK (97, 98). Six phosphorylation sites on RAF1 and

four phosphorylation sites on BRAF have been identified, which contribute to the down-

regulation of RAS/ERK pathway by inhibiting binding to activated RAS or disrupting

BRAF/RAF1 heterodimers (98). Finally, ERKs have also been demonstrated to negatively

regulate themselves by phosphorylating MAP kinase phosphatases (MKPs), which reduces the

degradation of these phosphatases through the ubiquitin-directed proteasome complex (99, 100).

MAPK-interacting kinase 1 (MNK1) and MNK2 are cytosolic serine/threonine protein

kinases initially discovered as ERK-interacting proteins (101). Both ERK and p38, but not JNK,

activate MNK by phosphorylation at Thr197 and Thr202. Activated MNK1, and possibly

MNK2, upregulates eukaryotic initiation factor-4E (eIF-4E) in vitro through phosphorylation at

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Figure 1-3. Downstream targets of ERK signaling.

ERK1 and ERK2 positively regulate transcription directly and indirectly via phosphorylation of

p90 ribosomal protein S6 kinases (RSKs), mitogen- and stress-activated protein kinases (MSKs),

and ternary complex factors (TCFs). Additionally, ERK1/2 indirectly regulate translation.

ERK1/2 also provide negative feedback loops for the RAS/ERK signaling pathway.

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Ser209, which is believed to enhance translation efficiency (102). The 90 kDa ribosomal S6

kinases (RSK) family of proteins are directly activated by ERK1/2 in response to growth factors,

many polypeptide hormones, chemokines and other stimuli (103, 104). RSKs are characterized

by the presence of two functional domains, the N-terminal kinase domain and the C-terminal

kinase domain, connected by a linker region, which are activated in a sequential manner by a

series of phosphorylation events following the binding of active ERK1 or ERK2 to an ERK

docking site (D domain) located at the extreme carboxyl terminus of cytoplasmic RSK (105).

RSKs phosphorylate many cytosolic and nuclear targets, regulating diverse cellular processes,

including cell proliferation, survival, growth and motility (106). For example, RSKs regulate

translation by modulating the PI3K-mTOR pathway at various steps (107-110). RSKs also play a

direct role in cell-cycle regulation. For example, RSKs have been shown to phosphorylate the

cyclin-dependent kinase (CDK) inhibitor p27KIP1, which promotes its association with 14-3-3,

prevents its translocation to the nucleus, and thereby promotes G1-phase progression (111).

Activated RSKs also translocate to nucleus and regulate transcription by direct phosphorylation

of transcription factors involved in IEG expression or by post-translational modification of IEG

products (104, 112) (Figure 1-3).

Upon phosphorylation, ERK1 and ERK2 translocate into the nucleus (113), where they

phosphorylate a wide range of targets, including transcription factors and a family of RSK-

related kinases, the mitogen- and stress-activated protein kinases (MSKs) (114). MSK1 and

MSK2 share the same tandem kinase structure as the RSKs, and also appear to be activated by

sequential phosphorylation following ERK docking. MSKs phosphorylate and activate the

activating transcription factor 1 (ATF1) at Ser63 (115), and the cAMP response element binding

protein (CREB) at Ser133 (116). MSKs also are the major kinases for Histone H3 and high-

mobility-group protein HMG-14, facilitating rapid induction of IEGs in response to mitogenic

and stress stimuli in fibroblasts (117). The best-characterized transcription factor substrates of

ERKs are ternary complex factors (TCFs), including Ets-like gene 1 (ELK-1) and SRF

Accessory Protein 1 (SAP1) and SAP2, which are directly phosphorylated by ERKs at multiple

sites (118, 119). Phosphorylated TCFs form complexes with the serum response factor (SRF) and

activate the transcription of numerous mitogen-inducible genes regulated by serum response

elements (SREs) (120). Another direct target of ERK is the proto-oncogene protein MYC, which

is a transcription factor that regulates cell-fate decisions, including proliferation, growth and

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apoptosis (121). Phosphorylation on Ser62 by ERK stabilizes MYC, and allows it to activate

transcription as a heterodimeric partner with MAX (122). ERK can also directly phosphorylate

components of activating protein-1 (AP1) family of transcription factors, c-JUN and c-FOS

(123).

Finally, the ERK pathway has been demonstrated to directly link growth factor signaling

to ribosome biogenesis. Following serum stimulation, ERK directly binds to and phosphorylates

the BRF1 subunit of the RNA polymerase (pol) III-specific transcription factor TFIIIB, which

enhances translational efficiency by affecting tRNA and 5S rRNA synthesis (124). ERK also

activates ribosomal RNA transcription following EGF stimulation by POL I through

phosphorylation of upstream binding factor (UBF), preventing its interaction with DNA (125).

1.2 Noonan syndrome and the RASopathies

1.2.1 Noonan syndrome

Noonan syndrome (NS; OMIM 163950) is the eponymous name for the genetic disorder

described by the pediatric cardiologist Jacqueline Noonan (126). NS is thought to be relatively

common, although its prevalence has not been determined accurately to date. Most authors cite

the figure of 1 in 1,000–2,500 live births first reported by Nora and colleagues (127). NS is

characterized by short shature, distinctive facial dysmorphic features, a wide spectrum of

congenital heart defects (CHD), and an increased risk of hematopoietic malignancy. Other

relatively common features are bleeding diathesis, ectodermal anomalies, lymphatic dysplasias,

cryptorchidism, and cognitive deficits (128-130) (Table 1-1).

Facial features of NS include high forehead, hypertelorism, downslanting palpebral

fissures, epicantal folds, ptosis, and low-set and/or posteriorly rotated ears (Figure 1-4). Besides

the short and/or webbed neck, a low posterior hairline is common. Cardiac defects and short

stature are the major reasons that patients with NS seek medical attention. While birth length is

typically normal, growth parameters usually drop below the 3rd centile during the first years of

life. Because there is often some attenuation and/or delay of the pubertal growth spurt, the

prevalence of short stature in NS is highest during the age of normal puberty. This is

accompanied by a delay in bone age. Although there have been reports of growth hormone (GH)

deficiency, neurosecretory dysfunction, or GH resistance in NS (131-133), a consistent pattern of

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Table 1-1. Clinical features of Noonan syndrome.

Short stature

Facial dysmorphism

Triangular face shape

Broad forehead

Hypertelorism (widely set eyes)

Epicanthal folds (extra fold of skin at the inner corner of the eye)

Ptosis (drooping of the eyelids)

Down-slanting palpebral fissures

Low set and/or backward rotated ears

Cardiovascular defects

Pulmonic stenosis

Atrial septal defects

Ventricular septal defects

Mitral valve defects

Hypertrophic cardiomyopathy

Hematological disorders

Bleeding diathesis

Thrombocytopenia

Leukemia

Lymphedema

Developmental

Delay

Attention deficit/hyperactivity disorder

Skeletal

Pectus excavatum and/or carinatum

Vertebral anomalies

Webbed neck with low posterior hairline

Cryptorchidism (undescended testicles)

Dental/Oral problems

Feeding difficulties

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Figure 1-4. Facial dysmorphism in Noonan syndrome.

Series of affected individuals heterozygous for mutations in different NS-associated genes are

shown. Adopted from Tartaglia M et al. Best Practice & Research Clinical Endocrinology &

Metabolism. 2011. 25(1): 161-179.

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abnormal GH secretion or action has not been shown so far, and it seems unlikely that there is a

simple link between GH and the growth deficits in NS. GH treatment in NS is still a matter of

debate. Recent research using our NS mouse model expressing the D61G mutant of Shp2,

showed that NS-causing Shp2 mutants inhibit insulin-like growth factor 1 (IGF-I) release via

GH-induced Erk hyperactivation, contributing to early postnatal growth delay (134).

NS patients exhibit a wide spectrum of cardiac disease (135, 136). Pulmonic stenosis

(PS), septal defects, and HCM occur most commonly, but other lesions also are observed.

Feeding problems are noted in the majority of affected infants and can cause failure to thrive

(137). Developmental delay and learning problems are quite common. Some motor delay can be

attributed to the hypotonia often observed in affected infants. An increased incidence of attention

deficit/hyperactivity disorder and frank mental retardation also are observed. Available data

indicate that the heterogeneity in the cognitive abilities observed in NS is at least partially

attributed to the specific causative gene mutation (138, 139). Skeletal anomalies most frequently

consist of pectus deformities, cubitus valgus, vertebral defects, and scoliosis. Children with NS

also are at increased risk of developing myeloproliferative disorder (MPD) (140). Juvenile

myelomonocytic leukemia (JMML; OMIM 607785), a rare MPD of childhood (141), arises at

increased prevalence in NS, although it affects only a small percentage of patients (142). The

MPD in most children with NS usually regresses without treatment, but occasionally follows an

aggressive clinical course.

The diagnosis of NS depends primarily on clinical features, although the prevalence of

the characteristic features among affected individuals has not been assessed rigorously and

depends on the patient’s age (143). In newborns, the facial features can be less apparent and

length is typically normal, but lymphedema and excess nuchal folds may be present. With time,

several features become more obvious, including facial dysmorphism, pectus deformities, and

reduced growth. Hypertrophic cardiomyopathy (HCM) also may develop during the first few

years of life. The facial features also can become more difficult to detect in later adolescence and

adulthood.

NS is a Mendelian trait transmitted in an autosomal dominant manner, and as observed

for other dominant disorders, a significant percentage of cases is due to de novo mutations.

Although NS is genetically heterogeneous (31, 32, 144), all known cases are caused by germ-line

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mutations in conserved components of the canonical RAS/ERK pathway. Mutations in PTPN11,

which encodes the protein tyrosine phosphatase SHP2, account for approximately half of NS

cases (145). Other known NS genes include SOS1 (~10%) (146, 147), RAF1 (3-5%) (148, 149),

KRAS (<2%) (150, 151), NRAS (152) and BRAF (153) (<1%). The causative genes responsible

for the remaining 30% of NS cases remain to be identified. Mutations in some of these genes, as

well as in genes encoding other RAS/ERK pathway components, also cause phenotypically

related disorders, such as neurofibromatosis type 1 (NF1), Costello syndrome, cardio-facio-

cutaneous syndrome (CFCS), and LEOPARD syndrome; together with NS, these syndromes are

now termed “RASopathies” (32) (Figure 1-5).

The most common gene associated with NS is PTPN11, which accounts for

approximately 50% of cases. SHP2, the protein product of PTPN11, is a non-receptor protein-

tyrosine-phosphatase (PTP) that positively modulates RAS/ERK signaling (154). SHP2 is

composed of two tandem N-terminal SH2 domains (N-SH2 and C-SH2), which function as

phospho-tyrosine binding domains and mediate the interaction of this PTP with its substrates,

followed by a catalytic PTP domain and a C-terminal tail with tyrosyl phosphorylation sites and

a proline-rich motif (155). In the basal state, the catalytic function of the protein is auto-inhibited

by interaction between the N-SH2 domain and the PTP domain, which blocks substrate access.

Binding of an appropriate phosphotyrosyl peptide alters the conformation of the N-SH2 domain,

prevents its binding to the PTP domain and causes catalytic activation of the protein (156). Most

NS-causing missense mutations in PTPN11 affect residues involved in the N-SH2/PTP auto-

inhibitory binding and up-regulate SHP2 function by impairing the switch from the active to

inactive conformations. There also are mutations affecting residues located in the

phosphopeptide-binding pocket of the N-SH2 or C-SH2 domains or in the linker stretch

connecting these domains, which promote SHP2 gain of function by increasing the binding

affinity, altering the binding specificity, or changing the flexibility of the N-SH2 domain in a

manner that inhibits the N-SH2/PTP interaction (157-159).

SOS1 is the second most frequently mutated NS disease gene, accounting for

approximately 10% of cases (146, 147, 160). The majority of NS-associated SOS1 mutations

affect multiple domains that are responsible for stabilizing the protein in a catalytically

autoinhibited conformation. Approximately half affect residues located in the short helical linker

connecting the plekstrin homology (PH) and the RAS exchanger motif (REM) domains. A

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Figure 1-5. RASopathies and the RAS-RAF-MEK-ERK MAPK pathway.

Schematic diagram showing the RAS/ERK pathway and affected disease genes in RASopathies.

NS, Noonan syndrome; NS/LAH, Noonan-like syndrome with loose anagen hair; LS,

LEOPARD syndrome; CS, Costello syndrome; CFCS, cardiofaciocutaneous syndrome; NF1,

neurofibromatosis type 1; NFLS, neurofibromatosis type 1-like syndrome (also known as Legius

syndrome).

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second mutation cluster is located within the PH domain, while a third functional cluster resides

at the interacting regions of the Dbl homology and REM domains. A single amino acid change

(E846K) within the Cdc25 domain accounts for more than 10% of NS cases with SOS1

mutations. Biochemical data demonstrate that NS-causing SOS1 mutations disrupt the auto-

inhibition of SOS1 RAS-GEF activity resulting in gain-of-function of SOS1 and a subsequently

enhanced RAS/ ERK activation (146, 147).

Germline KRAS mutations account for a small percentage (< 2%) of affected subjects in

NS (150, 151, 161). NS-causing KRAS mutations up-regulate protein function and increase

signaling down the RAS/ERK pathway by either reducing the intrinsic or GAP-stimulated

GTPase activity and, consequently, impairing the switch between the active and inactive

conformation, or by interfering with the binding of KRAS to guanine nucleotide (150, 162).

More recently, two germline missense mutations of NRAS (T50I and G60E) conferring enhanced

stimulus-dependent MAPK activation, were reported to account for a few NS cases (152).

Two groups (148, 149) identified multiple missense mutations of RAF1 in NS, which

cluster in three regions (Figure 1-6). Approximately 70% of NS-associated RAF1 alleles alter the

motif flanking S259 within the so-called CR2 domain, which binds to 14-3-3 proteins and is

critical for auto-inhibition (163, 164). The second group of mutations (~15%) affects residues

within the activation segment of the kinase domain (D486 and T491). The remaining alleles

(~15%) involve two adjacent residues (S612 and L613) located C-terminal to the kinase domain.

Transient transfection studies indicate that mutations affecting the 14-3-3 binding motif or the C-

terminus of the protein enhance RAF1 kinase activity and increase MEK/ERK activation in cells.

By contrast, mutations that cluster in the activation segment are kinase-impaired and reportedly

act as dominant negative or null alleles (148, 149). Previous work suggested that the increased

kinase activity of NS-associated CR2 domain mutants results from decreased S259

phosphorylation and consequent dissociation from 14-3-3 (149, 165, 166), but the mechanism

underlying increased kinase activity of the RAF1 C-terminal mutants remained unclear.

Likewise, how kinase-defective RAF1 alleles cause NS had remained obscure, if not paradoxical.

Studies of kinase-defective BRAF alleles strongly implicate enhanced MEK/ERK activation and

heterodimerization with RAF1 in human melanoma pathogenesis (167, 168). The paradoxical

activation of the MEK/ERK pathway in wild type cells treated with selective small molecule

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Figure 1-6. RAF1 domain structure and location of residues altered in NS.

RAF1 protein domains are indicated (CR, conserved region) along with two serine residues

(below the cartoon), phosphorylation of which serve as 14-3-3 binding sites. The mutations

observed in NS are shown above the cartoon.

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BRAF inhibitors also has been attributed to the ability of these inhibitors to induce BRAF/RAF1

heterodimer formation (169, 170). The relevance of these observations for RAF1 alleles

expressed at physiological expression levels remained to be determined.

Germline BRAF mutations have also been documented in a small percentage of subjects

with phenotypes fitting or suggestive of NS (< 1% of cases) (153). Of note, BRAF has previously

been identified as a major disease gene underlying CFCS (50-75% of cases) (171, 172). NS-

associated mutations largely do not overlap with those occurring in CFCS, suggesting a

genotype-phenotype correlation.

NS is characterized by marked phenotypic variability, which can be explained, in part, by

the underlying molecular lesions. PTPN11 mutations are more prevalent among subjects with PS

and short stature, and less common in individuals with HCM and/or severe cognitive deficits

(173, 174). JMML is associated with a narrow spectrum of mutations affecting the PTPN11 gene

(142, 175), but also can be associated with certain germline KRAS mutations (150). Germline

KRAS mutations are generally associated with a highly variable but generally severe phenotype

(150, 151, 161). Subjects with a mutated SOS1 allele tend to exhibit a distinctive phenotype

characterized by ectodermal abnormalities and generally associated with a lower prevalence of

cognitive deficits and short stature (146, 160). NS patients with RAF1 mutations have a much

higher incidence (~75%) of hypertrophic cardiomyopathy (HCM) than is found in the overall NS

population (~18%). Notably, kinase-activating and kinase-impaired RAF1 alleles are associated

with different syndromic phenotypes. Only RAF1 alleles encoding kinase-activated mutants are

highly associated (~95%) with HCM (148, 149).

1.2.2 RASopathies: disorders clinically related to Noonan syndrome

1.2.2.1 LEOPARD syndrome

LEOPARD syndrome (LS; OMIM 151100) is a rare autosomal dominant disorder that

overlaps phenotypically with NS, including a ‘Noonan-like’ appearance as well as Lentigines,

ECG conduction abnormalities, Ocular hypertelorism, Pulmonic stenosis, Abnormal genitalia,

Retardation of growth, and sensorineural Deafness (acronym LEOPARD). Similar to NS, there

are age-dependent aspects of the phenotype. Craniofacial dysmorphism is similar to that of NS

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but is usually milder (176). Multiple lentigines, which are flat, black-brown macules, are

dispersed primarily on the face, neck, and upper part of the trunk, sparing the mucosae. Growth

retardation is observed in 25% of affected individuals. Approximately half of affected

individuals have heart defects, which are similar to those in NS but occur with different

frequencies. ECG anomalies, progressive conduction defects and HCM are the most frequent

features. Of note, HCM is detected in up to 80% of LS patients with heart defects, most

commonly appearing during infancy. LS is allelic with NS with a restricted spectrum of

mutations in PTPN11 that cause decreased phosphatase activity and account for the vast majority

of affected individuals (~90%) (Figure 1-5) (177-180). In a small proportion of cases, LS has

been linked to mutations in RAF1 or BRAF (149, 153, 181).

1.2.2.2 Noonan-like syndrome with loose anagen hair

Patients with Noonan-like syndrome with loose anagen hair (NS/LAH; OMIM 607721)

show facial features reminiscent of NS, including macrocephaly, high forehead, hypertelorism,

and low-set and posteriorly rotated ears, in addition to short and wedded neck and pectus

abnormalities (182-184). Ectodermal involvement, severe short stature associated with proven

GH deficiency (GHD), significant cognitive deficits and distinctive hyperactive behavior are

other common features associated with these subjects. The hair anomalies include easily

pluckable, sparse, thin, and slow-growing hair in the anagen phase, which fit a well-known

condition termed loose anagen hair (LAH) syndrome (185), and suggest that this is a disorder

distinct from NS. Most affected individuals also have darkly pigmented skin with eczema or

ichthyosis. Cardiac anomalies are observed in the majority of cases, with mitral valve and septal

defects significantly overrepresented compared with the general NS population. The voice of

affected individuals is characteristically hypernasal. To date, NS/LAH appears to be genetically

homogeneous, as all affected individuals share the same 4A>G missense mutation (an S2G

amino acid substitution) in SHOC2 (Figure 1-5) (183), which encodes a leucine-rich repeat-

containing scaffold protein required for the efficient transmission of information from RAS to

the MAPK cascade (186). Functional studies of the SHOC2-S2G mutant demonstrated aberrant

protein N-myristoylation, which resulted in aberrant targeting of SHOC2 to the plasma

membrane and impaired translocation to the nucleus upon growth factor stimulation.

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1.2.2.3 CBL-mutation associated Noonan-like syndrome

Three independent studies recently reported that heterozygous germline mutations in the

CBL tumor suppressor gene, which is mutated in myeloid malignancies and encodes a

multivalent adaptor protein with E3 ubiquitin ligase activity, underlie a previously unrecognized

condition with clinical features fitting or partially overlapping NS in some individuals (Figure

1-5) (187-189). Missense mutations alter the evolutionarily conserved residues located in the

RING finger domain or the linker connecting this domain to the N-terminal tyrosine kinase

binding domain, a known mutational hot spot in myeloid malignancies that affects CBL-

mediated receptor ubiquitylation and dysregulates signal flow through RAS (187). Common

features occurring in CBL mutation-positive subjects include variable developmental delay,

reduced growth, facial dysmorphism, café-au-lait spots, and predisposition to JMML during

childhood (188, 189).

1.2.2.4 Neurofibromatosis type 1 and related phenotypes

Neurofibromatosis type 1 (NF1; OMIM 162200) is one of the most common autosomal

dominant disorders (1 in 3000–4000 live births) caused by a single gene (190, 191). NF1 is

characterized by multiple café-au-lait spots, axillary and inguinal freckling, Lisch nodules of the

iris, cutaneous, subcutaneous and/or plexiform neurofibromas, skeletal dysplasia including short

stature, dystrophic scoliosis and sphenoid wing dysplasia, learning deficits and behavioral

abnormalities, and a predisposition for developing benign and malignant neoplasms, such as

malignant peripheral nerve sheath tumors (MPNST) and optic pathway gliomas (OPG).

Cardiovascular abnormalities in NF1 include congenital heart defects such as pulmonary artery

stenosis, vasculopathy and hypertension. Heterozygous mutations of the NF1 gene, which

encodes Neurofibromin, a RAS-GTPase activating protein (RASGAP) that activates the intrinsic

GTPase activity of RAS and negatively regulates its role in signal transduction, are observed in

the vast majority of affected individuals (>90%) (Figure 1-5) (192). Loss-of-function mutations

or deletions of the NF1 gene lead to increased RAS/ERK signalling.

A disorder related to NF1 is NF1-like syndrome (NFLS; OMIM 611431), also known as

Legius syndrome (193, 194). The most common clinical features of NFLS are multiple café-au-

lait spots, axillary freckling, macrocephaly, NS-like facial dysmorphism, learning disabilities,

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and attention deficit-hyperactive disorder (ADHD). Despite the clinical overlap with NF1, some

typical features of NF1, such as Lisch nodules, neurofibromas and central nervous system tumors

are absent in NFLS patients. NFLS is caused by loss-of-function mutations of the SPRED1 gene

(193, 195), which encodes Sprouty-related EVH1 domain-containing protein 1 (SPRED1), a

negative modulator of RAS/ERK pathway that suppresses phosphorylation and activation of

RAF (Figure 1-5) (196).

1.2.2.5 Cardiofaciocutaneous syndrome

Cardiofaciocutaneous syndrome (CFCS; OMIM 115150) is an extremely rare and severe

genetic disorder characterized by cardiac abnormalities, distinctive craniofacial dysmorphism,

cutaneous abnormalities, failure to thrive, severe feeding problems, developmental delay,

reduced growth, and abnormalities of the gastrointestinal tract and central nervous system (197,

198). Recurrent craniofacial features include macrocephaly, which is usually associated with

prominent forehead, bitemporal constriction, and facial dysmorphia that is coarser compared

with NS. The skin abnormalities include dryness and hyperkeratosis, ichthyosis, eczema,

unusually sparse, brittle and curly scalp hair, and absent/sparse eyebrows and eyelashes.

Pigmentary changes (such as café-au-lait spots, nevi or lentigines) and hemangiomas are

observed. Cardiac defects occur in the majority of affected individuals, and consist of pulmonic

stenosis and other valve dysplasias, septal defects, HCM and rhythm disturbances. Some form of

neurologic and/or cognitive delay (ranging from mild to severe) is seen in all affected

individuals; seizures are also frequent. CFCS is genetically heterogeneous, and is usually the

result of a de novo dominant mutations in the KRAS, BRAF, MAP2K1 or MAP2K2 genes, which

occurr in approximately 60-90% of affected individuals (Figure 1-5) (171, 172).

1.2.2.6 Costello syndrome

Costello syndrome (CS; OMIM 218040) is the eponymous name for a rare genetic

disorder described by pediatrician Dr. Jack Costello in 1970, as a condition characterized by

prenatal overgrowth followed by severe failure to thrive, delayed development and mental

retardation, distinctive coarse facial features, short stature, and cardiac defects (most commonly

HCM, septal defects, valve thickening and/or dysplasia, and tachycardia), and musculoskeletal

and skin abnormalities (unusually flexible joints and loose folds of extra skin on hands and feet)

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(199, 200). Beginning in early childhood, affected individuals are predisposed to certain

malignancies (most commonly rhabdomyosarcoma, neuroblastoma and transitional cell

carcinoma) (201). Cutaneous papillomas in the perinasal and perianal regions are the most

common benign tumors seen with this condition. CS is caused by germline missense mutations in

HRAS (Figure 1-5) (202), but not in other RAS family genes. CS-associated HRAS mutations at

codons 12 and 13 promote the active, GTP-bound conformation and constitutively activate

downstream effectors such as MAP kinase, PI-3 kinase and RALGDS (203).

1.2.3 Noonan syndrome mouse models

In order to study the molecular pathogenesis of NS, several mouse models carrying NS-

associated mutations have been generated and characterized in our and other laboratories. Araki

et al. generated the first knock-in mouse model expressing the NS-associated gain-of-function

mutant Shp2D61G (204). When homozygous, the D61G mutant causes embryonic lethality,

whereas heterozygotes have decreased viability. Surviving Ptpn11D61G/+ embryos (approximately

50%) have short stature, craniofacial abnormalities similar to those in NS patients, and MPD.

Severely affected Ptpn11D61G/+ embryos (approximately 50%) have multiple cardiac defects,

including atrial, atrioventricular or ventricular septal defects, double-outlet right ventricle

(DORV), and markedly enlarged outflow tract and atrioventricular valve primordia, similar to

those in mice lacking Nf1 (205, 206). Their endocardial cushions have increased Erk activation,

but Erk hyperactivation is cell- and pathway-specific. These data show that a single Ptpn11 gain-

of-function mutation can evoke all major features of Noonan syndrome by acting on multiple

developmental lineages in a gene dosage-dependent and pathway-selective manner. Transgenic

mice expressing a different NS-associated Ptpn11 mutant (Q79R) also show valvulospetal

defects and facial abnormalities seen in NS patients, which are prevented by the genetic ablation

of Erk1/2 or pre-natal pharmacological inhibition of Mek (207-209).

Later, using an inducible gene knock-in approach, our laboratory further elucidated the

mechanism underlining the cardiac defects in NS caused by Ptpn11 mutations, and showed that

all cardiac defects in NS result from mutant Shp2 expression in the endocardium, not in the

myocardium or neural crest (210). Moreover, the penetrance of NS defects is affected by genetic

background and the specific Ptpn11 allele. Finally, ex vivo assays and pharmacological

approaches showed that NS mutants cause cardiac valve defects by increasing Erk MAPK

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activation. However, we did not observe HCM in these NS mouse models caused by Ptpn11

mutations.

Chen et al., in collaboration with our laboratory, observed that activation of multiple

signaling pathways causes developmental defects in mice with a Noonan syndrome–associated

Sos1E846K gain-of-function mutation (211). Both heterozygous and homozygous mutant mice

showed many NS-associated phenotypes, including growth defects, distinctive facial dysmorphia,

hematologic abnormalities, and cardiac defects. The phenotypes in Sos1E846K mice and our NS-

associated Ptpn11 mutation mouse models (204, 210) overlap to some degree, but are

distinguishable. Sos1E846K/+ mice develop left ventricular hypertrophy (LVH) and fibrosis, with

or without aortic stenosis (AS). Sos1E846K mutation appears to selectively affect semilunar valves,

whereas Ptpn11D61G has a more profound effect on atrioventricular valves. Although NS patients

with SOS1 mutations develop PS, PS was not observed in the Sos1E846K/+ hearts. The Ras/Erk

pathway, as well as Rac and Stat3, are activated in Sos1E846K mutant hearts, suggesting that Rac

and Stat3 activation might also contribute to NS phenotypes. Furthermore, prenatal

administration of a MEK inhibitor ameliorated the embryonic lethality, cardiac defects, and NS

features of the homozygous mutant mice.

1.3 Function and regulation of RAF1

The first raf gene was described in 1983 as a retroviral oncogene, v-raf, transduced by

the murine sarcoma virus (MSV-3611) (212). Later, an avian homolog, v-mil, was found in the

acutely transforming avian retrovirus MH2 (213). These two retroviruses encoded the first

oncogene with serine/threonine kinase activity to be discovered (214). After the cellular proto-

oncogenes c-raf (215) and c-mil (216) were cloned, RAF proteins have been studied intensely.

Initial studies demonstrated that CRAF (also known as RAF1) plays a critical role in mediating

the cellular effects of growth factor signals (217-219). Later, RAF proteins were identified as the

direct activators of MEK (220, 221) and as downstream effectors of RAS (222-226), acting as

essential connectors between RAS and the MEK-ERK pathway. Most subsequent work focused

on understanding this role and the regulation of RAF proteins in detail, until new kinase-

independent functions of RAF1 in the regulation of apoptosis (227-229) and cell migration (230)

emerged in the last decade.

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Three different RAF isoforms, originating from three independent genes, are present in

mammals: RAF1/CRAF, BRAF, and ARAF. All Raf isoforms are expressed ubiquitously in

embryonic and adult mouse tissues. However, the Araf and Braf genes are more restricted in their

expression, with Araf mRNA expressed particularly highly in urogenital organs and Braf mRNA

abundant in neuronal tissues (231-233). Genetic studies in mice have shown that the Raf proteins

have non-redundant functions in development. On a predominantly C57BL/6 genetic background,

Araf-deficient mice survive to birth, but die between 7 and 21 days post-partum, displaying

neurological and gastrointestinal defects (234). By contrast, Araf-/- animals on a 129/OLA

background, survive to adulthood, are fertile and do not display obvious intestinal abnormalities,

although they do have a subset of the neurological defects seen on the C57BL/6 background.

Mice with a targeted disruption in Braf die of vascular defects during mid-gestation. Braf-/-

embryos show growth retardation, an increased number of endothelial precursor cells,

dramatically enlarged blood vessels and apoptotic death of differentiated endothelial cells (235).

Raf1-deficient embryos also are growth retarded and die at mid-gestation with vascular defects in

the yolk sac and placenta, abnormalities in the fetal liver, as well as increased apoptosis of

embryonic tissues (236-238). The lack of compensation between the Raf proteins in mice does

not seem to be the result of differences in expression patterns, which implies that the different

isoforms have distinct functions.

1.3.1 Structure of RAF family kinases

All RAF isoforms share a common structure, consisting of three conserved regions (CR)

with distinct functions (Figure 1-7). The CR1 region contains elements required for membrane

recruitment, including a RAS binding domain (RBD), which binds to active RAS-GTP, and a

cysteine-rich domain (CRD), which has a zinc finger structure homologous to those of protein

kinase C (PKC). The CRD stabilizes the association with RAS through interaction with the lipid

moiety present on processed RAS, and also is necessary for the interaction of CR1 with the

kinase domain for RAF autoinhibition (222, 239-242). CR2 contains important inhibitory

phosphorylation sites participating in negative regulation of RAS binding and RAF activation

(243). Finally, CR3 comprises the kinase domain, including the activation segment, whose

phosphorylation is crucial for kinase activation (244). In addition, phosphorylation of residues in

the negatively-charged (N) region upstream of the CR3 is necessary for RAF activation (245).

Functionally, the RAF structure can be split into a regulatory N-terminal region, containing CR1

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Figure 1-7. Common structure of RAF proteins.

Schematic structure of RAF proteins depicting the three conserved region (CR1-3), the RAS-

binding domain (RBD) and cysteine-rich domain (CRD), as well as the negative-charged

regulatory region (N-region), glycine-rich loop (G-loop) and activation segment in which the

phosphorylation sites crucial for activation are located.

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and CR2, which is critical for activation as well as inhibitory phosphorylation, and a catalytic C-

terminal region, which includes the phosphorylation sites necessary for kinase activation. The

regulatory domain restrains the activity of the kinase domain (240, 246, 247) and its removal

results in constitutive oncogenic activation (248). However, the activity of the isolated RAF1

kinase domain is subjected to further regulation and can be stimulated by phorbol esters, v-Src,

and phosphorylation (247, 249). This observation is in keeping with the finding that the most

common oncogenic mutation in BRAFV600E activates BRAF kinase activity by mimicking

phosphorylation of the activation loop that releases its inhibitory interaction with the ATP-

binding domain (167).

1.3.2 Regulation of RAF1

RAF regulation is highly complex and involves many steps, including membrane

recruitment, dimerization, protein-protein interaction, conformational changes and

phosphorylation (250-252).

1.3.2.1 The RAF1 activation/deactivation cycle

In the quiescent state, RAF1 is thought to exist in a closed, inactive conformation

wherein the N-terminal regulatory region folds over and blocks the catalytic region (Figure 1-8)

(253). This conformation is stabilized by 14-3-3 dimer binding to an N-terminal site, phospho-

S259 (pS259), and a C-terminal site, pS621 (Figure 1-9). Although RAF1 activation is

incompletely understood, numerous studies have suggested the following sequence of events

(Figure 1-8).

Dephosphorylation of pS259 at the cell membrane by specific phosphatases, including

protein phosphatase 2 (PP2A) and PP1, releases 14-3-3 from its N-terminal binding site on RAF1,

allowing conformational changes to occur that unmask the RBD and CRD in the CR1 region to

enable RAS binding and membrane recruitment (164, 254-258). RAS activation is under

negative feedback regulation mediated by ERK and its downstream substrate RSK, which

phosphorylate and inhibit SOS1 (95, 259). On the other hand, binding of RAF1 to RAS can be

facilitated by the scaffolding protein SUR-8/SHOC2 (186). The CRD is necessary, but not

sufficient, for stable membrane recruitment and activation of RAF1 (260, 261). RAF

translocation to the membrane also is aided by the ability of RAF to interact with lipids (262,

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263). These data suggest that the CRD may stabilize the primary recruitment of RAF1 exerted by

the RBD through forging interactions with the lipid tails of RAS proteins (264). Furthermore,

RAS isoforms reside in different subcellular compartments, which can influence interactions

with RAF family members, and can profoundly influence the mechanism and kinetics of RAF

activation (265).

Phosphorylation of multiple residues in the N-region upstream of CR3 and in the

activation segment in CR3 is required for full RAF1 activation (Figure 1-8 and Figure 1-9). The

N-region in RAF1 contains the S338/9 and Y340/1 phosphorylation sites, which are not only

essential for full kinase activation but also for interaction with the substrate MEK (266-268).

PAK (269, 270) and SRC family tyrosine kinases (271, 272), respectively, reportedly

phosphorylate these sites. Other studies suggest that Ser338 is autophosphorylated upon RAF

dimerization induced by mitogens (273), whereas others demonstrate that casein kinase 2 (CK2)

phosphorylates Ser338 as a component of the KSR1 scaffold complex recruited to RAF (274).

S338/9 phosphorylation itself only slightly elevates RAF1 kinase activity and mainly seems to

serve as a priming event that initiates further activating modifications (266). S471 has been

identified as a growth factor-induced RAF1 phosphorylation site, which is critical for RAF1

kinase activity and MEK binding (275). Finally, the phosphorylation of T491/S494 in the

activation loop is RAS-dependent and required for full activation (244, 276), but the identity of

the respective kinase(s) is unknown. Beside these major sites, phosphorylation at several minor

sites also has been reported, including T268/269 (277, 278), T481 (275) and S497/499 (279).

The role of these phosphorylations in RAF1 regulation remains unresolved, but they do not

notably affect RAF1 activation. RAF homodimerization and heterodimerization have recently

emerged as important regulatory mechanisms that drastically enhance kinase activity and

downstream signaling, and are discussed in detail later.

Deactivation of RAF1 is initiated by specific binding of protein phosphatase 5 (PP5) to

activated RAF1, which results in the dephosphorylation of pS338 (280). The phosphorylated N-

region also serves as a binding site for the RAF kinase inhibitor protein (RKIP) (281, 282),

which dissociates RAF1 from its substrate MEK (283, 284). In addition, RAF1 is subjected to

direct feedback phosphorylation on multiple sites by ERK (Figure 1-9), which inhibit the

activation of RAF1 by RAS and promote the subsequent dephosphorylation and resensitization

of RAF1 by PP2A (97). Negative feedback from ERK to RAF1 also is suggested by a systematic

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Figure 1-8. Overview of RAF1 activation/deactivation.

In quiescent cells, intramolecular autoinhibition of RAF1 is stabilized by the binding of 14-3-3 to

the pS259 and pS621 residues. Upon membrane recruitment by activated RAS, pS259 is

dephosphorylated by PP1 or PP2A. Subsequently, phosphorylation of the N-region and

activation loop by as yet incompletely defined upstream kinases, and homodimerization or

heterodimerization with other RAF isoforms, cause full activation of RAF1. Following

MEK/ERK activation, RAF1 is deactivated through PP5-mediated dephosphorylation of pS338,

and ERK-mediated feedback phosphorylation, which desensitize the kinase. PP2A, through the

prolyl isomerase PIN1, and maybe other unknown phosphatases, dephosphorylate the remaining

activating sites and the ERK feedback sites. RAF1 ultimately reverts to its closed, inactive

conformation upon rephosphorylation of S259 by AKT, PKA or unknown kinases, allowing

intramolecular bidentate 14-3-3 rebinding. Activating events are colored red and deactivating

processes are in black.

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Figure 1-9. Regulatory phosphorylation sites of RAF proteins.

The structure and phosphorylation residues of the three RAF isoforms. Red residues indicate

activating phosphorylation sites, black are ERK feedback phosphorylation sites, and blue are 14-

3-3 binding sites.

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analysis of feedback regulation of the RAS/ERK pathway based on mathematical modeling (285),

although several ERK feedback phosphorylation sites also were described as stimulating RAF1

activity (286).

RAF1 also is negatively regulated by cAMP–activated kinase (PKA)-catalyzed

phosphorylation of several sites, including S43, S233, S259 and S621. The phosphorylation of

S43 interferes with RAS binding (287, 288), whereas phosphorylation of S233 and S259

enhances the binding of 14-3-3 and suppresses RAF1 kinase activity (289, 290). Studies also

show that S259 could be phosphorylated by AKT directly or indirectly (291-293). The

phosphorylation of S621 and binding of 14-3-3 to pS621 appears to have multiple roles in RAF1

regulation. On the one hand, it serves as a negative regulatory phosphorylation site to stabilize

the closed, inactive conformation in resting cells (294, 295); on the other hand, its inhibitory

function is converted into an essential component of RAF1 activation following stimulation

(296). For example, pS621 increases the stability of RAF1 by preventing proteasome-mediated

degradation (297). Binding of 14-3-3 to pS621 also enhances RAF dimerization (98, 168, 298,

299), and is required for the kinase domain to bind ATP (296). However, several studies

demonstrate that S621 phosphorylation is largely mediated by autophosphorylation (296, 297).

The phosphatase(s) responsible for dephosphorylating S621 is(are) currently unknown.

1.3.2.2 RAF homodimers and heterodimers

Dimerization is a frequent mechanism for the activation of kinases. Homodimerization

was initially highlighted as a potentially important step of RAF1 activation by two studies

showing that forced interaction of RAF1 monomers tagged with inducible dimerizing tags

robustly induced kinase activity (300, 301). Both studies proposed that active RAS would

promote the formation of dimers. This hypothesis was later extended to heterodimerization

between RAF1 and BRAF, which was found to be inducible by active RAS (302). These initial

studies showed that homodimerization and heterodimerization can hyperactivate RAF kinases.

Later, Rushworth et al. demonstrated that endogenous BRAF and RAF1 heterodimerize in

multiple cell lines in response to mitogens (298). Biochemical fractionation of RAF heterodimers

from homodimers and monomers showed that RAF1/BRAF heterodimers accounted for the

majority of the mitogen-induced RAF kinase activity, suggesting that RAF1 and BRAF

cooperate to drastically elevate their kinase activity when forming heterodimers. Given that

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heterodimers between wild-type RAF1 and kinase-defective BRAF still display elevated kinase

activity, as do heterodimers between wild-type BRAF and kinase-impaired RAF1, either kinase-

competent RAF isoform is sufficient to confer high catalytic activity to the heterodimers.

Heterodimerization plays a pathological role in certain cancers. When BRAF mutations

were discovered in cancer (35), a puzzling observation was that while the most frequent mutation,

V600E, massively enhanced BRAF kinase activity, several less frequent BRAF mutations only

mildly increased, or even impaired, kinase activity (167). Nevertheless, even kinase-impaired

BRAF mutants could hyperactivate the MEK/ERK pathway. Intriguingly, this activation was

dependent on the presence of RAF1, and a subsequent study demonstrated that the kinase-

impaired BRAF mutants found in human cancers indeed could promote RAF1 heterodimer

formation (168). Although physiological RAF1/BRAF heterodimerization is induced by RAS

activation, oncogenic BRAF mutants constitutively dimerize with RAF1 (168, 298). More

recently, an important role for RAF heterodimers in response to ATP-competitive RAF kinase

inhibitors was described. The original observation that RAF inhibitors paradoxically induce ERK

cascade signaling can now been explained by the ability of RAF inhibitors to promote RAF

heterodimerization and activation in the presence of oncogenic or normally activated RAS (169,

170, 303).

Mutation of S621 abrogates RAF heterodimerization, suggesting that 14-3-3 binding to

pS621 is essential for RAF heterodimerization (302). Indeed, heterodimerization is enhanced by

wild type 14-3-3, but not a dimerization-negative 14-3-3 mutant, suggesting that the 14-3-3

dimer crosslinks RAF1 and BRAF by binding to the C-terminal sites on each kinase (298). This

observation suggests a mechanism for how 14-3-3 can stabilize both inactive and active RAF1

conformations. In the inactive conformation, 14-3-3 clasps the RAF1 regulatory domain to the

kinase domain via intramolecular binding to pS259 in the N-terminus and p621 in the C-terminus.

Dephosphorylation of pS259 and binding to activated RAS displaces 14-3-3 from S259, leaving

one 14-3-3 arm free to contact with the 14-3-3 binding site on BRAF in order to facilitate

heterodimerization. RAF heterodimerization also is regulated by ERK-mediated feedback

phosphorylation on RAF (97, 298). The feedback phosphorylation mainly serves to limit the

lifetime of RAF1/BRAF heterodimers, and mutation of the relevant sites enhances ERK

signaling and the associated biological activities. Other regulators include KSR1 (304) and

MLK3 (305), which both enhance heterodimerization.

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The precise mechanism by which the juxtaposition of RAF molecules stimulates kinase

activity was elucidated recently by Rajakulendran et al. (299). They demonstrated that activation

of the kinase domain of RAF is controlled by an allosteric interaction between two kinase

domains in a specific side-to-side dimer configuration, which is thought to position a critical

helix (helix αC) in the kinase domain in a productive conformation necessary for catalytic

activity (Figure 1-10). The dimer interface region in the RAF kinase domain is conserved in the

KSR kinase domain, which also can serve as an allosteric activator of RAF by forming

KSR/RAF side-to-side heterodimers. Given that all isoforms of RAF and KSR share a nearly

identical side-to-side dimer interface, the question still remains as to how the cell regulates

specific dimer formation between the multiple isoforms of RAF and KSR proteins found in

higher-order metazoans. Presumably, cells must have a separate mechanism for selectively

driving dimer formation between any two specific isoforms. For example, a recent study showed

that most RAF inhibitors induce KSR1/BRAF binding, but promote little complex formation

between KSR1 and RAF1 (306). These works also found that KSR1 competes with RAF1 for

inhibitor-induced binding to BRAF.

1.3.2.3 Scaffolds and modulators of RAF signaling

Multiple studies have revealed scaffolding as a mechanism that helps the ERK cascade to

transduce signals with high efficiency and specificity. Scaffolds also appear to be involved in the

spatial restriction of ERK activity and signaling to distinct subcellular compartments (Figure

1-11) (307). Therefore, scaffolds can have a huge impact on the biochemical and biological

behavior of the ERK pathway (308, 309). However, our knowledge of their role in the functional

modulation of the pathway and their exact mechanism of action is still limited.

The best-characterized scaffold of the ERK pathway is kinase suppressor of Ras (KSR)

(310). KSR1 has a kinase domain with high homology with RAF1. But this kinase domain

contains mutations in residues critical for catalytic activity. Whether KSR1 has kinase activity or

is a pseudokinase have been debated. As a scaffold for the ERK pathway, KSR1 can interact

with all kinases of the ERK pathway. MEK is bound constitutively, while RAF and ERK are

recruited to KSR1 upon mitogen stimulation (311, 312). However, KSR1 binds less than 5% of

endogenous RAF1 (313), indicating that it might affect only a subset of RAF functions.

Experiments with KSR-deficient mice indicate that KSR is not absolutely required, but enhances

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Figure 1-10. Allosteric mechanism for activation of the kinase domain of RAF1.

RAF1 exists in an inactive conformation characterized by the monomeric state of its kinase

domain. The N- and C-terminal lobes of RAF or KSR kinase domains are indicated. Following

RAS activation, the kinase domain of RAF1 transitions from an inactive monomeric state to an

active side-to-side dimer, characterized by the juxtaposition of a critical helix (helix αC) in the

N-lobe. The cellular pool of RAF/KSR proteins contains multiple isoforms, and the mechanism

by which the cell selectively drives dimer formation between two given isoforms is currently

unknown.

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Figure 1-11. Scaffolding proteins in RAF-MEK-ERK signaling.

Scaffolding proteins form RAF-MEK-ERK signaling platforms at different subcellular

localizations. See text for details.

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signaling from RAS (314). Recent studies have shown that KSR is a functional protein kinase,

presumably stimulated allosterically by forming side-to-side dimers with RAF, and catalyzes

phosphorylation of MEK1 on non-activation segment residues to facilitate RAF-mediated

phosphorylation of MEK1 activation segment serines (315, 316).

Another group of RAF scaffolds is the connector enhancer of KSR (CNK) family of

proteins, which lack kinase activity but contain different protein-protein interaction domains that

can bind a variety of proteins, including RAF (309, 317). CNK1 can augment RAF1 activation

by increasing tyrosine phosphorylation of the N-region through the recruitment of c-SRC (318).

Interestingly, CNK1 also can bind Ras association domain-containing protein 1A (RASSF1A)

and enhance apoptosis in a MST2-dependent manner, which may play a role in balancing

apoptosis and proliferation by coordinating MST2 binding to RASSF1A or RAF1 (319). Another

family of multi-domain scaffolding proteins is IQ motif containing GTPase-activating proteins

(IQGAPs), which directly interact with, and modulate the functions of, BRAF, MEK, and ERK

(320, 321). Prohibitin (PHB) facilitates the displacement of 14-3-3 from RAF1 by activated RAS,

thereby promoting plasma membrane localization and phosphorylation of RAF1 at the activating

S338 (322, 323).

Scaffolding proteins also are crucial for the localization of members of the ERK pathway

to different subcellular signaling platforms. Similar expression to FGF (SEF1), which is located

at the Golgi apparatus, is a transmembrane scaffold for MEK and ERK (324). Importantly, SEF1

only binds activated MEK and inhibits the dissociation of the MEK-ERK complex, which blocks

nuclear translocation of ERK, allowing the activation only of cytoplasmic ERK targets (324).

The β-arrestins are proposed to augment ERK activation in clathrin-coated pits by scaffolding

RAF1, MEK, and ERK (325). Similar to SEF1, β-arrestins also prevent ERK nuclear

translocation and therefore restrict RAS signaling to cytoplasmic effectors of the pathway. The

small scaffold MEK partner-1(MP1) is an obligatory heterodimer with p14, and this complex

interacts with MEK and ERK, targeting them to late endosomes (326, 327). In addition, MP1

may target MEK-ERK to high molecular weight protein complexes (328), organized by MAPK

organizer 1 (MORG1), which was identified as an interaction partner of MP1 as well as RAF1,

BRAF, MEK, and ERK (329). MP1 also forms the Ragulator complex with p14 and p18, which

is required for the recruitment and activation of mTORC1 at the lysosomal surface (330). The

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multidomain protein paxillin is a component of focal adhesions, providing a structural and

signaling link between the actin cytoskeleton and the extracellular matrix (ECM) (331). Paxillin

constitutively interacts with MEK, but also binds to activated RAF and ERK in response to

growth factors, directing activated ERK to focal adhesions (332).

1.3.3 MEK-independent functions of RAF1

Studies using conventional and conditional Raf1 knockout mice (237, 238, 333) led to the

discovery of new RAF1 effector pathways in which regulation occurs through protein-protein

interactions. For some of these pathways, RAF1 kinase activity is dispensable (Figure 1-12).

1.3.3.1 RAF1 and apoptosis

RAF1 regulates apoptosis through multiple targets. A mitochondrial pool of RAF1 has

been shown to protect cells from apoptosis (Figure 1-12). RAF1 can be targeted to the

mitochondria via its interaction with BCL2 (334). In addition, RAF1 serves as a scaffold to

recruit protein kinase C theta (PKCθ) to phosphorylate and inactivate the pro-apoptotic BCL-2

family member BAD (335, 336). Another possible mechanism is the direct interaction between

RAF1 and mitochondrial voltage-dependent anion channels (VDACs), which may be responsible

for the RAF-induced inhibition of cytochrome c release from mitochondria (337).

Raf1 ablation in mice demonstrated that Raf1 is required for survival and protection

against apoptosis (237, 238). Surprisingly, reconstituting Raf1–/– mice with a kinase-impaired

Raf1 mutant (Raf-1YY340/1FF) fully rescues the apoptotic phenotype and produces viable mice

(238). Studies have shown that several mechanisms account for the anti-apoptotic function of

Raf1 (Figure 1-12). These may operate in a tissue-specific manner, but none requires Raf1 kinase

activity. One mechanism is through the deregulation of the Rho effector kinase Rok-α, which is

up-regulated and mislocalized to the membrane in Raf1-deficient cells (229). This results in a

defect in the internalization of the Fas death receptor, which increases Fas clustering and

membrane expression, enhancing Fas sensitivity.

The other targets of RAF1 in apoptosis suppression include two pro-apoptotic kinases,

apoptosis signal-regulating kinase 1 (ASK1) (227, 338) and MST2 (228, 339) (Figure 1-12).

ASK1 is a member of the MAP kinase kinase kinase family that selectively activates JNK and

p38 to promote apoptosis induced by various cytotoxic stresses, such as reactive oxygen species

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Figure 1-12. MEK-independent RAF1 signaling pathways.

RAF1 can suppress apoptosis in a MEK-independent fashion in several ways: 1) by binding to

and inhibiting ASK1; 2) by suppressing cytochrome C release from mitochondria through

voltage-dependent anion channels (VDACs); 3) by acting as scaffold to recruit PKCθ to

phosphorylate and inactivate BAD; 4) by binding to and inhibiting the mammalian MST2

pathway; and 5) by inhibiting ROK-α-induced FAS maintenance and clustering at the cell

membrane. Inhibition of ROK-α by RAF1 binding also is required for regulating motility

through the actin cytoskeleton and for skin tumorigenesis by preventing keratinocyte

differentiation and sustaining MYC expression.

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(ROS), or by death receptors, such as the TNF-α receptor or FAS (340). RAF1 interacts with the

N-terminal regulatory fragment of ASK1 and inhibits its proapoptotic function (227). Cardiac-

specific disruption of Raf1 results in heart dilation and dysfunction with a significant increase in

cardiomyocyte apoptosis, all of which are rescued by the ablation of Ask1 (338). MST2 was

identified in a proteomics screen for RAF1-associated proteins (228). RAF1 binds to the SARAH

domain of MST2, thereby preventing the dimerization and phosphorylation of the activation loop

of MST2, independent of RAF1 kinase activity. RASSF1A, a tumor suppressor, can disrupt the

RAF1–MST2 complex and promote the assembly of a proapoptotic signaling complex consisting

of RASSF1A, MST2, LATS1, and YAP1 (339, 341). Activated YAP1 translocates into the

nucleus and interacts with p73, promoting the expression of proapoptotic BH3-only protein p53-

upregulated modulator of apoptosis (PUMA) to induce apoptosis (341). Interestingly, RAS

binding to RAF1 enables RAF1 to activate the MEK-ERK pathway and promote proliferation,

but also induces dissociation of the RAF1-MST2 complex and promotes apoptosis (342).

1.3.3.2 RAF1 regulates cell motility and differentiation through ROK-α

Conditional gene ablation of Raf1 in keratinocytes delays migration and wound healing

(230). The target of Raf1 in motility is Rok-α (Figure 1-12). Raf1 knockout fibroblasts and

keratinocytes show a contracted appearance and fail to migrate due to the hyperactivity and

incorrect localization of Rok-α to the plasma membrane. Similar to Raf1, Rok-α is regulated by

auto-inhibition, and its C-terminal regulatory region features a domain highly homologous to the

CRD in Raf1. The Raf1 regulatory domain can cross-regulate Rok-α by binding to the Rok-α

kinase domain and repressing its function in trans (343). The biological relevance of the Raf1:

Rok-α complex was observed in a Ras-induced skin tumor mouse model (344). In this model, the

inhibition of Rok-α by Raf1 was required for Ras transformation by decreasing the expression of

epidermal differentiation cluster (EDC) and activating the Stat3/Myc pathway, thereby

promoting dedifferentiation of tumor cells.

1.4 Hypertrophic cardiomyopathy and the RAS/ERK pathway

1.4.1 Cardiac hypertrophy and hypertrophic cardiomyopathy

The mammalian heart is a dynamic organ, which is composed of cardiomyocytes, non-

myocytes (e.g., fibroblasts, endothelial cells, mast cells and vascular smooth muscle cells) and

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surrounding extracellular matrix (345, 346). Cardiomyocytes are specialized muscle cells

composed of bundles of myofibrils, which contain repeating contractile units known as

sarcomeres (347). It is generally believed that most cardiomyocytes lose their ability to

proliferate at or soon after birth, and that the subsequent growth of heart occurs primarily as a

result of an increase in myocyte size (348). In adults, the growth of heart is closely matched to its

functional load. In response to elevated workload, the heart undergoes a hypertrophic response,

which is characterized by an increase in the size of individual cardiomyocytes, to compensate for

the increase in wall stress (345). This hypertrophic response can be classified as one of two

general types, physiological or pathological, which are caused by different stimuli, are

functionally distinguishable, and are associated with distinct structural and molecular phenotypes

(Figure 1-13) (349, 350).

Physiological growth of the heart includes the embryonic and fetal stages of development

and the rapid postnatal growth stage (345). In the adult, physiological hypertrophy usually occurs

in response to chronic exercise training (351) or pregnancy (352). Pathological cardiac

hypertrophy is caused by genetic mutations (primary) (353, 354) or in response to diverse

stimuli, such as hypertension, valvular stenosis or myocardial infarction (secondary) (345, 355).

With increased cardiac stress, pathological cardiac hypertrophy might initially represent a

compensatory mechanism for increasing cardiac function and decreasing ventricular wall

tension. However, chronic pathological hypertrophy eventually leads to functional

decompensation, and predisposes individuals to ventricular dilatation, heart failure, arrhythmia

and/or sudden death (356, 357). By contrast, physiological hypertrophy does not decompensate

into dilated cardiomyopathy or heart failure (351, 358).

Physiological and pathological hypertrophy can be further sub-classified as concentric

(hearts with thick walls and relatively small cavities) or eccentric (hearts with chamber

enlargement and a proportional change in wall thickness) (359, 360). Isotonic exercise (e.g.,

running, walking or swimming) causes volume overload and produces eccentric physiological

hypertrophy, whereas isometric or static exercise (e.g., weight lifting) causes pressure overload

and results in concentric physiological hypertrophy (345, 358). Pathological stimuli causing

pressure overload, such as hypertension or aortic stenosis, produce concentric hypertrophy. By

contrast, volume overload (e.g., mitral or aortic regurgitation) that causes increased diastolic wall

stress or myocardial infarction (MI), can result in eccentric hypertrophy (359, 360). Concentric

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Figure 1-13. Types of cardiac hypertrophy.

RV, right ventricle; LV, left ventricle.

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hypertrophy is characterized by assembly of sarcomeres in parallel, resulting in a relative

increase in the width of individual cardiomyocytes. By contrast, eccentric hypertrophy is

characterized by assembly of sarcomeres in series, leading to a relatively greater increase in the

length than in the width of cardiomyocytes. As mentioned above, concentric pathological

hypertrophy may progress to eccentric hypertrophy, and then eventually to cardiac dilation with

associated systolic heart failure (361).

Animal studies have demonstrated that physiological and pathological hypertrophy are

associated with distinct histological and molecular characteristics (362-364). In mice, a surgical

model of pathological hypertrophy can be established by transverse aortic constriction (TAC),

which causes pressure overload on the left ventricle (365, 366). TAC initially leads to

compensated hypertrophy, which often is associated with a temporary increase of cardiac

contractility. However, over time, the response to the chronic hemodynamic overload becomes

maladaptive, resulting in cardiac dilatation and heart failure. Pathological hypertrophy usually is

associated with increased apoptosis, necrosis and interstitial fibrosis (367). In response to

pathological stimuli, cardiac fibroblasts and extracellular matrix proteins accumulate

disproportionately and excessively surrounding the cardiomyocytes, which can lead to

mechanical stiffness, and contribute to cardiac dysfunction (368). Pathological cardiac

hypertrophy typically is associated with reactivation of fetal genes, including atrial natriuretic

peptide (ANP), B-type natriuretic peptide (BNP) and genes for the fetal isoforms of contractile

proteins, such as skeletal α-actin and β-myosin heavy chain (β-MHC). These changes can be

accompanied by down-regulation of genes normally expressed at higher levels in the adult, such

as α-MHC and sarcoplasmic reticulum Ca2+-ATPase (SERCA2a) (369). This altered gene

expression program affects cardiac function, leading to altered contractility, elevated end-

diastolic pressure and arrhythmias. By contrast, these histological and molecular changes are not

observed in models of physiological cardiac hypertrophy induced by exercise training (364, 370,

371).

Primary hypertrophic cardiomyopathy (HCM) is the prototypical genetic form of

pathological hypertrophy. The hallmark of HCM is cardiac hypertrophy in the absence of an

obvious inciting hypertrophic stimulus (372). Complications of HCM include left ventricular

outflow tract obstruction, arrhythmias, diastolic dysfunction, MI, stroke, infective endocarditis

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and sudden cardiac death (373). Hearts from HCM patients commonly show cardiomegaly and

left ventricular hypertrophy (LVH). In end-stage disease, cardiac remodeling occurs, leading to

dilatation of the ventricles and atria and thinning of the ventricular wall due to myocardial

scarring (374). Myocyte disarray, the loss of the normal parallel alignment of cardiomyocytes, is

the most characteristic and common histological finding in HCM (373). Also, HCM usually is

associated with interstitial myocardial fibrosis and variation in the size of myocyte nuclei.

Autosomal dominant mutations in several genes encoding proteins of the cardiac

sarcomeric apparatus (e,g., β-MHC, cardiac troponin T, and myosin-binding protein C) account

for 60-70% of all cases of primary HCM (373). Such mutations usually alter sarcomere structure

and function and result in mechanical, biochemical and/or bioenergetic stresses that activate

cardiomyocyte signaling pathways to mediate the hypertrophic phenotype (375-378). Non-

sarcomeric genes responsible for myocardial metabolism also have been identified in HCM, such

as lysosome-associated membrane protein-2-α-galactosidase (LAMP2) (379). Aberrant

activation of hypertrophic signaling pathways can themselves result in hypertrophy. For

example, germ line mutations in adenosine monophosphate-activated protein kinase (AMPK) are

a rare cause of HCM (380-382).

Moreover, the discovery of mutated RAS/ERK pathway-related genes in RASopathies

has underlined the relevance of this signaling pathway to HCM (32). HCM occurs with a high

incidence in CS (200, 383) and LS (176, 384), whereas it is less common in CFCS (171, 385)

and NS (146, 173) patients. However, NS-associated kinase-activating RAF1 mutants are highly

associated with HCM (148, 149).

1.4.2 Signaling pathways involved in cardiac hypertrophy

Studies of transgenic and knockout mice, in combination with surgical and exercise

models, have revealed signaling cascades that play important roles in regulating cardiac

hypertrophy, such as the GPCR signaling, the calcineurin-nuclear factor of activated T cells

(NFAT) pathway, the insulin-like growth factor-I (IGF-I)-phosphoinositide-3 kinase (PI3K)

cascade, and the MAPK pathway (349, 350, 386) (Figure 1-14). To date, the best characterized

examples of pathways that play distinct roles in pathological and physiological hypertrophy are

Gαq/α11- dependent and IGF-I signaling, respectively.

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The secretion of cardiac paracrine and/or autocrine factors, including angiotensin-II

(Ang-II), endothelin-1 (ET-1) and catecholamines (α-adrenergic), in response to a pathological

stimulus plays an important role in the development of pathological cardiac hypertrophy (387-

391) (Figure 1-14). These ligands bind to specific GPCRs that are coupled to heterotrimeric G

proteins of the Gαq/α11 subclass, which leads to the activation of downstream signaling molecules,

such as protein kinase C (PKC), and the mobilization of internal Ca2+. Ca2+ stores have been

shown to regulate cardiac hypertrophy by activating the calcineurin-NFAT pathway (392, 393)

and by promoting the calmodulin-dependent kinase (CaMK)-mediated nuclear export of histone

deacetylases (HDACs) (394). Calcineurin, which is a Ca2+-dependent serine/threonine protein

phosphatase, binds to and dephosphorylates transcription factors of the NFAT family, resulting

in their nuclear translocation and the activation of pro-hypertrophic gene expression (392). The

class II HDACs are implicated in cardiac hypertrophy by means of chromatin remodelling and

altered gene expression. Nuclear export of class II HDACs triggered by CaMKII-mediated

phosphorylation releases the suppression of hypertrophic gene transcription and induces cardiac

hypertrophy (394). Activation of Gαq/α11 also can induce MAPK pathways in cardiomyocytes,

although the exact mechanism remains unknown (395).

IGF-I has been implicated in the regulation of developmental and physiological growth of

the heart, and is released in response to exercise training (396-400) (Figure 1-14). IGF-I acts via

the IGF-I receptor, a receptor tyrosine kinase that activates class IA PI3K (PI3Kα). PI3K

activation results in the sarcolemmal recruitment and activation of the kinase AKT (also known

as PKB) (401). Glycogen synthase kinase-3β (GSK-3β), which is inhibited by AKT-mediated

phosphorylation, is an important downstream target of AKT in the heart. GSK-3β negatively

regulates hypertrophic transcriptional effectors (e.g., GATA4, β-catenin, c-Myc and NFAT), and

inhibits the translation initiation factor elF2B. AKT also enhances protein synthesis and cell

growth by activating the mTOR pathway (402). However, the role of the PI3K-AKT signaling

pathway in promoting cardiac hypertrophy is complex, as each of the effectors in this pathway

can have adaptive or maladaptive influences on the myocardium, depending on the type and

duration of the stimuli (350, 354). For example, short-term activation of AKT in cardiomyocytes

results in physiological adaptive hypertrophy, whereas chronic activation produces pathological

hypertrophy (403). Excessive PI3K-AKT-mTOR signaling also has been recently associated

with pathological cardiac hypertrophy in LS, wherein rapamycin treatment reverses LS cardiac

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Figure 1-14. Signaling pathways involved in cardiac hypertrophy.

ANP, atrial natriuretic peptide; Ang II, angiotensin II; BNP, B-type natriuretic peptide; CaMK,

calmodulin-dependent kinase; CDK, cyclin dependent kinase; DAG, diacylglycerol; Endo-1,

endothelin-1; GC-A, guanyl cyclase-A; GPCR, G-protein coupled receptors; GSK3β, glycogen

synthase kinase-3β; HDAC, histone deacetylases; IκB, inhibitor of NF-κB; IGF-I, insulin-like

growth factor-I; IKK, inhibitor of NF-κB kinase; MEF, myocyte enhancer factor; mTOR,

mammalian target of rapamycin; NFAT, nuclear factor of activated T cells; NF-κB, nuclear

factor-κB; NIK, NF-κB-inducing kinase; PDK, phosphoinositide-dependent kinase; PI3K,

phosphatidylinositol 3-kinase; PKB, protein kinase B; PKC, protein kinase C; PKD, protein

kinase D; PLA2, phospholipase A2; PLC, phospholipase C; Pol II, RNA polymerase II; TAK,

TGFβ-activated kinase; TGFβ, transforming growth factor-β; TNFα, tumour necrosis factor-α.

Adopted from Heineke and Molkentin, Nature Reviews Molecular Cell Biology 7, 589-600 (350).

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defects (404).

MAPK signaling is initiated in cardiomocytes by GPCRs (395), RTKs (e.g. IGF-I, EGF

and FGF receptors) (405), receptor serine/threonine kinases (e.g. transforming growth factor-β1

(TGF- β1) receptor) (406), glycoprotein-130 (gp130), the receptor for the interleukin-6 (IL-6)-

related cytokines (407), and by stress stimuli, such as mechanical stretch (408) (Figure 1-14).

Activated MAPKs each phosphorylate numerous downstream targets, including transcription

factors that induce the reprogramming of cardiac gene expression.

The role of the RAS/ERK pathway in cardiac hypertrophy has been controversial. Some

data argue that excessive activity of this pathway causes HCM, whereas other evidence suggests

its involvement in physiological, but not pathological, cardiac hypertrophy (409, 410).

Transgenic mice with cardiac-specific expression of oncogenic HRas (G12V) display significant

cardiac hypertrophy, decreased contractility, diastolic dysfunction associated with interstitial

fibrosis, induction of cardiac fetal genes and sudden death (411-413), all of which are consistent

with HCM. However, besides RAF-MEK-ERK activation, RAS also can activate the JNK

branch of the MAPK cascade, PI3K and other signaling pathways. Therefore, the cardiac defects

observed with constitutive RAS activation could be independent of MEK-ERK signaling. In

cultured cardiomyocytes, depletion of Erk1/2 with antisense oligonucleotides, or

pharmacological inhibition of Mek1/2, attenuates the hypertrophic response to agonist

stimulation (414, 415). Mice with cardiac-specific over-expression of “dominant-negative” Raf1

show reduced phosphorylation of the Erk target Elk1 and have no overt phenotype, but they are

resistant to the development of cardiac hypertrophy in response to pressure overload (416).

These data suggest that signals from Raf1 are necessary for promoting the pathological

hypertrophic response. The RAS/ERK pathway also has been implicated in inducing the re-

expression of fetal cardiac genes by altering the levels and activities of cardiac transcription

factors, such as GATA4, MEF2 and NFAT (417).

On the other hand, transgenic mice expressing an activated Mek1 allele under the control

of the α-MHC promoter have concentric hypertrophy with enhanced contractility, show no signs

of decompensation over time and reportedly do not progress to pathological hypertrophy up to 6

months of age (418). This study also found that the Mek1-Erk1/2 pathway induces cardiac

hypertrophy partially by enhancing the transcriptional activity of Nfat, indicating crosstalk

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between the Ras/Erk and the calcineurin-Nfat pathways (419). Another study argued against any

role for Erk1/2 in cardiac hypertrophy, as Erk1-/-Erk2+/- mice, as well as transgenic mice with

cardiac-specific expression of dual specificity phosphatase 6 (Dusp6), an Erk1/2-specific

phosphatase, showed a normal hypertrophic response to pressure overload and exercise (420). A

more recent study from the same group using mice lacking all Erk1/2 proteins in the heart

demonstrated that Erk1 and Erk2 regulate the balance between eccentric and concentric cardiac

growth (421). Although loss of Erk1/2 in the heart (Nkx2.5-Cre) or specifically in the

cardiomyocytes (α-MHC-Cre) does not block the cardiac hypertrophic response to either aging

or pathological stimuli, it does dramatically alter how the heart grows: adult cardiomyocytes

from the Erk1-/-Erk2fl/fl:Cre heart show eccentric growth with a significant increase in length,

whereas myocytes from activated Mek1 transgenic hearts show concentric growth with increased

width.

The MEK5-ERK5 branch of MAPKs pathways also has been identified as a regulator of

cardiac growth. Transgenic mice with cardiac-specific expression of activated Mek5 show

eccentric cardiac hypertrophy, and progress to dilated cardiomyopathy and sudden death (422).

Consistent with this, targeted deletion of Erk5 in the heart attenuates the hypertrophic response

following pressure overload, and induces apoptosis in the heart (423). By contrast, activation of

p38 (transgenic mice expressing activated Mek3 or Mek6) (424) or JNK (transgenic mice

expressing activated Mek7) (425, 426) in the heart does not induce cardiac hypertrophy. Such

mice develop lethal cardiomyopathy as juveniles, characterized by reduced functional

performance, fibrosis and dilated ventricular walls.

Finally, in addition to signaling events that are initiated by ligand-receptor interactions,

cardiomyocytes can directly sense biomechanical deformation or stretch through an internal

sensory apparatus (427, 428). The integrin-interacting molecule melusin has been implicated as a

mechanical stress sensor in cardiomyocytes, essential for the inactivation of GSK-3β (428). The

muscle LIM-domain protein (MLP) might be a second sensory apparatus at the level of the Z-

disc within sarcomeres (427). MLP anchors to specific proteins at the Z-disc and functions as an

internal stretch sensor through a complex of transducing proteins that regulate calcineurin-NFAT

signaling (429).

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1.5 Rationale and hypothesis

The RAS/ERK pathway controls fundamental cellular processes, such as proliferation,

differentiation and apoptosis, so it is not surprising that malfunction of this pathway results in the

multiple clinical manifestations seen in RASopathies. Yet, how mutations in the same signaling

pathway cause similar, yet clearly distinct, phenotypes remains unclear. Consequently, detailed

understanding of RASopathy pathogenesis could yield new insights into RAS/ERK pathway

regulation.

Primary HCM is the most common inherited cardiovascular disorder (1 in 500

individuals in the general population) (373, 430) and a leading cause of sudden death in the

young (431). Genetic and cellular models have identified multiple signaling pathways that can

cause or contribute to pathological hypertrophy, but the detailed mechanism by which aberrant

activation of these pathways evokes HCM remains incompletely understood. Delineating the

molecular pathways that distinguish physiological and pathological cardiac hypertrophy, and

identifying ways to reverse the latter, are of obvious medical importance. The role of the

RAS/ERK pathway in cardiac hypertrophy has been controversial, and remains to be resolved.

Given the fact that HCM is found in nearly all (~95%) NS patients bearing RAF1 mutations that

cause increased kinase activity (148, 149), this disorder provides a good opportunity to study the

role of RAS/ERK pathway in the pathogenesis of HCM.

The hypotheses of my project were: (1) Specific RAF1 mutations identified in NS

patients cause the common features of NS; (2) Kinase-activating RAF1 mutations cause NS with

HCM, whereas kinase-impaired RAF1 mutations are not associated with HCM; (3) Different

RAF1 mutations cause NS through distinct molecular mechanism(s), and/or by acting in different

cell types. To test my hypotheses, I generated and analyzed lines of knock-in mice with HCM-

associated kinase-activating (L613V) or non-HCM-associated kinase-impaired (D486N) Raf1

mutations, respectively. I also investigated the effects and biochemical properties of various

other NS-associated RAF1 mutants expressed at more physiological levels than in previous

transient transfection studies. Together, my project aimed to advance the field in understanding

the molecular pathogenesis of NS as well as HCM, which could provide us with potentially

important prognostic, diagnostic and even therapeutic implications for this disease. My findings

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could also increase our understanding of other forms of cardiac disease, as well as normal

cardiac development.

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Chapter 2

Materials and methods

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2.1 Mice

All animal studies were approved by the University Health Network Animal Care

Committee (Toronto, ON, Canada) and performed in accordance with the standards of the

Canadian Council on Animal Care.

2.1.1 Generation of Raf1L613V knock-in mice

To construct the targeting vector for inducible Raf1L613V knock-in mice, a “short arm”,

containing Raf1 exon 12 (SacII-NotI genomic fragment) and a “long arm”, which includes exons

13-16 (BamHI-ClaI genomic fragment) were ligated into the vector pGK Neo-HSV-1 TK (432).

The L613V (exon 16) mutation, marked by a unique DraIII site, was introduced by site-directed

mutagenesis. A splice acceptor sequence, a Raf1 cDNA fragment encoding wild-type exons 13-

16 and a pGK-Neo (Neo) gene were positioned after the first loxP site as a SalI-XbaI fragment.

The targeting vector was linearized with SacII and electroporated into G4 ES cells (129S6 x

C57BL/6 F1 background). Genomic DNA, isolated from doubly G418/1-(2-deoxy-2-fluoro-β-D-

arabinofuranosyl)-5 iodouracil (FIAU)-resistant (positive and negative selection, respectively)

ES clones, was screened by PCR using primers outside and inside the targeting vector (Table

2-1), followed by NotI digestion, which marks the targeting vector. Homologous recombinants

were confirmed by Southern blotting using Neo and external (5’ and 3’) probes (Table 2-1). For

these experiments, genomic DNA was digested with XbaI (5’ and Neo probes) or BamHI (3’

probe).

To validate the desired properties of the targeted locus, correctly targeted ES cells were

transfected with a Cre-expressing plasmid (MSCV-GFP-Cre) to excise the cDNA-Neo cassette

(see below for detailed methods). Expression of Raf1L613V mRNA was confirmed by RT-PCR

(Table 2-1), followed by digestion with DraIII, which marks the L613V allele. Chimeras were

generated by outbred morula aggregation (Toronto Centre of Phenogenomics), and germ line

transmission was obtained (L613Vfl/+ mice). L613Vfl/+ mice (129Sv X C57BL/B6) were

crossed to CMV-Cre (C57BL/B6) mice, which express Cre ubiquitously, and then to WT

(129S6) mice to generate mice with global Raf1L613V expression (L613V/+, 129Sv X

C57BL/B6). Mice on a 129Sv X C57BL/B6 mixed background were used for all experiments.

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PCR Screening

sense 5’-TCCAGCTAATTGACATTGCCCGACAGACAGCTCAG-3’

antisense 5’-GAACGGGTTGTCATCCTGCATCCGGATTACTTCTG-3’

Neo probe sense 5’-GGATTGCACGCAGGTTCTCCG-3’

antisense 5’-CGCCGCCAAGCTCTTCAGCAA-3’

5’ probe sense 5’-TGCTCTGGAGCTCAAACCCTCAGTGTAG-3’

antisense 5’-CATGGCTGAGTGGACGGTCAGGCTG-3’

3’ probe sense 5’-GAGACGGCAGATCCTCAGTAGTACTTG-3’

antisense 5’-ACGGTGGTAGTTGTGTCTTTGGCCATG-3’

RT-PCR for Raf1 mRNA

sense 5’-TCTCCATGAAGGCCTCACGGTG-3’

antisense 5’-AGACTGGTAGCCTTGGGGATGTAG-3’

Genotyping for Raf1L613V

sense 5’-ATCCCCTGATCTCAGCAGGCTCTAC-3’

antisense 5’-AGTAGTCTAGGTCCTTAGCAGCAGC-3’

Table 2-1. PCR primers for generating the Raf1L613V mice.

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For genotyping, genomic DNA was prepared from tails, and then subjected to PCR (Table 2-1)

and digestion with DraIII.

2.1.2 Generation of Raf1D486N knock-in mice

To construct the targeting vector for our inducible Raf1D486N knock-in mice, a “short

arm”, containing Raf1 exon 12 (SacII-NotI genomic fragment), and a “long arm,” which includes

exons 13-16 (BamHI-ClaI genomic fragment), were ligated into the vector pGK Neo-HSV-1 TK

(432). The D486N (exon 13) mutation, marked by a unique ApoI site, was introduced by site-

directed mutagenesis. A splice acceptor sequence, a Raf1 cDNA fragment encoding wild-type

exons 13-16, and a pGK-Neo (Neo) gene were positioned after the first loxP site as a SalI-XbaI

fragment. The targeting vector was linearized with SacII and electroporated into G4 ES cells

(129Sv x C57BL/6 F1 background). Genomic DNA, isolated from doubly G418/FIAU-resistant

ES clones, was screened by PCR using primers outside and inside the targeting vector (Table

2-2). Homologous recombinants were confirmed by Southern blotting, using Neo and external

(5’ and 3’) probes (Table 2-2). For these experiments, genomic DNA was digested with XbaI (5’

and Neo probes) or BamHI (3’ probe).

To validate the desired properties of the targeted locus, correctly targeted ES cells were

transfected with a Cre-expressing plasmid (pMSCV-GFP-Cre) to excise the cDNA-Neo cassette

(see below for detailed methods). Expression of Raf1D486N mRNA was confirmed by RT-PCR

(Table 2-2) followed by digestion with ApoI, which marks the D486N allele. Chimeras were

generated by outbred morula aggregation (Toronto Centre of Phenogenomics), and germ line

transmission was obtained (D486Nfl/+ mice). D486Nfl/+ mice (129Sv X C57BL/6 background)

were crossed to EIIa-Cre (129Sv) mice, which express Cre ubiquitously, and then to WT

(C57BL/6) mice to generate mice with global Raf1D486N expression (D486N/+, 129Sv X

C57BL/6). Mice on a 129Sv X C57BL/6 mixed background were used for all experiments. For

genotyping, genomic DNA was prepared from tails, and then subjected to PCR (Table 2-2) and

digestion with ApoI.

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PCR screening

for 5’loxP site

sense 5’-TCCAGCTAATTGACATTGCCCGACAGACAGCTCAG-3’

antisense 5’-GAACGGGTTGTCATCCTGCATCCGGATTACTTCTG-3’

PCR screening

and probe for

sense 5’-GGA TTG CAC GCA GGT TCT CCG-3’

antisense 5’-CGC CGC CAA GCT CTT CAG CAA-3’

5’ probe

sense 5’-TGC TCT GGA GCT CAA ACC CTC AGT GTA G -3’

antisense 5’-CAT GGC TGA GTG GAC GGT CAG GCT G-3’

3’ probe

sense 5’-GAG ACG GCA GAT CCT CAG TAG TAC TTG-3’

antisense 5’-ACG GTG GTA GTT GTG TCT TTG GCC ATG-3’

RT-PCR for

Raf1 mRNA

sense 5’-TCT CCA TGA AGG CCT CAC GGTG-3’

antisense 5’-AGA CTG GTA GCC TTG GGG ATG TAG-3’

PCR screening

& genotyping

for Raf1D486N

sense 5’-TGTGCGCATGCCATCGTTCCCTGTC -3’

antisense 5’-GCACCCTACTCTGGCCCAGTAATTC -3’

Table 2-2. PCR primers for generating the Raf1D486N mice.

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2.2 Cell culture

2.2.1 Mouse embryonic stem (ES) cells

ES cells (G4) were cultured on γ-irradiated mouse embryonic fibloblast (MEF) feeders in

knockout Dulbecco's modified Eagle's medium (DMEM) (Invitrogen), containing 15% ES-tested

(HyClone, Thermo Scientific) fetal bovine serum (FBS), 2mM L-glutamine (Invitrogen), 0.1mM

Non-Essential Amino Acids (NEAA) (Invitrogen), 0.1mM β-mercaptoethanol (Sigma), 100

U/ml penicillin/streptomycin (Invitrogen), and 500U/ml LIF (ESGRO, Chemicon). ES cells

were transfected with MSCV-GFP-Cre plasmid using Lipofectamine 2000 reagent (Invitrogen),

according to the manufacturer’s protocol. GFP-positive cells were purified by Fluorescence

Activated Cell Sorting (FACS) at 48 hours post-transfection, and then used for RNA isolation.

For biochemical studies, ES cells were cultured under feeder-free condition, starved in

knockout DMEM containing 1% FBS for 6 hr and then stimulated with 103U/ml LIF before

harvesting.

2.2.2 Mouse embryonic fibroblasts (MEFs)

Primary MEFs were prepared from E13.5 embryos as described (433). In brief, embryos

were incubated in 0.25% trypsin/EDTA (Invitrogen) for 30 min at 37°C, and dissociated cells

were collected by centrifugation and cultured in DMEM (WISENT INC.) containing 10% FBS

and 100 units/ml penicillin/streptomycin (Invitrogen). Different independent MEF strains were

used for experiments, with similar results.

Primary MEFs were immortalized by the 3T3 protocol (434). Immortalized MEFs were

cultured in DMEM containing 10% FBS and 100units/ml penicillin/streptomycin.

For biochemical studies, MEFs were starved in serum-free DMEM for 16 hr before

stimulation with 10ng/ml EGF or 50ng/ml PDGF (both from PeproTech) before harvesting.

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2.2.3 Neonatal cardiomyocytes and cardiac fibroblasts

Neonatal mouse ventricular myocytes (neonatal cardiomyocytes) were isolated using

methods adapted from a previous study (435). In brief, 1-day-old mouse hearts were harvested

and pre-digested with 0.15 mg/mL trypsin (Invitrogen) at 4°C for 12-16 hours, followed by

50U/ml Type II Collagenase (Worthington Biochemical) and 0.2 mg/mL trypsin in calcium- and

bicarbonate-free Hank’s buffer with HEPES (pH 7.5) (137mM NaCl, 5.36mM KCl, 0.81mM

MgSO4, 5.55 mM Dextrose, 0.44mM KH2PO4, 0.34mM NaH2PO4·H2O and 20mM HEPES) for

1-2 hr at 37°C. Non-cardiomyocytes were depleted by differential plating for 1 hour.

Cardiomyocytes were counted, seeded at 2.5 × 105 cells/ml on Falcon Primaria™ tissue-culture

plates (BD Biosciences), and cultured at 37°C in DMEM/Ham's F-12 [1:1 (v/v); Invitrogen],

10% FBS, and 100 U/ml penicillin/streptomycin (Invitrogen), supplemented with 0.1 mM

bromodeoxyuridine (Sigma-Aldrich) and 20 μM arabinosylcytosine (Sigma-Aldrich) to inhibit

rapidly proliferating cells. For biochemical studies, the medium was replaced with serum-free

DMEM/Ham’s F12 (1:1) medium supplemented with 1% insulin-transferrin-selenium

supplements-X (Invitrogen) after 24 hr. After an additional 24 hr, cardiomyocytes were

stimulated with 10ng/ml IL-6 (PreproTech), 1µg/ml Angiotensin II (Sigma-Aldrich), 100ng/ml

IGF-I (PreproTech), 50ng/ml EGF (PreproTech) or 100ng/ml NRG (heregulin β1, PeproTech)

before harvesting.

Non-cardiomyocytes from the above preparation, mainly comprising cardiac fibroblasts,

were cultured in DMEM containing 10% FBS and 100 units/ml penicillin/streptomycin. For

biochemical studies, cardiac fibroblasts were starved in serum-free DMEM for 16 hr, and then

stimulated with EGF (50 ng/ml), IGF-I (100ng/ml), PDGF (100ng/ml), or FGF2 (100ng/ml), all

from PeproTech, before harvesting.

2.2.4 Flp-In T-REx 293 cell lines

Flp-In T-REx 293 cell lines were cultured in DMEM (WISENT INC.) containing 10%

tetracycline-tested GIBCO® FBS (Invitrogen) and 100units/ml penicillin/streptomycin

(Invitrogen). To induce expression of the indicated gene, a final concentration of 1µg/ml

tetracycline (Sigma-Aldrich) was added to the cells, followed by incubation for 24 hours before

harvesting for analysis.

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For biochemical studies, Flp-In T-REx 293 cell lines were starved in serum-free DMEM

for 16 hr before stimulation with 50ng/ml EGF before harvesting.

2.3 Generation of Flp-In T-REx 293 expression cell lines

RAF1 mutations were introduced into a Flag-tagged human RAF1 construct (a gift from

Dr. Bruce Gelb, Mt Sinai Hospital, NY, NY) by site-directed mutagenesis. Flag-tagged human

WT or mutant RAF1 coding sequences were cloned into the KpnI and XhoI restriction sites of

the vector pcDNA5/FRT/TO (Invitrogen). Flp-In T-Rex 293 host cells (Invitrogen) were co-

transfected with pOG44 (Invitrogen) and pcDNA5/FRT/TO expression plasmid DNAs, using

FuGENE HD Transfection Reagent (Promega), according to the manufacturer’s protocol.

Hygromycin-resistant colonies were picked and expanded to assay for tetracycline-regulated

expression of Flag-RAF1.

2.4 Inducible RAF1/BRAF heterodimerization

Inducible RAF1/BRAF heterodimerization was achieved by using the ARGENT

regulated heterodimerization kit (ARIAD). A Flag-tagged human RAF1R401H/D486N cDNA was

cloned into the EcoRI and XbaI restriction sites of the vector pC4EN-F1. A human BRAF cDNA

was cloned into the XbaI restriction site of the vector pC4-RHE. The resultant FKBP and FRB

fusion protein constructs were co-transfected into Flp-In T-REx 293 host cells, using FuGENE

HD Transfection Reagent (Promega), as above.

2.5 Lentivirus production and transduction

Lentiviral shRNA expression plasmids (hairpin-pLKO.1) were obtained from Dr. Jason

Moffat (University of Toronto). shRNA oligonucleotide sequences against murine Braf (5’

CCACATCATTGAGACCAAATT 3’ or 5’ CGAGGATACCTATCTCCAGAT 3’), murine Araf

(5’ CAGGCTCATCAAAGGAAGAAA 3’) and a control shRNA against Luciferase (5’

CAAATCACAGAATCGTCGTAT 3’) were used. To generate lentiviruses, 2X106 293T

packaging cells in growth medium (DMEM+10%FBS) were transfected with 3µg lentiviral

shRNA construct, 2.7 µg packaging plasmid (pCMV-dR8.91) and 0.3 µg envelope plasmid

(VSV-G) in 10cm cell culture plates, using FuGENE HD Transfection Reagent (Promega)

according to the manufacturer’s protocol. Transfection medium was changed the following

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morning and replaced with 10ml high serum growth medium (DMEM+30%FBS) for viral

harvest. Virus-containing supernatants were collected at 48h post-transfection, and subsequently

passed through a 0.45µm filter to remove cell debris, aliquotted, and stored at -80°C.

For knock-down experiments, cells were transduced in 10cm cell culture plates with 1ml

virus in the presence of 6µg/ml polybrene for 24 hours, followed by selection with 2.5µg/ml

puromycin for 48 hours. After puromycin selection, transduced cells were re-plated for

biochemical analysis.

2.6 Retrovirus production and transduction

A Myc epitope tag was added to the N-terminus of human BRAF cDNA (The Centre for

Applied Genomics) by using PCR. The BRAFR509H mutation was introduced by site-directed

mutagenesis. RNAi-insensitive BRAFWT and BRAFR509H mutants were generated by introducing

three silent mutations into the shRNA target sequence in BRAF by site-directed mutagenesis.

Blunt-ended Myc-BRAFWT or Myc-BRAFR509H coding sequence was cloned into the blunt-ended

XhoI site of the pMSCV-IRES-EGFP vector upstream of the IRES sequence.

To generate retroviruses, 1X106 293T packaging cells in growth medium

(DMEM+10%FBS) in 6cm cell culture plates were transfected with 3µg pMSCV retroviral and 3

µg EcoPac packaging plasmid using FuGENE HD Transfection Reagent (Promega), according to

the manufacturer’s protocol. Transfection media was changed the following morning and

replaced with 5ml fresh growth medium for viral harvest. Virus-containing supernatants were

collected at 48h post-transfection, and passed through a 0.45µm filter to remove cell debris.

MEFs were transduced in 15cm cell culture plates with 5ml virus in the presence of 6µg/ml

polybrene for 24 hours, followed by fluorescence-activated cell sorting (FACS) to select for GFP

positive cells.

2.7 Biochemical analysis

Total protein extracts from cells or tissues were prepared by homogenization in RIPA

buffer (50mM Tris-HCl, pH7.5, 150mM NaCl, 2mM EDTA, 1% NP40, 0.5% Na deoxycholate,

0.1% SDS) containing a protease and phosphatase inhibitor cocktail (40 g/ml PMSF, 20mM

NaF, 1mM Na3VO4, 10mM β-glycerophosphate, 10mM sodium pyrophosphate, 2g/ml antipain,

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2g/ml pepstatinA, 20g/ml leupeptin, and 20g/ml aprotinin). Homogenates were centrifuged

at 16,100 x g for 15 min at 4 °C, and the supernatants were collected. Lysates (10-25g protein)

were resolved by SDS-PAGE and analyzed by immunoblotting.

For immunoprecipitations, total cell extracts were prepared in NP40 buffer (20mM Tris-

HCl, pH8.0, 137mM NaCl, 2mM EDTA, 1% NP40 and 10% glycerol) containing the protease

and phosphatase inhibitor cocktail described above. Homogenates were centrifuged at 16,100 x g

for 15 min at 4 °C, and the supernatants were collected. Lysates were incubated with anti-Raf1

antibody (BD Biosciences) and Protein-G Sepharose 4 Fast Flow (GE Healthcare), or anti-FLAG

M2 affinity agarose gel (Sigma-Aldrich) for 3 hours at 4 °C with rotation. Beads were washed

four times with NP40 buffer, and immunoprecipitates were analyzed by immunoblotting.

Antibodies for immunoblots included: Raf1 (BD Biosciences), SH-PTP2 (C-18) and

ERK2 (D2) (Santa Cruz Biotechnology Inc.), and phospho-MEK1/2, MEK1/2, phospho-p44/42

MAPK, phospho-S6 (Ser235/236), p38, phospho-p38, phospho-JNK1/2, Akt1, phospho-Akt

(Ser473), phospho-GSK3β (Ser9), phospho-P70S6K (Cell Signaling Technology). Primary

antibody binding was visualized by IRDye infrared secondary antibodies using the Odyssey

Infrared Imaging System (LI-COR Biosciences). Quantification of immunoblots was performed

by using Odyssey V3.0 software.

2.8 Body size analysis and morphometry

For growth curves, body length [anal-nasal (AN) length] and weight were measured

weekly. For skeletal morphometry, mice were anesthetized with 2% isoflurane and scanned by

using a Locus Ultra microCT scanner (GE Healthcare). Three-dimensional images of the

skeleton were generated and analyzed with GEHC MicroView software (GE Healthcare). Skull

measurements were made according to the Standard Protocol and Procedures from the Jackson

Laboratory (http://craniofacial.jax.org/standard_protocols.html).

2.9 Histology and immunohistochemistry

Hearts for morphometry and histochemistry were fixed in the relaxed state by infusion of

1% KCl in PBS, followed by 10% buffered formalin. Hearts were then incubated in 10%

buffered formalin overnight and embedded in paraffin. Sections (5µm) were prepared and

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stained with H&E, Picro sirius red (PSR) or Masson-Trichrome. Cell membranes were stained

with TRITC-conjugated wheat germ agglutinin (WGA) (Sigma-Aldrich), according to the

manufacturer’s protocol. Nuclei were stained with DAPI. Cross-sectional area of cardiomyocytes

with centrally located nuclei (to ensure the same plane of sectioning) was measured by using

ImageJ. Five individual samples were analyzed for each genotype, with 200 cells measured in

each.

2.10 BrdU incorporation assays

For proliferation assays, pregnant mice (E16.5) were injected intraperitoneally (IP) with

BrdU (100 μg/g body weight) 1 h before sacrifice. Embryos were fixed in 10% buffered formalin

overnight and embedded in paraffin. BrdU incorporation was detected by using rat anti-BrdU

primary antibody (1:50; Abcam). Immune complexes were visualized using F(ab)2 biotin-

conjugated donkey anti-rat IgG (1:500; Research Diagnostics Inc.) and the VECTASTAIN Elite

ABC Kit (Vector Laboratories). Sections were counterstained with hematoxylin. For each

sample, BrdU+ cells were counted in 10 randomly selected fields.

2.11 Hematopoietic analysis

Myeloid colony assays (in the absence of added cytokines) were performed as described

previously (436). In brief, bone marrow (BM) cells were suspended in MethoCult® M3234

without cytokines (Stem Cell Technologies) at 105 cells/ml, and colonies were scored after 7-9

days. Complete blood counts were determined by using a Hemavet 850FS (Drew Scientific).

2.12 Echocardiographic and hemodynamic measurements

For echocardiography and cardiac catheterization, mice were anesthetized with

isoflurane/oxygen (2%/100%), and body temperature was maintained at ~37.5°C. Transthoracic

2D and M-mode echocardiography was performed from the long axis view of the heart at the

level of the papillary muscle with a Visualsonics Vevo 770 imaging system (Visualsonics)

equipped with a 30-MHz linear transducer (RMV707B). Measurements of the left ventricular

(LV) end-systolic diameter (LVIDs) and end-diastolic diameter (LVIDd) internal dimensions and

of the LV diastolic posterior wall thickness (LVPWd) were made under Time Motion-mode. The

papillary muscles were excluded from all measurements. Measurements were averaged from at

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least 3 separate cardiac cycles. From TM-mode measurements, end-diastolic volume (EDV) was

calculated as (4.5 x normalized LVIDd 2), end-systolic volume (ESV) was calculated as (3.72 x

normalized LVIDs 2), and stroke volume (SV) was calculated as EDV-ESV. Cardiac output (CO)

was calculated as SV X Heart rate. Fractional shortening (FS) was calculated as (LVIDd –

LVIDs)/ LVIDd x 100%, and ejection fraction (EF) was calculated as (EDV – ESV)/EDV x

100%.

For invasive hemodynamic assessments, a 1.2F catheter (model # FTS-1211B-0018,

Scisense Inc.) was inserted via the right carotid artery into the left ventricle. Hemodynamic

signals were digitized at a sampling rate of 1kHz and acquired to a computer using the MP100

imaging system and Acqknowledge software (BIOPAC Systems, Inc). Following recording of

left ventricular pressure, the catheter was relocated to the ascending aortic for measurement of

aortic blood pressure. Mean arterial pressure (MAP) was calculated as (Systolic pressure +

Diastolic pressure x 2)/3.

2.13 Transverse aortic constriction (TAC)

Eight to nine week-old male mice (25-30g body weight) were anesthetized with 2%

isoflurane, intubated, connected to a ventilator (Harvard Apparatus) and ventilated at a tidal

volume of ~230ul and 135 breaths/min. A para-sternal thoracotomy was performed to expose the

transverse aorta, which was then constricted with a 7/0 silk suture tied around a 27G needle.

Pressure overload was maintained for various times as indicated in Results and Figure Legends.

2.14 MEK inhibitor and Rapamycin treatment

N-[(R)-2,3-dihydroxy-propoxy]-3,4-difluoro-2-(2-fluoro-4-iodo-phenylamino)-

benzamide (PD0325901) was synthesized according to the disclosure in document

WO2007042885(A2). All chemicals necessary for the synthesis were purchased from Sigma

Aldrich.

PD0325901 was dissolved in DMSO at a concentration of 50mg/ml, then resuspended in

vehicle (0.5% hydroxypropylmethylcellulose with 0.2% Tween 80) at a concentration of

0.5mg/ml, and injected intraperitoneally (5mg/kg body weight) daily for the indicated times.

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Control mice were injected with vehicle. The same protocol was used to inject lactating females

for early post-natal treatment.

Rapamycin (LC Laboratories) was dissolved in DMSO at a concentration of 20mg/ml,

then resuspended in vehicle (0.5% hydroxypropylmethylcellulose with 0.2% Tween 80) at a

concentration of 0.2mg/ml, and injected intraperitoneally (2mg/kg body weight) daily for the

indicated times. Control mice were injected with vehicle.

2.15 Quantitative real time RT-PCR

Total RNA from the left ventricle was prepared using the RNeasy mini-kit (QIAGEN).

RNA (2 µg) was reverse transcribed using SuperScriptIII (Invitrogen). TaqMan probe-based

gene expression analyses (Applied Biosystems) for Myh7 (Mm00600555_m1), Myh6

(Mm00440359_m1), Nppa (Mm01255748_g1) and Nppb (Mm00435304_g1) were conducted

according to the manufacturer’s instructions. Each sample was measured in triplicate, and their

relative expression was normalized to Gapdh (4352932E).

2.16 Statistics

All data are presented as mean±SEM. Statistical significance was determined using

Student’s t-test, one-way ANOVA or two-way repeated measure ANOVA, as appropriate. If

ANOVA was significant, individual differences were evaluated using Bonferroni post-test.

Deviation of progeny from Mendelian frequency was assessed by χ2 test. Kaplan–Meier survival

curves were analyzed using the Log-rank test. For experiments in Figure 3-19, significant

outliers were identified using Grubbs’s test. All statistical analyses were performed with

GraphPad Prism 5. For all studies, p<0.05 was considered significant.

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Chapter 3

MEK-ERK Pathway Modulation Ameliorates Disease Phenotypes in

a Mouse Model of Noonan Syndrome Associated with the Raf1L613V

Mutation

This Chapter is a modified version of a paper published in the Journal of Clinical Investigation

(2011 Mar;121(3):1009-25).

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3.1 Abstract

Hypertrophic cardiomyopathy (HCM) is a leading cause of sudden death in children and

young adults. Abnormalities in several signaling pathways are implicated in the pathogenesis of

HCM, but the role of the RAS-RAF-MEK-ERK MAPK pathway has been controversial. Noonan

syndrome (NS) is one of several autosomal-dominant conditions known as “RASopathies”,

which are caused by mutations in different components of this pathway. Germ-line mutations in

RAF1 (which encodes the serine-threonine kinase RAF1) account for approximately 3–5% of

cases of NS. Unlike other NS alleles, RAF1 mutations that confer increased kinase activity are

highly associated with HCM. To explore the pathogenesis of such mutations, we generated

“knock-in” mice expressing the NS-associated Raf1L613V mutation. Like NS patients, mice

heterozygous for this mutation (referred to herein as L613V/+ mice) had short stature,

craniofacial dysmorphia, and hematologic abnormalities. Valvuloseptal development was

normal, but L613V/+ mice exhibited eccentric cardiac hypertrophy and aberrant cardiac fetal

gene expression, and decompensated following pressure overload. Agonist-evoked MEK/ERK

activation was enhanced in multiple cell types, and post-natal MEK inhibition normalized the

growth, facial, and cardiac defects in L613V/+ mice. These data show that different NS genes

have intrinsically distinct pathological effects, demonstrate that enhanced MEK-ERK activity is

critical for causing HCM and other RAF1-mutant NS phenotypes, and suggest a mutation-

specific approach to the treatment of RASopathies.

3.2 Background

Cardiac hypertrophy is a major way by which cardiomyocytes respond to various

stresses, including abnormal neuro-hormonal stimuli, hemodynamic overload and injury. There

are two general types of cardiac hypertrophy (349, 350): physiological, which is associated with

exercise or pregnancy, and pathological, caused by genetic defects (primary), or excessive

afterload, resulting from conditions such as hypertension or valvular stenosis (secondary). With

increased cardiac stress, cardiac hypertrophy may initially represent a compensatory response of

the myocardium. However, chronic pathological hypertrophy predisposes to ventricular

dilatation, heart failure, arrhythmia and/or sudden death (356, 357). Physiological hypertrophy is

typically concentric, with preservation of chamber shape, the absence of inflammation or fibrosis

and normal cardiac gene expression. By contrast, pathological hypertrophy eventually progresses

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to chamber dilatation (eccentric hypertrophy), is often associated with fibrosis, and typically

leads to the reactivation of a fetal gene expression program, characterized by increased levels of

(among others) atrial natriuretic peptide (ANP), brain natriuretic peptide (BNP) and beta-myosin

heavy chain (β-MHC) (437). Delineating the molecular pathways that distinguish physiological

and pathological hypertrophy, and identifying ways to reverse the latter, are of obvious medical

importance.

Primary hypertrophic cardiomyopathy (HCM), the prototypic genetic form of pathologic

hypertrophy, is a leading cause of sudden death in the young (431). The hallmark of HCM is

cardiac hypertrophy in the absence of an obvious inciting hypertrophic stimulus (372). Mutations

in genes encoding sarcomeric proteins (e,g., β-MHC, cardiac troponin T, and myosin-binding

protein C) account for most (~75%) cases of primary HCM. Such mutations usually alter

sarcomere structure and function and result in mechanical, biochemical and/or bioenergetic

stresses that activate cardiomyocyte signaling pathways to mediate the hypertrophic phenotype

(375-378). Aberrant activation of hypertrophic signaling pathways can themselves result in

hypertrophy. For example, germ line mutations in adenosine monophosphate-activated protein

kinase (AMPK) are a rare cause of HCM (380-382). Moreover, genetic and cellular models have

identified multiple signaling systems that can cause or contribute to pathological hypertrophy,

including the calcineurin/NFAT, PI3K/Akt/mTOR, GSK3β and JNK pathways (349, 350, 386).

The detailed mechanism by which aberrant activation of these pathways evokes pathological

hypertrophy remains incompletely understood.

The RAS-RAF-MEK-ERK MAPK pathway (hereafter, the RAS/ERK pathway) is a

central signaling cascade evoked by multiple agonists, including growth factors (e.g., Heregulin,

IGF-1, EGF, PDGF), cytokines (e.g., IL6, cardiotrophin, LIF), G-protein coupled receptor

agonists (angiotensin-II, beta-adrenergic agonists), and physical stimuli (e.g., mechanical

stretch), in cardiomyocytes as well as other cell types (26, 349, 350). The pathway is initiated by

the activation of RAS, which requires RAS-guanine nucleotide exchange factors (RAS-GEFs)

such as SOS1 and, in most cell types, the protein-tyrosine phosphatase SHP2 (encoded by

PTPN11 gene). RAS recruits RAF proteins (RAF1, BRAF, ARAF) to the cell membrane, where

they are activated and subsequently form complexes with MEK1/2 and ERK1/2, aided by

scaffolds, such as KSR. Activated RAF proteins phosphorylate MEK1,2 which, in turn,

phosphorylate ERK1,2. ERKs phosphorylate cytosolic substrates and also translocate to the

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nucleus to stimulate diverse gene expression programs by phosphorylating several transcription

factors (26, 28).

The role of the RAS/ERK pathway in cardiac hypertrophy has been controversial. Some

data argue that excessive activity of this pathway causes HCM, whereas other evidence suggests

involvement in physiological, but not pathological, hypertrophy (409, 410). Transgenic mice

with cardiac-specific expression of oncogenic HRas (G12V) display significant cardiac

hypertrophy, decreased contractility, diastolic dysfunction associated with interstitial fibrosis,

induction of cardiac fetal genes and sudden death (411-413), all of which are consistent with

HCM. In cultured cardiomyocytes, depletion of Erk1/2 with antisense oligonucleotides or

pharmacological inhibition of Mek1/2 attenuates the hypertrophic response to agonist stimulation

(414, 415). Mice with cardiac-specific over-expression of “dominant-negative” Raf1 have no

overt phenotype, but they are resistant to the development of cardiac hypertrophy in response to

pressure overload (416), suggesting that signals from Raf1 are necessary for the hypertrophic

response. On the other hand, transgenic mice expressing an activated Mek1 allele under the

control of the alpha-MHC promoter have concentric hypertrophy with enhanced contractile

performance, show no signs of decompensation over time and reportedly do not progress to

pathological hypertrophy (418). A recent study even argued against any role for ERK1/2 in

cardiac hypertrophy, as Erk1-/-Erk2+/- mice, as well as transgenic mice with cardiac-specific

expression of dual specificity phosphatase 6 (Dusp6), an ERK1/2-specific phosphatase, showed a

normal hypertrophic response to pressure overload and exercise (420).

Over the past ten years, germ line mutations in genes encoding several members of the

RAS/ERK pathway have been identified in a set of related, yet distinct, human developmental

syndromes (31, 32, 144, 438, 439), now collectively termed the RASopathies (32, 439). These

disorders, some (but not all) of which include HCM as a syndromic phenotype, present an

opportunity to clarify the role of the RAS/ERK pathway in cardiac hypertrophy. Noonan

syndrome (NS), a relatively common autosomal dominant disorder (~1/1,000–2500 live births),

typically presents with proportional short stature, facial dysmorphia, and cardiovascular

abnormalities. Many (25-50%) NS patients exhibit some form of myeloproliferative disorder

(MPD), which is usually transient and resolves spontaneously; rarely, NS patients develop the

severe childhood MPD, juvenile myelomonocytic leukemia (JMML) or other forms of leukemia

(440). Mutations in PTPN11 that increase SHP2 phosphatase activity account for ~50% of NS

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cases (145); other known NS genes include SOS1 (~10%) (146, 147), RAF1 (3-5%) (148, 149),

KRAS (1-2%) (150, 151), NRAS (<1%) (152) and SHOC2 (<1%) (183).

Although NS patients typically have valvuloseptal defects, ~20% have HCM (441).

Moreover, different NS genes are differentially associated with HCM. Only ~10% of NS patients

with PTPN11 mutations (442) and ~20% of those with mutations in SOS1 (146) develop HCM.

By contrast, HCM is found in nearly all (~95%) patients bearing RAF1 mutations that cause

increased kinase activity (148, 149). The frequency of HCM also varies in other RASopathies.

HCM is the most frequent (~80%) cardiovascular manifestation of LEOPARD syndrome (LS),

caused by phosphatase-inactivating mutations of PTPN11 (158, 179, 180, 384), but also is

common (~50% in each) in Costello Syndrome (CS), caused by gain-of-function mutations in

HRAS (202, 383), and Cardio-facio-cutaneous (CFC) syndrome, caused by BRAF, MEK1 or

MEK2 mutations (171, 172, 385). Whether these differences represent differential effects of

specific RAS/ERK pathway mutations, the effects of modifiers in the outbred human population,

or both, remains unclear.

Mouse models have begun to address such issues and to provide insight into the detailed

pathogenesis and potential therapeutic approaches to these disorders. For example, we previously

generated a knock-in mouse model of the NS-associated Ptpn11D61G mutation, which

recapitulates the major features of NS, including short stature, facial dysmorphia, mild MPD and

valvuloseptal defects. These mice, like most PTPN11 mutant NS patients, do not have HCM

(204). Transgenic mice expressing a different NS-associated Ptpn11 mutant (Q79R) also show

valvulospetal defects and facial abnormalities seen in NS patients, which are prevented by the

genetic ablation of Erk1/2 and or pre-natal pharmacological inhibition of Mek, respectively (207-

209). Genetic ablation of Erk1 also prevents the development of valvuloseptal defects in mice

expressing a highly activated Ptpn11 mutant in endocardial cells (210). A knock-in mouse model

of CS, caused by HRasG12V mutation shows HCM, but these mice also have aortic stenosis,

making it unclear whether hypertrophy is primary or secondary (443).

Here, we have generated “knock-in” mice expressing the kinase-activating NS mutant

Raf1L613V (L613V). We find that, whereas similar to Ptpn11 mutant mice, mice expressing this

Raf1 allele have short stature, facial dysmorphia, and hematological abnormalities, they do not

have valvuloseptal abnormalities but instead develop HCM. Remarkably, nearly all phenotypic

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abnormalities in Raf1 mutant mice are reversed by post-natal MEK inhibitor treatment. Our

results show that different NS genes have intrinsically distinct pathological effects, and

demonstrate that enhanced MEK/ERK activity is critical for causing HCM and other RAF1-

mutant NS phenotypes. Along with the companion study on LS-associated HCM by Marin et al.

(444), these findings suggest a mutation-specific approach to the treatment of RASopathies.

3.3 Results

3.3.1 Generation of L613V/+ mice

Expression of an activated Raf1 mutant during development might cause embryonic

lethality. Therefore, to investigate the effects of the NS-associated, kinase-activating RAF1L613V

mutant, we designed an “inducible knock-in” Raf1L613V allele (L613Vfl) (Figure 3-1A). The

targeting vector included a cassette containing a splice acceptor sequence, a Raf1 cDNA

fragment encoding wild type (WT) exons 13-16 and a pGK-Neo (Neo) gene. The fusion

cDNA/Neo cassette was flanked by loxP sites and was positioned upstream of exons 13-16 of the

Raf1 gene itself, with an L613V mutation introduced into exon 16 and an HSV-TK cassette for

negative selection. In the absence of Cre recombinase (445), Raf1 exon 12 should be spliced to

the cDNA (exon 13-16), leading to the production of WT Raf1. When Cre is present, the floxed

cassette should be excised, evoking transcription of the mutant Raf1 allele.

The targeting construct was electroporated into G4 embryonic stem (ES) cells, and

correctly targeted clones (L613Vfl/+) were identified by PCR and confirmed by Southern

blotting (Figure 3-1B). We also validated the desired properties of the targeted locus in

L613Vfl/+ ES cells (Figure 3-1C). As expected, expression of the mutant allele was undetectable

(by RT- PCR) in the absence of Cre, but it was induced effectively upon introduction of a Cre-

expression vector (MSCV-GFP-Cre). Mutant Raf1 (protein) also was expressed at levels

comparable to WT Raf1. Chimeras were then generated by outbred morula aggregation, and

germ line transmission was obtained. L613Vfl/+ progeny were crossed to CMV-Cre mice, which

express Cre ubiquitously, and then to WT mice, thereby generating mice with global Raf1L613V

expression (L613V/+ mice) on a 129S6 x C57BL/6 mixed background.

L613V/+ mice were obtained at the expected Mendelian ratio at weaning, indicating that

on this mixed background, Raf1L613V expression during development is compatible with

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Figure 3-1.Generation of inducible Raf1L613V knock-in mice.

(A) Targeting strategy. Structures of the Raf1 locus, targeting vector, mutant allele and location

of probes for Southern blotting are shown. (B) Correct targeting of ES cells. Genomic DNA

from WT ES cells and PCR-positive L613Vfl/+ ES clones was digested with XbaI (5’ and Neo

probe) or BamHI (3’ probe) and subjected to Southern blotting with 5’, 3’ or Neo probes,

respectively. Blots with 5’ and 3’ probes represent non-adjacent lanes on the same gel. (C)

Expression of Raf1L613V allele is inducible. RNA was isolated from WT and L613Vfl/+ ES cells

with or without prior transfection of MSCV-Cre-GFP plasmid and reverse transcribed into

cDNA. A PCR product, obtained by using primers within exon 11 and at the end of exon 16 of

the Raf1 cDNA, was digested with DraIII. Note that the mutant allele is silent until Cre is

introduced, and then is expressed efficiently.

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embryonic viability. However, similar to mice expressing NS-associated Ptpn11 mutant alleles

(204), L613V/+ mice could not be obtained after (>three generations) backcrossing to C57BL/6

mice. Consequently, all experiments herein were performed on the 129S6 x C57BL/6 mixed

background.

3.3.2 L613V/+ mice show multiple NS phenotypes

L613V/+ newborns showed normal size at birth (Figure 3-2A). At weaning, though,

L613V/+ mice were significantly smaller than their WT littermates, and they remained shorter

throughout their lives (Figure 3-2B and Figure 3-2C). Although their overall body proportions

were normal, L613V/+ mice exhibited facial dysmorphia (Figure 3-3). Consistent with their

decreased body size, the skulls of L613V/+ mice were significantly shorter than WT. Their skull

width was increased, however, resulting in a significantly greater width/length ratio. As a result,

L613V/+ mice had a ‘triangular’ facial appearance with a blunter snout and widely-set eyes

(increased inner canthal distance). These features are reminiscent of the facial phenotype of mice

expressing NS-associated Ptpn11 mutations (204, 209, 210) and represent the mouse equivalent

of the facial abnormalities seen in NS patients (446).

Like mouse models of Ptpn11 mutation-associated NS (204, 210) and many NS patients

(447), L613V/+ mice had hematological defects. There was abnormal expansion of myeloid

progenitors, and bone marrow (BM) from L613V/+ mice yielded factor-independent myeloid

colonies (Figure 3-4A). L613V/+ mice also developed splenomegaly, which became more severe

as they aged (Figure 3-4B). Peripheral blood counts were normal at 4 months, but by 1 year,

L613V/+ mice had developed subtle but statistically significant leukocytosis, neutrophilia and

monocytosis (Figure 3-4C) with normal hematocrit and platelet counts.

3.3.3 L613V/+ mice show cardiac hypertrophy with chamber dilatation

Unlike PTPN11 alleles, which are negatively associated with hypertrophic

cardiomyopathy (HCM) in NS patients (442) and in mouse models (204, 208, 210), RAF1

mutations that encode proteins with increased kinase activity are strongly associated with HCM

(148, 149). Remarkably, L613V/+ mice showed evidence of cardiac hypertrophy as early as 2

weeks after birth, as indicated by an increased heart weight to body weight ratio (Figure 3-5A)

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Figure 3-2. Short stature in L613V/+ mice.

(A) Body weight of neonatal WT (n=30) and L613V/+ (n=35) mice. ns, not significant. (B)

Gross appearance of 2-month-old WT and L613V/+ male mice. (C) Growth curves of WT

(n=45) and L613V/+ (n=45) male and female mice. Differences were significant at all time

points (p<0.0001, two-way repeated measure ANOVA; *** p<0.0001, Bonferroni post-test).

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C

Morphometry of skulls

Genotype WT (n=13) L613V/+ (n=11)

Length (mm) 22.9±0.1 21.4±0.3***

Width (mm) 10.4±0.1 10.9±0.1***

Width/Length 0.46±0.01 0.51±0.01***

Inner canthal distance (mm) 6.1±0.1 6.5±0.1***

Figure 3-3. L613V/+ mice have facial dysmorphia.

(A) Gross facial appearance of WT and L613V/+ mice. (B) Representative microCT scans of

skulls from WT and L613V/+ mice. Double-headed arrows indicate inner canthal distance. (C)

Morphometric measurements from microCT scans of a cohort of 2-month-old WT and L613V/+

male mice. *** p<0.0001, 2-tailed Student’s t-test.

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Figure 3-4. L613V/+ mice have hematological defects.

(A) Cytokine-independent myeloid colonies from bone marrow of 2 month-old mice (n=6 for

each genotype). *** p<0.0001, 2-tailed Student’s t-test. (B) Splenomegaly in L613V/+ mice.

Representative gross appearance (left) and spleen weight/body weight (mg/g) ratio (448) in WT

(n=25) and L613V/+ (n=25) mice at 4 months. *** p<0.0001, 2-tailed Student’s t-test. (C)

Increased total white blood cells (WBC), neutrophils (NE) and monocytes (MO) in 1 year-old

L613V/+ mice (n=8 for each genotype). * p<0.05; *** p<0.0001, 2-tailed Student’s t-test.

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Cardiac enlargement became even more obvious in adult L613V/+ mice, with histological

analysis revealing substantial thickening of the ventricular wall and septum (Figure 3-5B).

Increased heart size can reflect a larger number of cardiomyocytes (e.g., as a consequence of

excess proliferation during development) and/or cardiomyocyte hypertrophy. Cardiomyocyte

proliferation, as measured by BrdU incorporation assays, was comparable in E16.5 L613V/+ and

WT embryos (Figure 3-5C). By contrast, cross-sectional area was markedly (~35%) increased in

cardiomyocytes from 8-week-old L613V/+, compared with WT, mice (Figure 3-5D), indicative

of cardiac hypertrophy.

Cardiac hypertrophy can be secondary to pressure overload caused by stenotic valves or

hypertension. Notably, mice expressing the NS-associated Ptpn11D61G mutation have severe

valvuloseptal abnormalities, including atrial, atrioventricular or ventricular septal defects and

double-outlet right ventricle (204). By contrast, valvuloseptal development, as assessed by

histology, appeared normal in 1 week-old L613V/+ mice (Figure 3-5E). Invasive hemodynamic

studies established that ventricular pressure was actually lower in L613V/+ mice compared with

WT controls (see below).

To assess cardiac morphology and function, we performed echocardiography on

L613V/+ mice and littermate controls at 2 and 4 months of age. As expected, left ventricular

diastolic posterior wall thickness (LVPWd) was increased in L613V/+ mice (Figure 3-6A and

Figure 3-6B). Although chamber size was normal in 2 month-old mice, by 4 months, L613V/+

hearts showed an increase in left ventricular internal end-diastolic dimension (LVIDd). Left

ventricular end-systolic dimension (LVIDs) remained within normal limits (Figure 3-6A and

Figure 3-6C), indicating preserved or enhanced function. Consistent with the latter interpretation,

stroke volume (SV), ejection fraction (EF), fractional shortening (FS), and cardiac output (CO)

were increased in L613V/+ mice (Table 3-1).

Invasive hemodynamic studies confirmed and extended these conclusions (Figure 3-6D

and Table 3-2). L613V/+ mice showed increased dP/dt Max, consistent with enhanced

contractility, but no change in cardiac relaxation (-dP/dt). Afterload (systolic pressure) was

slightly lower in L613V/+ mice. Although this finding rules out hypertension as a cause of

hypertrophy in L613V/+ mice, it complicates comparison of dP/dt Max values. For this reason,

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Figure 3-5. L613V/+ mice show cardiac hypertrophy with normal cardiomyocyte

proliferation and valve development.

(A) L613V/+ mice show cardiac hypertrophy, as indicated by heart weight/body weight ratio

(mg/g) as early as 2 weeks after birth (n=14 for each genotype). (B) Representative gross

appearance (left top) and H&E-stained cross-sections (left bottom; original magnification, 4X;

black bar, 2mm) of WT (n=25) and L613V/+ (n=25) hearts (8 weeks); heart weight/body weight

(mg/g) ratio of 4 month-old WT and L613V/+ mice is shown at right. (C) No difference in BrdU

incorporation in E16.5 WT (n=4) and L613V/+ (n=3) hearts. (D) Cross-sectional area of

cardiomyocytes (original magnification, 400X; white bar, 100µm), measured in WGA-strained

sections from 8 week-old mice (n=5 samples for each genotype, with 200 cells counted for each

sample using ImageJ). (E) Representative H&E-stained cross-sections of aortic valves in 1 week-

old mice (original magnification, 40X; two individual samples are shown for each genotype). No

obvious abnormalities were noted in other cardiac valves either. *** p<0.0001, 2-tailed Student’s

t-test.

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Figure 3-6. L613V/+ mice show cardiac hypertrophy with chamber dilatation.

(A) Representative echocardiograms of hearts from 4 month-old mice. Arrows indicate left

ventricular diastolic dimension. (B) Left ventricular diastolic posterior wall thickness (LVPWd)

at 2 and 4 months, measured by echocardiography (n=13 for WT; n=11 for L613V/+). (C) Left

ventricular chamber dimensions of 2 and 4 month-old WT (n=13) and L613V/+ hearts (n=11).

LVIDd, left ventricular internal end-diastolic dimension; LVIDs, left ventricular internal end-

systolic dimension. (D) Cardiac contractility of 4-month-old WT (n=13) and L613V/+ hearts

(n=11) hearts, as measured by invasive hemodynamic analysis. * p<0.05; ** p<0.005, 2-tailed

Student’s t-test.

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2 month 4 month

WT (n=13) L613V/+ (n=11) WT (n=12) L613V/+ (n=11)

Heart rate (bpm) 456±19 454±17 485±12 485±22

SV (µl) 38±1 49±3** 40±1 57±3***

EF% 56±1 63±2* 54±2 63±2**

FS% 29±1 34±2** 28±1 34±1***

CO (ml/min) 18±1 22±2* 19±1 28±2***

Table 3-1. Echocardiographic parameters in WT and L613V/+ mice.

Shown are data from 2 and 4 month-old mice. SV, stroke volume; EF, ejection fraction; FS,

fractional shortening; CO, cardiac output. * p<0.05; ** p<0.005; *** p<0.0001, 2-tailed

Student’s t-test.

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we also compared dP/dt estimated at left ventricular pressure (LVP) of 40mm Hg

(dP/dt@LVP40), therefore reducing or eliminating the influence of afterload (449). Importantly,

dP/dt@LVP40 was increased in L613V/+ animals, providing conclusive evidence of increased

contractility (Figure 3-6D). Moreover, there was no pressure gradient across the aortic valves of

L613V/+ mice (Table 3-2), ruling out aortic valve stenosis as a cause of their cardiac

hypertrophy. Taken together, our finding of eccentric cardiac hypertrophy in the absence of

pressure overload is consistent with the conclusion that L613V/+ mice have pathological

hypertrophy.

Mice (and humans) with pathological hypertrophy often reactivate specific fetal genes

(431, 437). There are two isoforms of cardiac myosin: alpha-myosin heavy chain (alpha-MHC,

faster kinetics) and beta-myosin heavy chain (beta-MHC, slower kinetics). In rodents, the Myh7

(beta-Mhc) gene is expressed mainly in late fetal life, whereas Myh6 (alpha-Mhc) is expressed

predominantly in the adult. Re-expression of Myh7 and a shift from alpha-Mhc to beta-Mhc is a

marker for phenotypic reprogramming and HCM (437). Indeed, Myh6 mRNA levels were

decreased significantly in L613V/+ hearts, and there was a trend (p=0.09, 1-tailed Student’s t-

test) towards increased Myh7 expression (Figure 3-7A). Consequently, the Myh7/Myh6 ratio

increased significantly. Expression of Nppa (atrial natriuretic peptide, Anp) and Nppb (brain

natriuretic peptide, Bnp), two other fetal genes often associated with cardiac hypertrophy (437,

450), was unaffected in L613V/+ hearts (Figure 3-7B).

3.3.4 Enhanced hypertrophic response and functional decompensation in

L613V/+ hearts following pressure overload

Although L613V/+ mice show cardiac hypertrophy, they displayed enhanced cardiac

function without signs of heart failure for at least a year of life. To gain further insight into the

nature of the hypertrophy in L613V/+ mice, we assessed their response (compared with controls)

to biomechanical stress by transverse aortic constriction (TAC). L613V/+ mice had an unusually

high acute death rate after this procedure (Figure 3-8A). Furthermore, the hearts of surviving

L613V/+ mice showed dramatic ventricular, as well as left atrial, enlargement compared with

WT mice (Figure 3-8B). Although WT mice had an ~45% increase in heart weight to body

weight ratio following TAC, L613V/+ mice had an ~ 72% increase. L613V/+ mice also

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WT (n=12) L613V/+ (n=12)

Heart rate (bpm) 516±17 521±17

LVP (mmHg) 121±3 113±2*

EDP (mmHg) 4.1±0.7 3.9±0.5

Systolic P (mmHg) 117±3 109±2

Diastolic P (mmHg) 83±3 77±2

-dP/dt (mmHg/s) -11,010±332 -11,190±327

-dP/dt/MAP (1/s) -118±4 -127±3

Table 3-2. Additional hemodynamic parameters of hearts from 4 month-old mice.

Cardiac catherizations were performed and analyzed as described in Methods. LVP, left

ventricular systolic pressure; EDP, end diastolic pressure; MAP, mean arterial pressure. *

p<0.05, 2-tailed Student’s t-test.

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Figure 3-7. L613V/+ mice show a shift from alpha-Mhc to beta-Mhc expression in hearts.

Alpha- and beta-Mhc gene expression (A) and Anp and Bnp gene expression (B) in 4-month-old

WT (n=6) and L613V/+ (n=9) hearts, assessed by quantitative real-time PCR. * p<0.05; **

p<0.005, 2-tailed Student’s t-test. WT heart sample after 8-week transverse aortic constriction

(TAC) (WT-TAC) (n=1) was served as positive control.

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Figure 3-8. Abnormal response of L613V/+ mice to pressure overload.

(A) Survival curves of WT (n=25) and L613V/+ (n=24) mice following transverse aortic

constriction (TAC). ** p<0.005, Log-rank test. (B) Gross appearance of hearts (left) and heart

weight/body weight (mg/g) ratio (right) at 8 weeks post-TAC. Black dashed lines show markedly

enlarged left atrium in L613V/+, compared with WT, mice. ** p<0.005; *** p<0.0001

(Bonferroni post-test when ANOVA is significant); ## p<0.005 (1-tailed Student’s t-test). (C)

Severe interstitial fibrosis in L613V/+ hearts (Pico Sirius Red-PSR staining; original

magnification, 100X) at 8 weeks post-TAC. Percentage of pixels staining positive with PSR for

interstitial fibrosis was quantified by using ImageJ (n=14 for WT; n=13 for L613V/+). ***

p<0.0001, 2-tailed Student’s t-test. (D) Perivascular fibrosis in hearts (PSR staining; original

magnification, 200X) at 8 weeks post-TAC. Similar results were obtained when Masson’s

Trichrome stain was used to assess fibrosis.

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developed more severe interstitial fibrosis (Figure 3-8C) and perivascular fibrosis (Figure 3-8D

and Figure 3-9A) post-TAC. Two L613V/+ mice were excluded from analysis as by 8 weeks of

TAC these mice had sustained a large (~30% of free ventricular wall) spontaneous transmural

infarct; extensive fibrosis with impaired systolic and diastolic function was evident (Figure

3-9B).

These morphologic and histological findings established that L613V/+ mice have an

altered response to pressure overload. Consistent with this, TAC provoked increases in left

ventricular diastolic posterior wall thickness (LVPWd) in WT and L613V/+ mice, which was

more pronounced in the L613V/+ mice (Figure 3-10A). Left ventricular internal end-diastolic

dimension (LVIDd) did not change after TAC in WT or L613V/+ mice, but remained elevated in

the latter (Figure 3-10B). Most importantly, several parameters of cardiac function, including SV

and FS, deteriorated in L613V/+ mice, whereas these were unaffected in WT mice (Figure 3-

10C). There also was a trend towards decreased cardiac output in L613V/+ mice subjected to

TAC, although this did not reach statistical significance because these mice increased their heart

rate sufficiently to compensate for decreased ventricular function (Figure 3-10D). In addition,

cardiac contractility (measured as either dP/dt Max or dP/dt@LVP40) decreased in L613V/+, but

not in WT mice (Figure 3-11A). Cardiac relaxation assessed by –dP/dt, normalized to mean

arterial pressure (afterload), was reduced comparably, while end-diastolic pressure was increased

to similar extents in WT and L613V/+ mice (Figure 3-11B). Thus, while WT mice could adapt

appropriately to pressure overload, L613V/+ mice exhibited substantial, occasionally fatal,

functional decompensation with reductions in SV, FS and dP/dt Max and @LVP40, consistent

with early stages of heart failure by 8 weeks of TAC.

3.3.5 The Raf1L613V mutant increases Mek and Erk activation in response to

multiple stimuli

Compared with WT RAF1, RAF1L613V has increased kinase activity in vitro and an

enhanced ability to activate MEK/ERK in transfection studies (148, 149). We assessed the effect

of Raf1L613V, expressed at endogenous levels, on the RAS-RAF-MEK-ERK MAPK pathway.

Consistent with the earlier over-expression experiments, Mek and Erk activation (as inferred

from immunoblots with activation-specific antibodies) was enhanced in multiple cell types

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Figure 3-9. Severe perivascular fibrosis and infarct in L613V/+ mice after TAC.

(A) Severe perivascular fibrosis in L613V/+ heart after TAC (PSR staining; original

magnification, 200X). (B) Gross appearance of an LV/+ heart (left) with a severe infarct after

TAC, and severe fibrosis (right) in the infarcted region (PSR staining; original magnification,

100X).

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Figure 3-10. Echocardiographic parameters in WT and L613V/+ mice following pressure

overload.

Left ventricular diastolic posterior wall thickness (LVPWd) (A) and left ventricular internal end-

diastolic dimension (LVIDd) (B) at 8 weeks after TAC. (C) Decreased stroke volume (SV) and

fractional shortening (FS) in L613V/+ mice after TAC. (D) Cardiac output (CO) and heart rate at

8 weeks after TAC. *** p<0.0001 (Bonferroni post-test when ANOVA is significant); # p<0.05;

## p<0.005; ### p<0.0001 (1-tailed Student’s t-test); ns=not significant. n=12 for WT Sham;

n=11 for L613V/+ Sham; n=22 for WT TAC; n=13 for L613V/+ TAC.

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Figure 3-11. Hemodynamic parameters in WT and L613V/+ mice following pressure

overload.

(A) Decreased cardiac contractility in L613V/+ mice after TAC. Because left ventricular

pressures (LVP) are not identical in WT and L613V/+ mice, both dP/dt Max and dP/dt@LVP40

are shown. (B) Additional invasive hemodynamic parameters of hearts after TAC. EDP, end-

diastolic pressure; MAP, mean arterial pressure. * p<0.05; ** p<0.005; *** p<0.0001

(Bonferroni post-test when ANOVA is significant); ## p<0.005; ### p<0.0001 (1-tailed

Student’s t-test); ns=not significant.

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expressing Raf1L613V, in response to a variety of stimuli, including LIF-stimulated ES cells

(Figure 3-12A), and EGF or PDGF-stimulated embryonic fibroblasts (MEFs) (Figure 3-12B and

Figure 3-12C). Of direct relevance to the L613V/+ cardiac phenotype, Mek and Erk activation

also were higher in L613V/+ (compared with WT) neonatal cardiomyocytes stimulated with

RTK (heregulin-β1), cytokine receptor (IL-6) or GPCR (AngII) agonists (Figure 3-13). Recently,

cardiac fibroblasts were implicated in the genesis of cardiac hypertrophy (451, 452); notably,

L613V/+ cardiac fibroblasts also showed enhanced agonist-stimulated Mek/Erk activation

(Figure 3-14). The effects (quantitative and qualitative) of the mutant Raf1 allele on Mek and

Erk activation differed in detail in cardiomyocytes versus cardiac fibroblasts (or MEFs) and in

response to different stimuli. In some cases, mutant Raf1 affected only the peak level

(magnitude) of activation, in others, solely the duration of activation, and for still others, both

magnitude and duration. Such differences might reflect distinct feedback responses to the

agonists in various cell types.

Although it was difficult to detect Erk activation in the adult heart under basal conditions

(data not shown), basal Mek activity was significantly higher in adult L613V/+, compared with

WT, hearts (Figure 3-15A). To compare Mek and Erk activation in vivo, we monitored the

response of WT and L613V/+ mice to pressure overload evoked by TAC for up to 45 minutes.

Mek activation remained significantly higher in L613V/+ hearts throughout the period of acute

TAC (Figure 3-15A). Erk activation was significantly higher in L613V/+ hearts after 30 min

TAC (compared with WT hearts), but was similar to WT at other time points (Figure 3-15B).

We also assayed several signaling pathways implicated in other models of cardiac

hypetrophy/HCM by immunoblotting with appropriate p-specific antibodies. Activation of the

MAPK family members c-jun NH2-terminal kinase (JNK) and p38 was comparable in AngII-

stimulated neonatal cardiomyocytes (Figure 3-16A) and EGF-stimulated cardiac fibroblasts

(Figure 3-16B). Likewise, Akt, GSK3β and p70S6K activation (in response to the agonists

tested) were unaffected by Raf1L613V expression in either cell type. Importantly, in the same

experiments, Mek and Erk activation were enhanced in L613V/+ cardiomyocytes and cardiac

fibroblasts (Figure 3-16).

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Figure 3-12. Raf1L613V mutant causes increased Mek and Erk activation in multiple cell

types.

(A) WT and L613V/+ ES cells were removed from feeders, starved for 6 hr, and then stimulated

with LIF (103U/ml) or left unstimulated (0’). Cell lysates (20µg protein) were resolved by SDS-

PAGE, and analyzed by immunoblotting with the indicated antibodies. One of two experiments

with comparable results is shown. (B, C) Primary mouse embryo fibroblasts (MEFs) from WT

and L613V/+ mice were starved for 16 hr, and then either stimulated with 10 ng/ml EGF (n=4;

two independent experiments in two different MEF strains) or 50 ng/ml PDGF (n=2; one

experiment in each of two different MEF strains). Quantification of blots from all experiments is

shown at right.

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Figure 3-13. Raf1L613V mutant increases Mek and Erk activation in cardiomyocytes.

Cardiomyocytes prepared from neonatal WT and L613V/+ mice were starved for 24 hr, and then

either left unstimulated (0’), or stimulated for the indicated times with 1µg/ml Angiotensin II

(AngII) (A), 10ng/ml IL-6 (B) or 100ng/ml heregulin-β1 (C). Cell lysates (15µg protein) were

immunoblotted with the indicated antibodies. Quantification of blots is shown at right. One of

two experiments with comparable results is shown.

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Figure 3-14. Raf1L613V mutant increases Mek and Erk activation in cardiac fibroblasts.

Cardiac fibroblasts prepared from neonatal WT and L613V/+ mice were starved for 16 hr, and

then either left unstimulated (0’) or stimulated for the indicated times with 50ng/ml EGF (A),

100ng/ml IGF-I (B), 100ng/ml PDGF (C) or 50ng/ml FGF2 (D). Cell lysates (20µg protein) were

immunoblotted with the indicated antibodies. Quantification of blots is shown at right. One of

two experiments with comparable results is shown.

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Figure 3-15. Enhanced Mek and Erk activation in L613V/+ hearts after pressure overload.

Hearts from WT and L613V/+ mice were subjected to TAC for the indicated times (n=5 for each

group at each time point), then lysed and analyzed by immunoblotting with the indicated

antibodies. Each lane represents an individual animal. (A) Mek activation, with all samples from

a single time point analyzed on the same gel. Quantification is shown at right. (B) Representative

samples of Erk activation from each time point analyzed on the same gel. Quantification of all

samples is shown at right. In both cases, Erk2 levels are shown as a loading control. * p<0.05, 2-

tailed Student’s t-test.

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Figure 3-16. Other signaling pathways are unaffected in neonatal cardiac myocytes and

fibroblasts.

(A) Cardiomyocytes from neonatal WT and L613V/+ mice were starved for 24 hr, and then

either left unstimulated (0’) or stimulated with AngII (1µg/ml). (B) Cardiac fibroblasts from

neonatal WT and L613V/+ mice were starved for 16 hr, and then either left unstimulated (0’) or

stimulated with EGF (10ng/ml). Cell lysates (15-20µg protein) were analyzed by

immunoblotting with the indicated phospho-specific antibodies for other pathways implicated in

cardiac hypertrophy.

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3.3.6 MEK inhibitor treatment normalizes NS phenotypes in L613V/+ mice

The genetics of NS (and other RASopathies), and the ability of Raf1L613V to selectively

enhance Mek and Erk activation by multiple agonists in cardiomyocytes and cardiac fibroblasts,

strongly implicate enhanced Mek/Erk activation in the pathogenesis of NS phenotypes, including

HCM. We asked whether any of these phenotypes might be reversible if Mek/Erk activation

were normalized by treatment of L613V/+ mice with a MEK inhibitor. In initial experiments,

the ATP-uncompetitive inhibitor PD0325901 (453) or empty vehicle was injected

intraperitoneally (IP) daily to WT and L613V/+ mice (5mg/kg body weight), beginning at 4

weeks and continuing for the succeeding 6 weeks. Importantly, at the start of the treatment

period, L613V/+ mice already show significant growth defects, facial dysmorphia and cardiac

hypertrophy.

Remarkably, the body length of L613V/+ mice began to catch up with WT mice after 1

week of treatment, and by 2 weeks, L613V/+ mice were the same length as untreated WT mice

(Figure 3-17A). MEK inhibitor-treated WT mice also increased their body length, such that by

the last two weeks of treatment, they were significantly longer than control, untreated WT mice.

Notably, however, MEK inhibitor-treated L613V/+ mice achieved the same final body length as

treated WT mice, arguing that increased Mek/Erk activity is the primary cause of the growth

defect in L613V/+ mice (see Discussion). Inhibitor treatment also increased the body weight of

L613V/+ mice, but surprisingly, they, as well as inhibitor-treated WT mice, gained substantially

more weight than untreated WT mice (Figure 3-17B). Increased body weight in MEK-inhibitor-

treated mice was accompanied (and presumably in large part caused) by an obvious increase in

body fat; thus, increased adiposity/body mass is an unanticipated, and previously unreported,

side-effect of PD0325901 (and possibly, MEK inhibitor) treatment (see Discussion).

MEK inhibitor treatment also affected the L613V/+ cardiac phenotype. The heart weight

to body weight ratio (Figure 3-17C) in L613V/+ mice was restored to normal range (WT control)

after treatment, whereas there was no significant change in WT mice. Echocardiography (Figure

3-17D and Figure 3-17E) and invasive hemodynamic (Figure 3-19) studies showed significant

improvement in multiple parameters of cardiac morphology and function. Furthermore,

histological assessment of cross-sectional area of cardiomyocytes confirmed the normalization of

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Figure 3-17. MEK inhibitor treatment rescues growth defect and cardiac hypertrophy in

L613V/+ mice.

Mice were injected intraperitoneally (IP) daily with PD0325901 (5mg/kg body weight) or

vehicle, starting at 4 weeks of age and for the succeeding 6 weeks. Body length (A) and body

weight (B) were measured weekly. Note the rapid normalization of body length, as well as the

increase in body weight caused by inhibitor treatment. # p<0.05; ## p<0.005; ### p<0.0001

(two-way repeated measure ANOVA); * p<0.05; ** p<0.005; *** p<0.0001 (Bonferroni post-

test when ANOVA is significant; black asterisk for WT treatment vs. WT control; red asterisk

for L613V/+ treatment vs. L613V/+ control). (C) Heart weight/body weight (mg/g) ratio and (D)

Left ventricular diastolic posterior wall thickness (LVPWd) are restored to within normal limits

in inhibitor-treated mice. ** p<0.005; *** p<0.0001 (Bonferroni post-test when ANOVA is

significant); # p<0.05 (1-tailed Student’s t-test); ns=not significant. (E) Left ventricular end-

diastolic dimension (LVIDd). **p<0.005 (Bonferroni post-test when ANOVA is significant); #

p<0.05; ## p<0.005 (1-tailed Student’s t-test); ns=not significant. n=14 for WT control; n=10 for

L613V/+ control; n=6 for WT treatment (WT PD); n=14 for L613V/+ treatment (L613V/+ PD).

(F) Cross-sectional area of cardiomyocytes (original magnification, 400X; white bar, 100µm),

measured in WGA-strained heart sections (n=2 samples for each group, with 200 cells counted

for each sample using ImageJ). *** p<0.0001 (Bonferroni post-test when ANOVA is

significant); ns=not significant.

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Figure 3-18. Normalized cardiac morphology and function after MEK inhibitor treatment.

(A) Left ventricular diastolic posterior wall thickness (LVPWd) normalized by BW1/3. (B) Left

ventricular internal end-diastolic dimension (LVIDd) normalized by BW1/3. (C) Normalized

stroke volume (SV). End-diastolic volume (EDV) = (4.5 x normalized LVIDd2); End-systolic

volume (ESV) = (3.72 x normalized LVIDd2); SV=EDV-ESV. (D) Normalized cardiac output

(CO). CO=Normalized SV X Heart rate. ** p<0.005; *** p<0.0001 (Bonferroni post-test when

ANOVA is significant); # p<0.05; ## p<0.005 (1-tailed Student’s t-test); ns=not significant.

n=14 for WT control; n=10 for LV/+ control; n=6 for WT treatment (WT PD); n=14 for

L613V/+ treatment (L613V/+ PD).

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Figure 3-19. MEK inhibitor treatment normalizes cardiac function in L613V/+ mice.

(A) Echocardiographic parameters of hearts after MEK inhibitor treatment as described in Figure

3-17. Note normalization of stroke volume (SV) and fractional shortening (FS), with a trend

towards normalization of cardiac output (CO). * p<0.05; ** p<0.005; *** p<0.0001 (Bonferroni

post-test when ANOVA is significant); # p<0.05 (1-tailed Student’s t-test); ns=not significant.

(B) Hemodynamic parameters, assessed by cardiac catheterization, after MEK inhibitor

treatment. For calculating statistical significance, significant outliers (circled data points), as

accessed by Grubbs' test, were removed. *: p<0.05 (Bonferroni post-test when ANOVA is

significant); p=0.09 when outliers are not removed. # p<0.05 (1-tailed Student’s t-test); p=0.12

when outliers are not removed. ns, not significant. n=14 for WT control; n=10 for L613V/+

control; n=6 for WT treatment (WT PD); n=14 for L613V/+ treatment (L613V/+ PD).

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cardiomyocyte size in L613V/+ mice after treatment (Figure 3-17F). The significant increase in

body size and body mass caused by inhibitor treatment potentially complicates echocardiography

and invasive hemodynamic comparisons of pre- and post-treatment WT and L613V/+ mice,

respectively. Therefore, we compared all parameters using both nominal values and values

normalized by BW1/3 (Figure 3-18); overall, the two analysis regimes lead to similar conclusions.

First, there was a significant reduction (towards normal) in left ventricular posterior wall

thickness in L613V/+ mice after treatment (Figure 3-17D); this difference was even more

significant when normalized by BW1/3 (Figure 3-18). Nominal left ventricular end-diastolic

dimension was unchanged in inhibitor-treated L613V/+ mice (Figure 3-17E), although when this

value is normalized, chamber dilatation was improved significantly, becoming comparable to

WT controls (Figure 3-18B). Inhibitor treatment clearly reduced the abnormal SV and FS in

L613V/+ mice towards normal (untreated or treated WT) values (Figure 3-19A and Figure

3-18C). There also was a strong trend towards decreased CO in L613V/+ mice after inhibitor

treatment (Figure 3-19A and Figure 3-18D). Finally, the excessive cardiac contractility (dP/dt

and dP/dt@LVP40) in L613V/+ mice was ameliorated by inhibitor treatment, while cardiac

relaxation remained unchanged (Figure 3-19B).

MEK inhibitor treatment did not improve the facial dysmorphia in L613V/+ mice in the

above study, most likely because skull development had already been completed by the onset of

drug administration. We tested whether earlier, but still post-natal, MEK inhibitor treatment

could prevent/ameliorate L613V/+ facial defects. Lactating female mice were injected IP with

PD0325901 (5mg/kg body weight) daily, beginning at postnatal day 0 (P0) until weaning (P21).

Weaned mice were then injected individually with same dose of inhibitor for another 5 weeks.

As expected from our initial treatment regimen (Figure 3-17A), the growth defect in L613V/+

mice was again prevented in this new study. Remarkably, however, earlier MEK inhibitor

treatment had a dramatic effect on the appearance of L613V/+ mice: they no longer had

“triangular” faces and instead, appeared indistinguishable from control (treated or untreated) WT

mice (Figure 3-20A). MicroCT morphometry confirmed that inner canthal distance was reduced

significantly, while skull length was increased and skull width and width/length ratio were

decreased in inhibitor-treated L613V/+ mice; all of these values were indistinguishable from WT

(treated or untreated) mice by the end of the treatment period (Figure 3-20B).

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Figure 3-20. Early post-natal MEK inhibitor treatment rescues facial dysmorphia in

L613V/+ mice.

Lactating female mice were injected IP daily with PD0325901 (5mg/kg body weight) or vehicle,

starting at postnatal day 0 (P0) until weaning (P21). Weaned mice were then injected IP

individually with PD0325901 (5mg/kg body weight) or vehicle daily for another 5 weeks. (A)

Gross facial appearances for mice treated with PD0325901 (PD) or vehicle. (B) Morphometric

measurements of skulls from microCT scans. ** p<0.005; *** p<0.0001 (Bonferroni post-test

when ANOVA is significant). n=11 for WT control; n=10 for L613V/+ control; n=6 for WT

treatment (WT PD); n=7 for L613V/+ treatment (L613V/+ PD).

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3.4 Discussion

We describe here a knock-in mouse model for Noonan syndrome (NS) caused by a Raf1

gain-of-function mutation. Similar to mouse models of Ptpn11 mutation-associated NS,

Raf1L613V heterozygosity causes proportional short stature, facial dysmorphia, and hematological

defects. Unlike phosphatase-activating Ptpn11 alleles, which cause valvuloseptal abnormalities

(204, 207, 210), L613V/+ mice have normal valvuloseptal development and instead exhibit

eccentric cardiac hypertrophy that decompensates upon pressure overload. Agonist-evoked

Mek/Erk activation is enhanced in multiple cell types without changes in several other signaling

pathways implicated in cardiac hypertrophy/HCM. Remarkably, post-natal MEK inhibition

normalizes the growth, facial and cardiac defects in L613V/+ mice, demonstrating that continued

MEK/ERK activity is critical for causing HCM and other NS phenotypes and identifying MEK

inhibitors as potential therapeutic agents for the treatment of NS.

RASopathies are a class of human genetic syndromes caused by germ line mutations in

genes that encode components of the RAS/ERK pathway (32, 439). Not surprisingly, these

disorders share several features (albeit with varying degrees of penetrance), yet each also

exhibits unique and characteristic phenotypes. Conceivably, the specific mutant gene, possibly as

a consequence of its position in the pathway and susceptibility to feedback regulation, could

direct the phenotype. Alternatively, genetic modifiers in the highly outbred human population

could be determinative.

Previous mouse models suggest that both the gene and the genetic background are

important to the ultimate RASopathy phenotype. Clearly, different mutations in the same

RASopathy gene can result in distinguishable phenotypes: gain-of-function Ptpn11 mutations,

depending on the degree of their phosphatase activity, cause a variable spectrum of NS

phenotypes (204, 207, 210). The current study, along with a parallel analysis of knock-in mice

expressing a NS-associated Sos1E846K mutant (211), shows that mutations in different genes that

cause the same RASopathy syndrome yield different phenotypes: mice with phosphatase-

activating Ptpn11 mutations have valvuloseptal defects, but not HCM (204, 207); Sos1E846K/+

mice develop left ventricular hypertrophy with incompletely penetrant aortic stenosis; and

Raf1L613V/+ mice exhibit HCM with normal valvuloseptal development.

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On the other hand, mutations associated with different RASopathies also have distinct

effects in mice. In contrast to the NS mice discussed above, an HRasG12V knock-in mouse model

of Costello Syndrome (CS) shows abnormal cranial dimensions, papillomas and angiosarcomas.

These mice have cardiac hypertrophy, but also aortic stenosis, making it unclear whether the

hypertrophy is primary or secondary (439, 443). As described in the accompanying manuscript

(444), a mouse model of LEOPARD syndrome (LS) caused by Ptpn11Y279C, indicates that, unlike

Ptpn11 alleles with increased catalytic activity, catalytically impaired mutants develop HCM and

skeletal abnormalities (as well as short stature and facial dysmorphia).

While the specific mutation plays a major role in determining RASopathy phenotype,

modifier loci also clearly contribute: just as there is considerable phenotypic variation between

family members carrying the same NS or LS allele (454), there are differences in disease

spectrum and severity of mice with Ptpn11 (204, 210) and Raf1 (data not shown) mutations on

different strain backgrounds. Ptpn11D61G/+ mice show incomplete penetrance of valvuloseptal

defects on mixed background and various penetrance of embryonic lethality on different strain

backgrounds (204, 210). Raf1L613V/+ mice were obtained at the expected Mendelian ratio on

mixed background, whereas on the C57BL/6 background this mutant allele almost always was

lethal (data not shown). All of these data suggest that incomplete penetrance reflects strain-

specific modifiers. Genomic scans using SNP panels should help to determine whether cloneable

modifiers exist or heterosis accounts for the variable penetrance.

The role of the RAS/ERK pathway in cardiac hypertrophy has been controversial. Over-

expression of MAPK phosphatase 1 (MKP-1) blocks both agonist-induced hypertrophy in vitro

and pressure overload-associated hypertrophy in vivo (455). However, MKP-1 inactivates all

three major MAPKs, so this study could not address the specific effects of Ras/Erk pathway

activation. Depletion of ERK1/2 with antisense oligonucleotides or pharmacological inhibition

of MEK1/2 attenuates the hypertrophic response to agonist stimulation of cultured

cardiomyocytes (414, 415), consistent with a requirement for MEK/ERK activation in the

hypertrophic response. Transgenic mice with cardiac-specific expression of HRasG12V display

HCM associated with interstitial fibrosis and sudden death (411-413). Cardiac-specific Nf1-

deleted mice develop marked cardiac hypertrophy, progressive cardiomyopathy, and fibrosis as

adults (456).

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Conversely, other studies suggest that MEK/ERK activity is dispensable for

cardiomyocyte hypertrophy. Transgenic mice with cardiac-restricted expression of activated

Mek1 exhibit concentric hypertrophy without signs of cardiomyopathy (418). Although

hypertrophy in this model was interpreted as physiological, these mice also have impaired

diastolic function and reactivated cardiac fetal gene expression, which is more consistent with

pathologic hypertrophy. A recent study showed that Erk1-/-Erk2+/- mice, or transgenic mice with

cardiac-specific expression of dual specificity phosphatase 6 (Dusp6), an Erk1/2-specific

phosphatase, showed a normal hypertrophic response to pressure overload and exercise (420). In

both of these lines of mice, however, residual Erk activity cannot be excluded. Also, it is possible

that Dusp6 has other targets besides Erk1/2, which could complicate interpretation of these

results. Moreover, most of these earlier studies involved cardiomyocyte-specific expression or

deletion of potential hypertrophy-related genes, which excludes the potential contribution of

other cell types in the heart to the hypertrophic response. Recent studies show that cardiac

fibroblasts play key roles in myocardial development and function (457, 458). Embryonic

cardiac fibroblasts induce myocyte proliferation, whereas adult cardiac fibroblasts promote

myocyte hypertrophy (457), and evoke pathological hypertrophy and fibrosis in response to

disease stimuli (451, 452). Of particular note, enhanced Ras/Erk activation in cardiac fibroblasts

is implicated in pathological hypertrophy and fibrosis caused by over-expression of the beta-

adrenergic receptor in cardiomyocytes (452).

Our mouse model, in which a NS-associated Raf1 mutant is expressed globally under

normal promoter control, supports the conclusion that excessive Ras/Erk pathway activity causes

HCM. Several lines of evidence indicate that L613V/+ mice have pathologic cardiac

hypertrophy. Hypertrophy is eccentric in these mice, and they show the characteristic shift from

alpha-Mhc to beta-Mhc expression seen in pathological hypertrophy. In response to pressure

overload (TAC), they have an unusually high death rate, presumably due to inability to adapt to

this stress or arrythmia, while surviving mice show clear evidence of functional decompensation.

Importantly, in our model, unlike many previous studies (see above), the mutant allele is

expressed in both cardiomyocytes and cardiac fibroblasts (as well as multiple other cell types).

Moreover, Mek/Erk activation is enhanced in response to multiple agonists in these cells. It will

be important to determine whether mutant expression in cardiomyocytes, cardiac fibroblasts, or

both is important for HCM in L613V/+ mice; our inducible Raf1 allele should facilitate such

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analyses. Most importantly, post-natal MEK inhibitor treatment substantially normalizes the

cardiac defects in L613V/+ mice, providing strong evidence for the critical role of the RAS/ERK

pathway in initiating and maintaining the cardiac hypertrophic response.

Post-natal MEK inhibitor treatment also normalizes the growth defects and, if

administered early enough, the facial dysmorphia in L613V/+ mice. Notably, MEK inhibitor

treatment also increases the body length of WT mice, but there is no difference in the final body

length (after treatment) between the WT and mutant-treated groups. Likewise, MEK inhibitor

treatment (at doses that effectively normalized L613V/+ cardiac anatomy and function) had little

effect on cardiac function in WT mice. These results strongly suggest that all of these NS

phenotypes are due to excessive MEK/ERK activity (as opposed to the MEK inhibitor acting on

a parallel pathway to mitigate syndromic features).

Unexpectedly, we found that PD0325901 treatment caused a significant increase in body

weight with an obvious increase in body fat. Although we cannot exclude the possibility that this

is an idiosyncratic (i.e., off-target) effect of this specific MEK inhibitor, other evidence points to

a potential obesity-promoting effect of MEK/ERK inhibition. For example, leptin activates Erk

via an Shp2-dependent pathway (459, 460) and deletion of Shp2 in post-mitotic forebrain

neurons causes early-onset obesity with decreased ERK activation and evidence of leptin

resistance (461). We suspect that MEK inhibition may act in analogous ways to promote obesity

in our mice.

In summary, our data demonstrate a critical role of the RAS/ERK pathway in the genesis

of HCM in NS, and show that NS phenotypes can be rescued by pharmacological inhibition of

MEK1/2. Previous studies showed that genetic ablation of Erk1/2 (207, 210) or pre-natal

treatment with a MEK inhibitor (209, 211) can prevent some NS phenotypes. While these studies

provide evidence for the key role of Mek/Erk hyperactivity in NS pathogenesis, they did not

resolve whether MEK inhibition can reverse these phenotypes. By contrast, our results suggest

that MEK inhibition may be useful for the specific treatment of Raf1 mutant NS, and possibly for

other RASopathies associated with increased MEK/ERK pathway activity. Interestingly, a

parallel study show that LS-associated HCM is associated with hyper-activation of PI3K/Akt

pathway, and can be rescued by Rapamycin treatment (444). Taken together, these studies argue

for a mutation-specific, “personalized” approach to RASopathy therapy.

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Chapter 4

Increased BRAF heterodimerization is the common pathogenic

mechanism for Noonan Syndrome-associated RAF1 mutants

This Chapter is a modified version of a paper published in Molecular and Cellular Biology (2012

Oct;32(19):3872-90).

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4.1 Abstract

Noonan syndrome (NS) is a relatively common autosomal dominant disorder,

characterized by congenital heart defects, short stature and facial dysmorphia. NS is caused by

germ-line mutations in several components of the RAS-RAF-MEK-ERK MAPK pathway,

including both kinase-activating and kinase-impaired alleles of RAF1 (~3-5%), which encodes a

serine-threonine kinase for MEK1/2. To investigate how kinase-impaired RAF1 mutants cause

NS, we generated knock-in mice expressing Raf1D486N. Raf1D486N/+ (hereafter D486N/+) female

mice exhibited a mild growth defect. Male and female D486N/D486N mice developed

concentric cardiac hypertrophy and incompletely penetrant, but severe, growth defects.

Remarkably, Mek/Erk activation was enhanced in Raf1D486N-expressing cells compared with

controls. RAF1D486N, as well as other kinase-impaired RAF1 mutants, showed increased

heterodimerization with BRAF, which was necessary and sufficient to promote increased

MEK/ERK activation. Furthermore, kinase-activating RAF1 mutants also required

heterodimerization to enhance MEK/ERK activation. Our results suggest that increased

heterodimerization ability is the common pathogenic mechanism for NS-associated RAF1

mutations.

4.2 Background

Noonan syndrome (NS) is a relatively common (1 in 1,000–2,500 live births) autosomal

dominant disorder (127, 462, 463), characterized by short stature, craniofacial dysmorphia, a

wide spectrum of congenital cardiac anomalies, and an increased risk of hematopoietic

malignancy. Although NS is genetically heterogeneous (31, 32, 144), all known cases are caused

by germ-line mutations in conserved components of the canonical RAS-RAF-MEK-ERK MAPK

(hereafter, RAS/ERK) cascade, a key regulator of cell proliferation, differentiation and survival

(26, 27). Mutations in PTPN11, which encodes the protein-tyrosine phosphatase SHP2, account

for approximately half of NS cases (145). Other known NS genes include SOS1 (~10%) (146,

147), RAF1 (3-5%) (148, 149), KRAS (<2%) (150, 151), NRAS (152) and SHOC2 (183) (<1-2%).

Mutations in some of these genes, as well as in genes encoding other RAS/ERK pathway

components, also cause phenotypically related disorders, such as neurofibromatosis type 1

(NF1), Costello syndrome, cardio-facio-cutaneous (CFC) syndrome, and LEOPARD syndrome;

together with NS, these syndromes are now termed “RASopathies” (32). How mutations in the

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same signaling pathway cause similar, yet clearly distinct, phenotypes remains unclear.

Consequently, detailed understanding of RASopathy pathogenesis should yield new insights into

RAS/ERK pathway regulation.

RAF family serine-threonine kinases (250-252) function as key RAS effectors,

phosphorylating and activating the dual specificity kinases MEK1 and MEK2, which in turn

promote the activation of the MAPKs, ERK1 and ERK2. The three mammalian RAF family

members (RAF1, BRAF and ARAF) differ in their expression profiles and regulatory

mechanisms, and have distinct roles during development. RAF1 (also known as CRAF) is the

most intensively studied isoform. Nevertheless, controversy and disagreement surround the

precise molecular events required for RAF1 activation, which include RAS-dependent

membrane recruitment, conformational changes, dimerization or oligomerization, scaffold

protein binding, and distinct phosphorylation/dephosphorylation events. RAF1 also has

important kinase-independent functions. For example, it interacts with and inhibits apoptosis

signal-regulating kinase 1 (ASK1) (227, 338), Mammalian STE20-like kinase 2 (MST2) (228,

339) and Rok-α (229).

Two groups (148, 149) identified multiple missense mutations of RAF1 in NS, which

cluster in three regions. Approximately 70% of NS-associated RAF1 alleles alter the motif

flanking S259 within the so-called CR2 domain, which binds to 14-3-3 proteins and is critical for

auto-inhibition (163, 164). The second group of mutations (~15%) affects residues within the

activation segment of the kinase domain (D486 and T491). The remaining alleles (~15%)

involve two adjacent residues (S612 and L613) located C-terminal to the kinase domain.

Transient transfection studies indicate that mutations affecting the 14-3-3 binding motif or the C-

terminus of the protein enhance RAF1 kinase activity and increase MEK/ERK activation in cells.

By contrast, mutations that cluster in the activation segment are kinase-impaired and reportedly

act as dominant negative or null alleles (148, 149). Previous work suggested that the increased

kinase activity of NS-associated CR2 domain mutants results from decreased S259

phosphorylation and consequent dissociation from 14-3-3 (149, 165, 166), but the mechanism

underlying increased kinase activity of the RAF1 C-terminal mutants remains unclear. Likewise,

how kinase-defective RAF1 alleles cause NS has remained obscure, if not paradoxical. Studies

of kinase-defective BRAF alleles strongly implicate enhanced MEK/ERK activation and

heterodimerization with RAF1 in human melanoma pathogenesis (167, 168). The paradoxical

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activation of the MEK/ERK pathway in wild type cells treated with selective small molecule

BRAF inhibitors also has been attributed to the ability of these inhibitors to induce BRAF/RAF1

heterodimer formation (169, 170). The relevance of these observations for RAF1 alleles

expressed at physiological expression levels remains to be determined.

Kinase-activating and kinase-impaired RAF1 alleles also are associated with different

syndromic phenotypes. NS patients with RAF1 mutations have a much higher incidence (~75%)

of hypertrophic cardiomyopathy (HCM) than is found in the overall NS population (~20%).

However, only RAF1 alleles encoding kinase-activated mutants are highly associated (~95%)

with HCM. Recently, we reported that knock-in mice expressing the kinase-activated allele

Raf1L613V develop typical NS features (short stature, facial dysmorphia, haematological

abnormalities), as well as HCM (464). As expected, agonist-evoked MEK/ERK activation was

enhanced in multiple cell types expressing Raf1L613V. Moreover, postnatal MEK inhibition

normalized the growth, facial, and cardiac defects in L613V/+ mice, showing that enhanced

MEK/ERK activation is critical for evoking RAF1-mutant NS phenotypes.

Whether kinase-defective Raf1 alleles also faithfully model human NS, and if so, how

kinase-activating and kinase-defective mutants can cause similar phenotypes, remains to be

resolved. To investigate this paradox, we generated and analyzed knock-in mice expressing the

kinase-impaired NS mutant Raf1D486N, and also re-examined the effects of various other NS

mutants expressed at more physiological levels than in previous transient transfection studies.

Our results strongly implicate increased heterodimerization ability as the common pathogenic

mechanism for NS-associated RAF1 mutations.

4.3 Results

4.3.1 Generation of Raf1D486N mice

To avoid the possibility that expression of kinase-impaired Raf1 might cause embryonic

lethality, we designed an inducible Raf1D486N “knock-in” allele (D486Nfl; Figure 4-1A). The

targeting vector included a cassette containing a splice acceptor sequence, a Raf1 cDNA

fragment encoding WT exons 13-16, and a pGK-Neo gene. The fusion cDNA/Neo cassette was

flanked by LoxP sites and was positioned between Raf1 exons 12 and 13, with the D486N

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Figure 4-1. Generation of Raf1D486N knock-in mice.

(A) Targeting strategy. Structures of the Raf1 locus, targeting vector, mutant allele and location

of probes for Southern blotting are shown. (B) Selection of targeted ES cells. To assay for

inclusion of the 5’ loxP site, a PCR product, obtained by using primers within exon 11 and the

Raf1 cDNA, was digested with NotI. To ensure inclusion of the D486N mutation, a PCR

product, obtained by using primers around exon 13, was digested with ApoI. To assay for

inclusion of the Neo cassette, a PCR product was obtained by using primers within the PGK Neo

gene. (C) ES cells are targeted correctly. Genomic DNA from WT ES cells and PCR-positive

D486Nfl/+ ES clones was digested with XbaI (5’ and Neo probe) or BamHI (3’ probe) and

subjected to Southern blotting with 5’, 3’ or Neo probes, respectively. (D) Inducible expression

of the Raf1D486N allele. RNA was isolated from WT and D486Nfl/+ ES cells with or without

prior transfection of pMSCV-Cre-GFP plasmid and reverse transcribed. A PCR product,

obtained by using primers within exon 11 and at the end of exon 16 of the Raf1 cDNA, was

digested with ApoI. Note that the mutant allele is silent until Cre is introduced, and then is

expressed efficiently. (E) Progeny from D486N/+ matings with the indicated littermates.

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mutation introduced into exon 13 and an HSV-TK cassette for negative selection. In the absence

of Cre recombinase (Cre), Raf1 exon 12 should be spliced to the cDNA (exons 13-16), resulting

in the expression of WT Raf1. When Cre is present, the floxed cassette is excised, leading to the

transcription of the mutant Raf1 allele (D486N).

The targeting construct was electroporated into G4 ES cells, and correctly targeted clones

(D486Nfl/+) were identified by PCR (Figure 4-1B) and confirmed by Southern blotting (Figure

4-1C). To test whether the mutant Raf1 allele could be induced in D486Nfl/+ ES cells, a Cre

expression vector (pMSCV-GFP-Cre) was introduced; as expected, this resulted in efficient

transcription of the mutant allele (Figure 4-1D). Chimeras were generated by outbred morula

aggregation, and germ-line transmission was obtained. D486Nfl/+ progeny were crossed to EIIa-

Cre mice, which express Cre ubiquitously, and then to WT mice, thereby generating mice with

global Raf1D486N expression (referred to as D486N/+ mice) on a 129Sv × C57BL/6 mixed

background. D486N/+ littermates were intercrossed to generate mice homozygous for Raf1D486N

(referred to as D486N/D486N mice). D486N/+ and D486N/D486N mice were obtained at the

expected Mendelian ratios at weaning (Figure 4-1E), indicating that Raf1D486N expression is

compatible with embryonic development.

4.3.2 Phenotypes of D486N/+ and D486N/D486N mice

Major features of NS include short stature, facial dysmorphia, cardiovascular

abnormalities and often, some form of myeloproliferative disease (MPD) (438). D486N/+ female

mice exhibited a mild, but reproducible growth defect compared with WT littermates, although

male heterozygotes had normal body size (Figure 4-2A). Male and female D486N/D486N mice

showed two distinct patterns of growth: about two thirds of these animals had a normal growth

pattern but in the remaining one third, body length and weight were markedly decreased (~50%

smaller than littermate controls). Hereafter, we refer to the normal sized mice as n-

D486N/D486N, and the smaller ones as s-D486N/D486N. The majority of s-D486N/D486N

mice died shortly after weaning, while those that survived continued to have reduced body size, a

hunched appearance with ruffled fur and frequent tremors, and ultimately, died between 4-8

months of age. Consistent with their decreased body size, the skulls of s-D486N/D486N mice

were significantly shorter than those of WT mice. However, skull width was decreased only

slightly, resulting in a significant increase in width/length ratio and a “triangular” facial

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appearance (Figure 4-2B). By contrast, n-D486N/D486N mice had normal lifespan, body size

and facial morphology (Figure 4-2B). Because D486N/D486N mice were maintained on a mixed

strain background, genetic modifiers presumably account for the variable phenotype.

Similar to RAF1D486N/+ NS patients, D486N/+ mice had normal heart size. By contrast, n-

D486N/D486N mice showed cardiac enlargement, manifested by a significantly increased heart

weight to body weight ratio compared with WT and D486N/+ littermates (Figure 4-2C).

Neonatal cardiomyocytes prepared from D486N/D486N mice showed significant increased

surface area compared with WT cardiomyocytes, indicating that cardiac enlargement was due to

hypertrophy (Figure 4-2D). Echocardiography performed on 4 month-old n-D486N/D486N mice

showed increased left ventricular diastolic posterior wall thickness (LVPWd) as expected, while

D486N/+ mice had normal LVPWd (Figure 4-2E). In contrast to Raf1L613V/+ mice, which develop

cardiac dilatation (464), left ventricular internal end-diastolic dimension (LVIDd) remained

normal in D486N/+ and n-D486N/D486N mice, while left ventricular internal end-systolic

dimension (LVIDs) tended to be reduced in D486N/+ hearts and was decreased significantly in

n-D486N/D486N hearts (Figure 4-2F). Stroke volume (SV), fractional shortening (FS), cardiac

output (CO), and ejection fraction (EF) also were increased in n-D486N/D486N mice (Figure

4-2G). Cardiac parameters could not be assessed in s-D486N/D486N mice because of their size

and general ill health. Overall, these findings indicate that D486N/D486N mice have concentric

cardiac hypertrophy with enhanced cardiac function.

Unlike other mouse models of NS (204, 210, 464, 465), D486N/+ and D486N/D486N

mice did not develop splenomegaly (Figure 4-3A), nor they show overt hematological defects

(Figure 4-3B and Figure 4-3C).

4.3.3 Raf1D486N expression increases Mek/Erk activation in response to

multiple stimuli

NS-associated RAF1 mutations include kinase-activating mutations (e.g., S257L and

L613V) and mutations with impaired kinase activity (e.g., D486N and T491I/R) (148, 149). To

begin to investigate why kinase-impaired and -activated RAF1 mutants cause similar

phenotypes, we assessed Mek and Erk activation in mouse embryonic fibroblasts (MEFs)

prepared from heterozygous and homozygous Raf1D486N embryos and stimulated with various

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Figure 4-2. Phenotypes of D486N/+ and D486N/D486N mice.

(A) D486N/+ females have mild growth defect. Growth curves of male (top) and female

(bottom; p<0.0001, two-way repeated measure ANOVA) WT (n=25) and D486N/+ (n=25) mice.

(B) D486N/D486N mice with severe growth defect (s-D486N/D486N) have facial dysmorphia.

Morphometric measurements from microCT scans of a cohort of 2-month-old WT (n=12),

normal size D486N/D486N (n-D486N/D486N) (n=10) and s-D486N/D486N (n=3) male mice.

(C) Representative gross appearance and heart weight/body weight (mg/g) ratios of WT (n=15),

D486N/+ (n=20) and D486N/D486N (n=15) male mice at 4 months. (D) Surface area of isolated

neonatal cardiomyocytes from WT and D486N/D486N mice. For each genotype, 350 cells were

analyzed using ImageJ. ### p<0.001, 2-tailed Student’s t-test. (E) Left ventricular diastolic

posterior wall thickness (LVPWd) at 4 months, measured by echocardiography (n=12 for WT;

n=20 for D486N/+; n=12 for D486N/D486N). (F) Left ventricular chamber dimensions of 4

month-old WT (n=12), D486N/+ (n=20) and D486N/D486N (n=12) hearts. LVIDs, left

ventricular internal end-systolic dimension; LVIDd, left ventricular internal end-diastolic

dimension. (G) Echocardiographic parameters of 4 month-old WT (n=12), D486N/+ (n=20) and

D486N/D486N (n=12) hearts. SV, stroke volume; FS, fractional shortening; CO, cardiac output;

EF, ejection fraction. * p<0.05; ** p<0.01; *** p<0.001 (Bonferroni post-test when ANOVA is

significant). ns, not significant.

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Figure 4-3. Additional phenotyping of D486N/+ and D486N/D486N mice.

(A) Spleen weight to body weight ratios in WT, D486N/+ and D486N/D486N male mice at 5

month (n=15 for each genotype). (B) White blood counts (WBC) in WT, D486N/+ and

D486N/D486N male mice at 1 year (n=6 for each genotype). (C) Myeloid colony formation in

the absence of cytokines of bone marrow cells from WT, D486N/+, and D486N/D486N mice.

Bulk bone marrow cells were extracted at 1 year (n=3 for each genotype). Colonies were

enumerated 7 days after plating. Bone marrow from an induced Mx-Cre:KrasG12D mouse (n=1),

serves as a positive control for cytokine-independent colony formation. ns, not significant.

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agonists. As reported previously (464), Raf1L613V expressing cells show enhanced Mek/Erk

activation in response to agonists for receptor tyrosine kinases (RTKs), cytokine receptors and G

protein-coupled receptors (GPCRs). Notably, in D486N/+ MEFs, Raf1 protein levels were only

~60% of WT levels, a finding consistent with previous work showing that Raf1 kinase activity is

required to prevent ubiquitin-mediated proteolysis of Raf1 (297). Nevertheless, D486N/+ MEFs

showed sustained Mek and Erk activation in response to EGF (Figure 4-4A) stimulation.

D486N/D486N MEFs had only ~30% of WT Raf1 levels, but showed enhanced and sustained

Mek and Erk activation (Figure 4-4A). Mek and Erk activation also were enhanced in

D486N/D486N neonatal cardiomyocytes stimulated with cytokine receptor (IL-6), GPCR (Ang-

II), or RTK (IGF-I, EGF and NRG) ligands (Figure 4-4B and Figure 4-5), providing a potential

explanation for cardiac hypertrophy in D486N/D486N mice (Figure 4-2). Cardiac fibroblasts

also are implicated in hypertrophy pathogenesis (452, 466), and compared with their WT

counterparts, neonatal D486N/D486N cardiac fibroblasts showed enhanced Mek/Erk activation

in response to multiple agonists (Figure 4-4C and Figure 4-6).

4.3.4 Quantitative differences in effects of NS-associated Raf1 D486N and

L613V mutants on Mek/Erk activation

Kinase-activating (e.g., L613V) and kinase-impaired (e.g., D486N) RAF1 mutants cause

NS, but HCM is only highly associated with the former (148, 149, 467). Remarkably, our

Raf1L613V (L613V/+) (464) and Raf1D486N mouse models showed analogous genotype-dependent

phenotypic differences (Table 4-1). L613V/+ mice exhibit HCM that progresses to chamber

dilatation (464). By contrast, D486N/+ mice had no obvious cardiac phenotype, but doubling the

dosage of this kinase-defective Raf1 allele (e.g., in D486N/D486N mice) resulted in concentric

cardiac hypertrophy with decreased left ventricular end-systolic dimension and normal end-

diastolic dimension (Figure 4-2).

We asked whether these phenotypic differences correlated with differential effects on

Mek/Erk activation. Indeed, Raf1L613V caused a dramatic increase of the magnitude of EGF-

evoked Mek/Erk activation in primary MEFs, whereas Raf1D486N resulted mainly in sustained

Mek/Erk activation (Figure 4-7A). More importantly, Mek activation was enhanced only slightly

in hearts from D486N/D486N mice, whereas L613V/+ hearts showed a more profound increase

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Figure 4-4. Raf1D486N mutant increases Mek and Erk activation.

(A) Primary mouse embryo fibroblasts (MEFs) from WT, D486N/+ and D486N/D486N mice

were serum-starved for 16 hr, and then stimulated with EGF for the indicated times. Cell lysates

were immunoblotted with the indicated antibodies. A representative blot and quantification of

blots from all experiments are shown (n=6; two independent experiments using three different

MEF strains). * p<0.05; ** p<0.01; *** p<0.001 (left bottom). * p<0.05; ** p<0.01; ***

p<0.001 (right; D486N/D486N vs. WT). # p<0.05; ## p<0.01 (right; D486N/+ vs. WT);

Bonferroni post-test, when ANOVA is significant. (B) Neonatal cardiomyocytes from WT and

D486N/D486N mice were starved for 24 hr, and then stimulated for the indicated times with the

indicated agonists. AngII, Angiotensin II. NRG, heregulin-β1. Cell lysates were immunoblotted,

as indicated. (C) Cardiac fibroblasts from WT and D486N/D486N neonates were starved for 16

hr, and then stimulated for the indicated times with various agonists. Cell lysates were

immunoblotted with the indicated antibodies.

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Figure 4-5. Mek/Erk activation in WT and D486N/D486N neonatal cardiomyocytes.

Cardiomyocytes prepared from neonatal WT and D486N/D486N mice were starved for 24 hr,

and then stimulated for the indicated times with different agonists, as indicated. Mek/Erk

activation (n=2) was analyzed by immunobloting with specific antibodies and quantified using

Odyssey software.

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Figure 4-6. Mek/Erk activation in WT and D486N/D486N neonatal cardiac fibroblasts.

Cardiac fibroblasts prepared from neonatal WT and D486N/D486N mice were starved for 24 hr,

and then stimulated for the indicated times with different agonists, as indicated. Mek/Erk

activation (n=2) was analyzed by immunobloting with specific antibodies and quantified using

Odyssey software.

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WT (n=20) D486N/D486N (n=12)

L613V/+ (n=11)

LVPWd (mm) 0.73±0.01 0.87±0.03*** 0.81±0.02 *

LVIDd (mm) 4.08±0.04 4.03±0.08## 4.47±0.14**

LVIDs (mm) 2.87±0.06 2.59±0.09** # 2.96±0.12

SV (µl) 41.4±0.9 47.9±3.4** # 57.5±3.3***

CO (ml/min) 20.3±0.5 23.8±1.2** # 27.8±2.1***

FS% 29.6±0.9 35.3±1.6** 34.0±1.0*

EF% 56.8±1.4 64.3±2.2* 62.9±1.5*

dP/dt Max (mmHg/s)

10390±347 10707±690# 12486±414**

Table 4-1. Comparison of cardiac phenotypes in D486N/D486N and L613V/+ mice.

Echocardiographic and invasive hemodynamic parameters of 4 month-old WT (n=12),

D486N/D486N (n=12) and L613V/+ (n=11) hearts. LVPWD, left ventricular diastolic posterior

wall thickness; LVIDs, left ventricular internal end-systolic dimension; LVIDd, left ventricular

internal end-diastolic dimension; SV, stroke volume; FS, fractional shortening; CO, cardiac

output; EF, ejection fraction. Phenotypic data for L613V/+ mice are derived from our previous

publication (464). * p<0.05; ** p<0.01; *** p<0.001 (WT vs. L613V/+ or WT vs.

D486N/D486N). # p<0.05; ## p<0.01 (D486N/D486N vs. L613V/+); Bonferroni post-test when

ANOVA is significant.

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Figure 4-7. NS-associated Raf1 mutants differentially activate Mek/Erk.

(A) Primary WT, D486N/D486N and L613V/+ MEFs were starved for 16 hr, and then

stimulated with EGF, as indicated. Cell lysates were immunoblotted with the indicated

antibodies. The top panel shows a representative blot; quantification of blots from all

experiments (n=4) is shown below. ** p<0.01; *** p<0.001 (D486N/D486N vs. WT); ##

p<0.01; ### p<0.001 (L613V/+ vs. WT); Bonferroni post-test when ANOVA is significant. (B)

Mek activation in heart tissues. Lysates from WT (n=7), L613V/+ (n=4) and D486N/D486N

(n=4) hearts were analyzed by immunoblotting with anti-pMek antibodies with Erk2 as a loading

control. Representative samples for each genotype are shown on the left. Each lane represents an

individual animal. Quantification of all samples is shown on the right. *p<0.05 (2-tailed

Student’s t-test).

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in Mek activation (Figure 4-7B). Although Raf1L613V and Raf1D486N enhanced Mek/Erk

activation in neonatal cardiomyocytes and cardiac fibroblasts, their effects on the response to

various stimuli differed (Figure 4-8). In some cases (e.g., IL-6 and FGF2), Raf1L613V and

Raf1D486N had similar effects on pathway activation, but in many others (e.g., AngII, NRG, EGF,

PDGF), Raf1L613V had a much stronger effect. Both the magnitude and the duration of Mek/Erk

activation were affected differentially by Raf1L613V and Raf1D486N. Such differences likely reflect

important, if sometimes subtle, differences in the regulation of the RAS/ERK pathway in

response to distinct agonists and in different cell types. However, given that Mek inhibitor

treatment reverses HCM in L613V/+ mice (464), these data are consistent with the idea that the

ability of kinase-activating Raf1 mutants to more profoundly enhance Mek/Erk pathway

activation underlies their distinct phenotypic effects (see Discussion).

4.3.5 Kinase-impaired Raf1 mutants enhance Mek/Erk activation by promoting

heterodimerization with Braf

To study the biochemical properties of RAF1 mutants in a more biochemically tractable

cell system, while avoiding the marked over-expression seen in transient transfection

experiments, we generated stable mammalian Flp-In T-REx 293 (T-REx 293) cell lines. Such

cells allow tetracycline-inducible expression of Flag-tagged WT and mutant RAF1 from the

same genomic locus at levels comparable to endogenous RAF1. As in primary MEFs,

cardiomyocytes and cardiac fibroblasts, expression of RAF1D486N in T-REx 293 cells resulted in

increased MEK/ERK activation, compared with the effects of RAF1WT expression (Figure 4-9A).

Transient transfection studies have shown that RAF1 can form heterodimers with BRAF, that

RAF1/BRAF heterodimers have increased kinase activity compared with the respective

homodimers or monomers, and that a single kinase-competent RAF isoform can confer high

catalytic activity to the heterodimer (168, 298, 302, 303). These findings suggested that

RAF1D486N might enhance MEK/ERK activation by promoting heterodimer formation. To test

this possibility, we immunoprecipitated Flag-tagged RAF1 from induced T-REx 293 cell lysates,

and subjected the immunoprecipitates to immunoblotting with anti-BRAF antibodies. Following

EGF stimulation, low levels of RAF1/BRAF heterodimers were found in WT RAF1-expressing

T- REx 293 cell lysates. By contrast, heterodimerization was enhanced dramatically in

RAF1D486N-expressing cells, even though, as in MEFs, RAF1D486N accumulated to levels

considerably lower than WT RAF1 (Figure 4-9A). Increased heterodimerization was not due to

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Figure 4-8. Severity of cardiac phenotype in D486N/D486N and L613V/+ mice correlates

with Mek/Erk activation.

Quantification of Mek/Erk activation from immunoblots (n=2) of cardiomyocytes (A) and

cardiac fibroblasts (B). Lysates from neonatal WT and D486N/D486N or WT and L613V/+

cells, starved for 24 hr and then stimulated as indicated. Values indicate fold change in Mek/Erk

activity (assessed by immunoblotting with phospho-specific antibodies) in D486N/D486N or

L613V/+ cells, compared with the corresponding WT control at 5 min post-stimulation.

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defective negative feedback (244), as heterodimerization was increased even when ERK

activation was blocked by MEK inhibitor treatment (Figure 4-9B). These results were confirmed

using primary D486N/+ and D486N/D486N MEFs, despite their markedly lower expression of

Raf1D486N (Figure 4-9C). Furthermore, enhanced ability to form RAF1/BRAF heterodimers was a

common feature of kinase-impaired, NS-associated RAF1 mutants: the T491I and T491R kinase

domain mutants also showed enhanced heterodimerization, and increased and sustained

MEK/ERK activation in EGF-stimualted T-REx 293 cells (Figure 4-10).

4.3.6 Heterodimerization with BRAF is required for RAF1D486N to enhance

MEK/ERK activation

We next asked whether heterodimerization with BRAF is required for enhanced

MEK/ERK activation in these cells. Indeed, infection of T-REx 293 cells expressing RAF1D486N

with a lentivirus expressing BRAF shRNA abolished MEK/ERK hyperactivation in these cells

(Figure 4-11A). Similar experiments were performed on primary D486N/D486N MEFs. Again,

Mek activation and Raf1/Braf heterodimer levels were reduced significantly in Braf knock-down

cells (Figure 4-11B and Figure 4-11C). Surprisingly, Erk activation was not affected, although

this could reflect significant up-regulation of Erk1 protein levels in Braf knock-down MEFs

(Figure 4-11B).

Mammals express three RAF family members, RAF1, BRAF and ARAF (252), all of

which share MEK1/2 as substrates. Co-immunoprecipitation experiments using lysates from

D486N/D486N MEFs showed that Araf also was present in Raf1 immunoprecipitates (Figure 4-

11D). However, Araf knockdown did not reduce Mek activation significantly in these cells,

compared with the marked effects of Braf depletion. Braf knockdown did cause enhanced

Araf/Raf1 heterodimerization, which led us to test whether heterodimerization with Araf helps to

explain persistent Mek/Erk activation in Braf knockdown cells. As expected, combined depletion

of Araf and Braf in these cells further impaired Mek/Erk activation. Taken together, these results

suggest that at least in MEFs, Raf1D486N enhances Mek/Erk activation primarily by promoting

heterodimerization with Braf, but raise the possibility that Araf/Raf1 heterodimers might

contribute to the effects of Raf1D486N in other cell types.

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Figure 4-9. RAF1D486N forms more heterodimers with BRAF.

(A) T-REx 293 cell lines expressing Flag-tagged human RAF1 WT or D486N were incubated

with 1µg/ml Tetracycline for 20 hours in serum-free medium, and then stimulated with EGF for

the indicated times. Heterodimers were detected by immunoprecipitation with anti-Flag

antibody, followed by blotting for endogenous BRAF. MEK and ERK activation in the same

lysates were assessed by immunoblotting. (B) T-REx 293 cell lines expressing Flag-tagged

human RAF1 WT or D486N were incubated with 1µg/ml Tetracycline for 20 hours in serum-

free medium, and then treated with 10µM PD0325901 for 1 hr before stimulation with EGF for

the indicated times. Heterodimers were detected by immunoprecipitation with anti-Flag

antibody, followed by blotting for BRAF. Total cell lysates from the same experiment were

immunoblotted with the indicated antibodies. (C) Primary MEFs from WT, D486N/+ and

D486N/D486N mice were starved for 16 hr, and then stimulated with EGF, as indicated.

Endogenous heterodimers were detected by immunoprecipitation with RAF1 antibody, followed

by blotting for Braf. Mek and Erk activation in the same lysates were assessed by

immunoblotting.

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Figure 4-10. Kinase-impaired RAF1 mutants associated with NS enhance MEK/ERK

activation and form more RAF1/BRAF heterodimers.

(A) T-REx 293 cell lines expressing Flag-tagged human RAF1 WT, D486N, T491I or T491R

were incubated with 1µg/ml Tetracycline for 20 hours in serum-free medium, and then

stimulated with EGF. Cell lysates from a representative experiment were subjected to

immunoblotting with the indicated antibodies. Quantification of blots from all experiments (n=4)

is shown on the right. *** p<0.001 (WT vs. D486N in blue; WT vs. T491I in red; WT vs. T491R

in green); Bonferroni post-test when ANOVA is significant. (B) T-REx 293 cell lines expressing

Flag-tagged human RAF1 WT or the indicated mutants were incubated with 1µg/ml Tetracycline

for 20 hours in serum-free medium, and then stimulated with EGF. Heterodimers were detected

by immunoprecipitation with anti-Flag antibody, followed by immunoblotting for BRAF. The

same lysates were immunoblotted with the indicated antibodies.

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Figure 4-11. BRAF is required for RAF1D486N to enhance MEK/ERK activation.

(A) T-REx 293 cell lines expressing Flag-tagged human WT or D486N mutant RAF1 were

infected with a lentivirus expressing BRAF shRNA (BRAF-KD) or a control (Luciferase) shRNA

(shluc). At 48 hours post-puromycin selection, cells were incubated with 1µg/ml Tetracycline for

20 hours in serum-free medium, and then stimulated with EGF. Cell lysates were immunoblotted

with the indicated antibodies. A representative experiment is shown at the left, and blots from all

experiments (n=3) are quantified on the right. * p<0.05; ** p<0.01; *** p<0.001 (WT-shluc vs.

D486N-shluc); ### p<0.001 (D486N-shluc vs. D486N-BRAF-KD); Bonferroni post-test when

ANOVA is significant. (B) Primary MEFs from D486N/D486N mice were infected with a

lentivirus expressing Braf shRNA (Braf-KD) or a control shRNA (shluc). Cell lysates were

immunoblotted with the indicated antibodies. A representative blot is shown on the top (arrow

indicates position of Erk1). Quantification of blots from all experiments (n=4) for Mek activation

is shown at the bottom. *** p<0.001; Bonferroni post-test when ANOVA is significant. (C)

Primary WT and D486N/D486N MEFs were infected with a lentivirus expressing Braf shRNA

(Braf-KD) or a control shRNA (shluc). Heterodimers were detected by immunoprecipitation with

RAF1 antibody, followed by immunoblotting for BRaf. Total cell lysates were immunoblotted

with the indicated antibodies. (D) Primary D486N/D486N MEFs were infected with lentiviruses

expressing shRNAs against Braf (Braf-KD) and/or Araf (Araf-KD) or a control shRNA (shluc).

Heterodimers were detected by immunoprecipitations with RAF1 antibody, followed by blotting

for Braf or Araf. Total cell lysates also were immunoblotted with the indicated antibodies.

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Although these experiments showed that Braf, and to a lesser extent, Araf, is necessary

for the effects of Raf1D486N, and that these effects correlate with the ability of this mutant to

increase heterodimerization, they did not establish a causal relationship between

heterodimerization and Mek/Erk hyperactivation. Structural studies of Drosophila Raf (299)

showed that Arg 481 (Arg401 in RAF1 and Arg509 in BRAF) is at the center of a side-to-side

RAF dimer interface, and directly participates in these interactions (Figure 4-12A). As in

Drosophila Raf, introduction of the R401H mutation into WT RAF1 or RAF1D486N abolished

RAF1/BRAF heterodimerization (Figure 4-12B). Moreover, the compound R401H/D486N

mutant no longer enhanced EGF-evoked MEK/ERK activation (Figure 4-12C). Similar results

were obtained when we tested another mutation (F408A) within the dimer interface (Figure 4-

12A and Figure 4-12D). Furthermore, re-expression of Myc-tagged WT Braf in Braf knock-

down D486N/D486N MEFs restored Mek and Erk activation, whereas BrafR509H could not

rescue (Figure 4-12E).

Finally, to test unambiguously the effects of dimerization per se (as opposed to other,

unanticipated structural consequences of dimer-interface mutants), we asked if forced

heterodimerization could restore the ability of RAF1R401H/D486N to promote MEK/ERK

activation. We fused Flag-RAF1R401H/D486N, which cannot form heterodimers with BRAF, to

FKBP, while FRB was fused to BRAF (Figure 4-13A). Upon co-transfection, these two proteins

can heterodimerize only upon addition of a rapamycin analog (A/C heterodimerizer).

Remarkably, MEK and ERK activation were restored upon A/C heterodimerizer addition (Figure

4-13B); importantly, the heterodimerizer itself had no effect (Figure 4-13C). Taken together,

these results show that heterodimerization with BRAF is necessary and sufficient for the NS-

associated RAF1D486N mutant to enhance MEK/ERK activation.

Surprisingly, we also noticed that the kinase-activating mutant RAF1L613V also formed

more RAF1/BRAF heterodimers in response to growth factor stimulation (Figure 4-12B and

Figure 4-14A) and, as for the kinase-defective Raf1 alleles, knock-down of Braf in primary

MEFs expressing Raf1L613V significantly reduced Mek activation in these cells (data not

shown). Moreover, the compound L613V and dimer mutant (R401H) abolished RAF1/BRAF

heterodimerization and no longer caused enhanced MEK/ERK activation (Figure 4-12B and

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Figure 4-12. Heterodimerization with BRAF is required for RAF1D486N mutant to enhance

MEK/ERK activation.

(A) Side-to-side dimer interface in Drosophila Raf (299). One protomer is displayed as a surface

representation in orange, while the other is shown as a ribbon representation in violet. The inset

shows a close-up of hydrogen bonding interactions involving R481 and F488 (R401 and F408 in

human RAF1). (B) T-REx 293 cell lines expressing Flag-tagged human RAF1 WT or the

indicated mutants were incubated with 1µg/ml Tetracycline for 20 hours in serum-free medium,

and then either left unstimulated (-) or stimulated with EGF for 5 min (+). Heterodimers were

detected by immunoprecipitation with anti-Flag antibody, followed by immunoblotting with

BRAF antibodies. Total cell lysates also were immunoblotted with the indicated antibodies. (C)

T-REx 293 cell lines expressing Flag-tagged human RAF1 WT or the indicated mutants were

incubated with 1µg/ml Tetracycline for 20 hours in serum-free medium, and then stimulated with

EGF for the times indicated. Cell lysates were immunoblotted with the indicated antibodies. A

representative immunoblot is shown, and quantification of the blots from all experiments (n=4) is

shown at the bottom. * p<0.05; ** p<0.01; *** p<0.001 (WT vs. D486N). ### p<0.001

(R401H/D486N vs. WT); Bonferroni post-test when ANOVA is significant. (D) T-REx 293 cell

lines expressing Flag-tagged human RAF1 WT, D486N and/or F408A were incubated with

1µg/ml Tetracycline for 20 hours in serum-free medium, and then stimulated with EGF for the

times indicated. Cell lysates were immunoblotted with the indicated antibodies. (E) Immortalized

D486N/D486N MEFs were infected with a lentivirus expressing Braf shRNA. At 72 hours post-

puromycin selection, cells were infected with MCSV-based retroviruses expressing GFP alone or

GFP plus myc-BRAFWT or myc-BRAFR509H. GFP+ cells, obtained by FACS, were serum-starved

for 16 hours, and then stimulated with EGF. Cell lysates were immunoblotted with the indicated

antibodies.

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Figure 4-13. Induced RAF1/BRAF heterodimerization restores activity of RAF1R401H/D486N

mutant to enhance MEK/ERK activation.

(A) RAF1R401H/D486N and BRAF coding sequences were sub-cloned into FKBP or PRB expression

vectors, respectively, as shown. (B) T-REx 293 cells were co-transfected with Flag-

RAF1R401H/D486N-FKBP and BRAF-FRB-HA expression plasmids for 24 hours, and then left

untreated (0h), or treated with A/C heterodimerizer (500nM) for the indicated times. Cell lysates

were immunoblotted with the indicated antibodies. (C) T-REx 293 host cells were co-transfected

with Flag-RAF1D486N or Flag-RAF1R401H/D486N and Myc-BRAF expression plasmids, and 24 hours

later were left untreated (-), or treated with A/C heterodimerizer (500nM) (+) for 30 min. Cell

lysates were immunoblotted with the indicated antibodies.

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Figure 4-14. RAF1L613V mutant enhances MEK/ERK activation via RAF1/BRAF

heterodimer formation.

(A) Primary WT, L613V/+ and L613V/L613V MEFs were starved for 16 hr, and then stimulated

with EGF for the indicated times. Endogenous heterodimers were detected by

immunoprecipitation with RAF1 antibody, followed by blotting for Braf. Total cell lysates from

the same experiments were immunoblotted with the indicated antibodies. (B) T-REx 293 cell

lines expressing Flag-RAF1 WT, L613V or R401H/L613V were incubated with 1µg/ml

Tetracycline for 20 hours in serum-free medium, and then stimulated with EGF for the times

indicated. MEK and ERK activation were assessed by immunoblotting with the indicated

antibodies. A representative immunoblot is shown, and quantification of the blots from all

experiments (n=3) is shown on the right. * p<0.05; *** p<0.001 (WT vs. L613V). ## p<0.01;

### p<0.001 (R401H/L613V vs. WT); Bonferroni post-test when ANOVA is significant.

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Figure 4-14B). These data provide an explanation for the ability of this class of kinase-activated

alleles to promote MEK/ERK hyper-activation.

4.4 Discussion

We generated and analyzed a knock-in mouse model for the NS-associated, kinase-

impaired Raf1D486N mutation. Unlike the kinase-activating mutation Raf1L613V (464), which

causes most major features of NS, including proportional short stature, facial dysmorphia,

hematological defects and HCM, Raf1D486N heterozygosity results only in a mild growth defect in

female mice. Raf1D486N homozygous mice (on a 129Sv × C57BL/6 mixed background) exhibit

concentric cardiac hypertrophy (but not HCM) and an incompletely penetrant severe growth

defect that is accompanied by facial dysmorphia, failure to thrive and early death. Detailed

analysis of the mechanism underlying the effects of this Raf1 allele, as well as other kinase-

defective RAF1 mutants, indicate that they paradoxically hyper-activate the RAS/ERK pathway

by promoting heterodimerization with BRAF and, to a lesser extent, ARAF. Finally, we

unexpectedly found that Raf1L613V, whose activation mechanism had remained unclear, also

promotes increased heterodimer formation. Taken together, these data identify increased

heterodimerization capacity as the common theme in the pathogenesis of RAF1 mutant-

associated NS.

The incompletely penetrant growth/facial dysmorphia/viability phenotype of

D486N/D486N mice suggests the existence of modifier gene(s) that vary between the 129Sv and

C57BL/6 strains. Indeed, preliminary mapping studies have identified a 129Sv locus on mouse

chromosome 8 that is strongly linked (LOD score ~15) to the s-D486N/D486N phenotype (see

5.2 Future directions). Like s-D486N/D486N mice, Araf-deficient mice have severe growth

defects, neurological abnormalities and post-natal lethality (234); notably, these phenotypes also

are sensitive to genetic background. The identification of loci that modify Raf1 function could

provide important insights into the regulation of Raf-dependent signaling, and it will be

interesting to see if the same modifier(s) affect the Raf1D486N and Araf mutant phenotypes.

Primary lung fibroblasts prepared from s- and n- D486N/D486N mice show comparable

Mek/Erk activation (data not shown), suggesting that the putative modifier(s) likely acts

downstream of Erk or parallel to the RAS/ERK pathway. Our previous NS mouse models,

Shp2D61G (204) and Raf1L613V (464), also show strain-specific differences in phenotype.

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However, different modifiers are likely to be involved, because increasing C57BL/6 content

causes lethality in these two mouse models. Given the marked genetic heterogeneity of the

human population, and the differences in penetrance of RASopathy phenotypes within a single

affected family, not to mention between unrelated patients with the same mutant allele, our

mouse models provide excellent opportunities to investigate the relative effects of RASopathy

mutations and genetic background.

Kinase-activating RAF1 mutants are strongly associated with HCM in human NS

patients, a phenotype reproduced by our Raf1L613V mouse model (464). Similarly, D486N/+ mice,

like NS patients with kinase-impaired RAF1 mutations, do not exhibit HCM. Notably, Raf1D486N

homozygosity (which is not seen in humans) results in concentric cardiac hypertrophy that does

not progress to heart failure within one year of observation (Figure 4-2), unlike the eccentric

cardiac hypertrophy seen in L613V/+ mice (464). Biochemical analysis revealed that, in contrast

to initial reports based on transient transfection experiments (149), expression of kinase-impaired

Raf1D486N at normal (endogenous) levels causes sustained and/or enhanced Mek/Erk activation in

multiple cell types in response to a variety of stimuli. In order to transmit signals downstream,

RAF proteins must assemble into multi-protein complexes (252). Presumably, the substantial

over-expression that occurs in transiently transfected cells results in artifactual dominant

negative effects of kinase-deficient RAF1 mutants that are not seen when these proteins are

expressed at appropriate, endogenous levels. Importantly, however, Raf1D486N is less potent than

Raf1L613V in enhancing Mek/Erk activation in response to most stimuli, both in cardiomyocytes

and in cardiac fibroblasts (Figure 4-8). Furthermore, Raf1L613V generally increases the magnitude

of Mek/Erk activation, whereas Raf1D486N most often prolongs signal duration. Given our

previous finding that HCM can be prevented or reversed by MEK inhibitor treatment of

Raf1L613V/+ mice, these data strongly suggest that differences in the ability of the D486N and

L613V mutants (and by inference, between kinase-impaired and kinase-activated RAF1 in

general) to promote Mek/Erk activation in the heart explains their distinct cardiac phenotypes. If

so, then agonists that show differential effects on Mek/Erk pathway activation in Raf1L613V/+ and

Raf1D486N/D486N cardiomyocytes (e.g., AngII, NRG, but not IL6) and/or cardiac fibroblasts (e.g.,

EGF, IGF, PDGF, but not FGF2) might be particularly important for cardiac hypertrophy in NS.

Raf1D486N accumulates to much lower levels than Raf1L613V, consistent with a previous report

that kinase activity is required to prevent Raf1 degradation (297). Consequently, we cannot

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ascertain whether lower levels of the Raf1D486N protein or its lower kinase activity accounts for

the decreased Mek/Erk hyperactivation seen in D486N/D486N, compared with L613V/+, mice

(also, see below).

The carboxyl oxygen of the highly conserved aspartic acid 486 (the “D” of the DFG

motif) plays a critical role in chelating Mg2+ and stabilizing ATP binding in the active site of

protein kinases (468). Mutation of this residue to alanine creates a kinase-inactive protein (297),

whereas mutation to asparagine severely impairs (but does not eliminate) activity (149). All RAF

family members require phosphorylation of key activation loop residues for maximal kinase

activation (244, 469), and T491 is one of these residues. As expected, NS-associated mutants

affecting T491 also show impaired kinase activity (149).

Our results clearly establish how all such mutants (D486N, T491I and T491R) enhance

downstream MEK/ERK activation. Previous studies showed that RAF1/BRAF

heterodimerization occurs during normal growth factor signaling, and indicated that

heterodimers have increased catalytic activity compared with homodimers or monomers (298).

Although some have argued that RAF1 inhibits BRAF activation in a kinase domain-dependent

fashion (470), other studies reported that human cancer-associated BRAF mutants with impaired

kinase activity promote MEK activation by binding to and trans-activating RAF1 (167, 168). Of

note, these kinase-impaired mutations alter the activation segment of BRAF, and some (D594G

and T599I) are analogous to NS-associated RAF1 mutants (471, 472). Marais and colleagues

recently reported that mice with a conditional kinase-dead Braf (BrafLSL-D594A) allele, when

combined with oncogenic Ras (KrasG12D), induced melanomas in mice. In tumor cells from these

mice, BrafD594A was bound to Raf1 constitutively (303). By analogy, we suspected that kinase-

impaired, NS-associated RAF1 mutants hyperactivated MEK/ERK by promoting

heterodimerization with BRAF. Indeed, we found that all of the kinase-impaired RAF1 mutants

form more heterodimers with BRAF upon growth factor stimulation (Figure 4-9 and Figure

4-10). Furthermore, the level of MEK/ERK hyperactivation depends on the level of RAF1/BRAF

heterodimerization, as RAF1T491I and RAF1T491R formed more heterodimers and caused more

MEK/ERK activation than did RAF1D486N (Figure 4-10).

Several subsequent lines of evidence establish a causal relationship between

heterodimerization and MEK/ERK activation. First, knockdown of BRAF in T-REx293 cells

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expressing Flag-RAF1D486N reduces MEK/ERK activation. Braf knockdown in primary

D486N/D486N MEFs also impairs Mek activation, although Erk activation is unaffected (Figure

4-11B). This difference might be more apparent than real, however, as total Erk1 levels are

increased in Braf-knockdown MEFs, suggesting potential compensation during the selection of

stable knockdown clones. Such feedback regulation presumably is impaired (or less effective) in

T-REx 293 cells. Second, mutation of either of two key residues (R401H, F408A) in the dimer

interface of the RAF1 kinase domain, as revealed by the crystal structures of BRAF (167) and

Drosophila Raf (299), abolishes the ability of RAF1D486N to enhance EGF-evoked MEK/ERK

activation (Figure 4-12). Most compellingly, forced heterodimerization of Raf1D486N/R401H and

BRAF by means of an inducible FKBP-FRB interaction system restored the ability of

Raf1D486N/R401H to enhance EGF-evoked MEK/ERK hyperactivation (Figure 4-13). The latter

result shows unambiguously that Raf1D486N/R401H cannot hyperactivate MEK/ERK

hyperactivation solely because it has lost its ability to promote RAF1/BRAF heterodimerization,

as opposed to some unanticipated effect of the second site R401H mutation on the conformation

of the D486N mutant. Taken together, our results demonstrate that heterodimerization with

BRAF is necessary and sufficient for such mutants to enhance MEK/ERK activation. Although

our knockdown studies show that RAF1D486N enhances MEK/ERK activation (at least in MEFs

and T-REx 293 cells) mainly through heterodimerization and transactivation of BRAF, we also

find that kinase-defective RAF1 mutants can heterodimerize with ARAF. Consequently, we do

not exclude the possibility that RAF1/ARAF heterodimers play an important role in the

pathogenesis of NS caused by kinase-defective RAF1 alleles in tissues where ARAF is a major

isoform (e.g., the brain).

Another cluster of NS-associated RAF1 mutations maps to the C-terminus (S612 and

L613) of RAF1. Like NS-associated CR2 domain mutants, these mutants have increased kinase

activity (149), even though S612 and L613 lie distal to the RAF1 kinase domain. Previous

studies have not implicated these residues in RAF1 regulation, so it has been unclear how such

mutations enhance kinase activity. Moreover, phosphorylation of Ser259 (an inhibitory binding

site for 14-3-3), and Ser 621 (149), a 14-3-3 binding site that promotes RAF1 activation (297),

are unaffected in RAF1L613V. We found that, compared with WT RAF1, RAF1L613V also forms

more heterodimers with BRAF upon growth factor stimulation. Moreover, superimposing the

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dimer interface mutant (R401H) on L613V blocked its ability to enhance MEK/ERK activation

(Figure 4-14).

These data strongly suggest that RAF1L613V also promotes MEK/ERK hyper-activation

via enhanced BRAF heterodimerization ability. As the kinase domain of RAF1L613V is not

altered, and this mutant is expressed at normal levels unlike kinase-deficient RAF1 alleles,

heterodimerization with BRAF presumably results in a greater increase in MEK/ERK activation,

compared with that evoked by kinase-impaired RAF1 alleles. The NS-associated CR2 domain

mutant RAF1S257L and RAF1P261T (98) (data not shown) also forms more heterodimers with

BRAF. Taken together, these results argue that increased ability to heterodimerize with BRAF

(and possibly ARAF) represents a general pathogenic mechanism for NS-associated RAF1

mutations, and suggest that agents that interfere with dimerization might have general utility for

the treatment of RAF1 mutant NS.

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Chapter 5

Summary and future directions

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5.1 Summary and Key Findings

NS is one of several autosomal dominant “RASopathies” caused by mutations in

components of the RAS/ERK pathway. Germ line mutations in RAF1 account for ~3-5% of NS,

and unlike other NS alleles, RAF1 mutations that confer increased kinase activity are highly

associated with hypertrophic cardiomyopathy (HCM). Surprisingly, some NS-associated RAF1

mutations show normal or decreased kinase activity. These observations raised the question of

why both kinase-activating and kinase-impaired RAF1 mutants can cause similar diseases, and

how NS-associated RAF1 mutations cause HCM. In this thesis, these issues were addressed by

generating and analyzing two lines of “knock-in” mice that express kinase-activating (L613V)

and impaired (D486N) RAF1 mutants, respectively.

In Chapter 3, I described a knock-in mouse model for NS caused by the kinase-

activating Raf1L613V mutation. Similar to mouse models of Ptpn11 mutation-associated NS,

Raf1L613V heterozygosity (L613V/+) causes proportional short stature, facial dysmorphia, and

hematological defects. Unlike phosphatase-activating Ptpn11 alleles, which cause valvuloseptal

abnormalities, L613V/+ mice have normal valvuloseptal development and instead exhibit

eccentric cardiac hypertrophy that decompensates upon pressure overload. Agonist-evoked

Mek/Erk activation is enhanced in multiple cell types without changes in several other signaling

pathways implicated in cardiac hypertrophy/HCM. Remarkably, post-natal MEK inhibition

normalizes the growth, facial and cardiac defects in L613V/+ mice, demonstrating that elevated

MEK/ERK activity is critical for causing HCM and other NS phenotypes and identifying MEK

inhibitors as potential therapeutic agents for the treatment of NS.

Whether kinase-defective Raf1 alleles also faithfully model human NS, and if so, how

kinase-activating and kinase-defective mutants can cause similar phenotypes, remained to be

resolved. In Chapter 4, I generated and analyzed knock-in mice expressing the kinase-impaired

NS mutant Raf1D486N, and also re-examined the effects of various other NS mutants expressed at

more physiological levels than in previous transient transfection studies. Unlike the kinase-

activating mutation Raf1L613V, which causes most major features of NS, Raf1D486N heterozygosity

results in only a mild growth defect in female mice. Raf1D486N homozygous mice (on a 129Sv ×

C57BL/6 mixed background) exhibit concentric cardiac hypertrophy (but not HCM) and an

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incompletely penetrant severe growth defect that is accompanied by facial dysmorphia, failure to

thrive and early death. Detailed analysis of the mechanism underlying the effects of this Raf1

allele, as well as other kinase-defective RAF1 mutants, indicate that they paradoxically hyper-

activate the RAS/ERK pathway by promoting heterodimerization with BRAF and, to a lesser

extent, ARAF. Finally, I unexpectedly found that Raf1L613V also promotes increased heterodimer

formation. Taken together, these data identify increased heterodimerization capacity as the

common theme in the pathogenesis of RAF1 mutant-associated NS.

5.2 Future Directions

5.2.1 Optimizing therapy for NS-associated HCM

RASopathies are rare, “orphan diseases”, so extensive pre-clinical evaluations are needed

to prioritize potential therapeutic approaches for these patients. Post-natal therapy for 6 weeks

with the MEK inhibitor PD0325901 (PD) rescues all NS phenotypes, including HCM, in

L613V/+ mice (Figure 3-17, Figure 3-19 and Figure 3-20; (464)). Whether the dose used for our

initial experiments (5mg/kg BW/day) is optimal is uncertain, nor is it clear whether HCM recurs

upon drug withdrawal. Furthermore, obesity was an unanticipated side effect of PD treatment.

Therefore, it is worthy to perform more extensive pre-clinical studies with PD treatment on our

NS mouse models, to ask whether its body mass effects are on- or off-target by testing a

structurally unrelated MEK inhibitor, and to test whether other agents can be used for L613V/+

HCM.

Our initial MEK inhibitor treatment studies were done with PD at 5mg/kg BW/day, a

dose below that used in most mouse xenograft tumor models of this agent for cancer therapy

(453). In humans, PD has side effects such as skin rash, diarrhea, fatigue, nausea, and visual

disturbances (473, 474) that may be tolerable for treatment of patients with advanced cancers,

but not for a developmental disorder such as NS, particularly if long term, continuous treatment

is required. Although we did not observe severe side effects except weight gain in our original

study, we would nevertheless like to use the lowest effective dose in patients. Therefore, I

performed a lower dose MEK inhibitor treatment (3mg/kg BW/day PD) in our L613V/+ NS

mouse model, beginning at 4 weeks of age and continuing for 6 weeks, as in our original study

(464). Mice were weighed and measured weekly. At the end of the treatment period, the hearts of

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these mice were analyzed by echocardiography. The results showed that the lower dose of MEK

inhibitor could still rescue the growth defect in L613V/+ mice (Figure 5-1A), although it takes

longer for L613V/+ mice to catch up with the WT controls (Figure 3-17). The increased body

mass effect also was reduced by decreasing the dose of PD (Figure 5-1B). However, at this dose,

MEK inhibitor treatment could not rescue the cardiac hypertrophy, nor the chamber dilatation of

the hearts (Figure 5-1C, D and E). Interestingly, MEK inhibitor treatment did caused a mild but

significant increase in LVIDs (Figure 5-1F), which is consistent with a decrease in FS and EF in

L613V/+ mice after treatment (Figure 5-2). However, other parameters of cardiac function (SV

and CO) did not change upon lower dose MEK inhibitor treatment (Figure 5-2). These data

suggest that there may be a different threshold of the level of Mek/Erk activation for causing the

growth defects and HCM in NS, and/or that an off-target effect of PD might contribute to the

rescue of HCM. Either way, we cannot lower the dose of PD in order to fully rescue the HCM in

our L613V/+ NS mouse model.

Future work should investigate whether growth retardation or HCM recur after drug

cessation. In order to test this, the same general design can be used as in our original study (464),

except that body size would be measured continuously post-treatment, and echocardiography

will be performed both at the end of the treatment period and at different time points post-

withdrawal. Another interesting question is whether we can rescue NS phenotypes in adult mice

(e.g., starting treatment at 10 weeks of age or later) instead of treating the mice at puberty as in

our original study (464) by MEK inhibitor treatment. All of these findings would provide

important guidelines for potential clinical treatment of NS patients with MEK inhibitors.

Finally, PD treatment causes a rapid and significant increase in body mass/body fat, an

obviously undesirable side effect. This could be an on-target effect of Mek-inhibition; notably,

deletion of Ptpn11 in the forebrain causes marked obesity (461), and Shp2, of course, is a major

regulator of Erk activation. Alternatively, it could be an idiosyncratic effect of PD. To

distinguish between these possibilities, we can perform similar studies to those described above

using the structurally unrelated MEK inhibitors, such as the compound ARRY162, which is

available commercially.

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Figure 5-1. Lower dose MEK inhibitor treatment rescues growth defect, but not cardiac

hypertrophy and chamber dilatation in L613V/+ mice.

Mice were injected IP daily with PD0325901 (PD; 3mg/kg) or vehicle, starting at 4 weeks of age

and for the succeeding 6 weeks. Body length (A) and body weight (B) were measured weekly. #

p<0.05, ### p<0.0001, two-way repeated-measures ANOVA; * p<0.05, Bonferroni post-test

when ANOVA was significant (black symbols, WT PD vs. WT control; red symbols, L613V/+

PD vs. L613V/+ control). (C) Heart weight/body weight (HW/BW) ratio, (D) LVPWd and (E)

LVIDd were not restored to within normal limits in lower dose MEK inhibitor-treated mice. (F)

LVIDs increased after MEK inhibitor treatment. LVPWd, LVIDd and LVIDs were also

normalized by BW1/3. ns, not significant. # p<0.05, ## p<0.005 1-tailed Student’s t test; *

p<0.05, ** p<0.005, *** p<0.0001, Bonferroni post-test when ANOVA was significant. n= 9

(WT); 10 (WT PD); 10 (L613V/+); 11 (L613V/+ PD).

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Figure 5-2. Lower dose MEK inhibitor treatment partially normalizes cardiac function in

L613V/+ mice.

Echocardiographic parameters at 4 month of age. ns, not significant. * p<0.05, ** p<0.005, ***

p<0.0001, Bonferroni post-test when ANOVA was significant. n= 9 (WT); 10 (WT PD); 10

(L613V/+); 11 (L613V/+ PD).

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Although PD treatment reverses all of the phenotypes in Raf1 mutant NS, no MEK

inhibitor is approved for use in children (or adults); therefore, it would be difficult to bring MEK

inhibitor therapy to the clinic quickly. Besides increased Mek/Erk activation, S6 phosphorylation

also is enhanced in Raf1 mutant NS (Figure 5-3). S6 can be a downstream target of the Erk target

Rsk (107); although p70S6K activation (and presumably mTorc1 activity) is normal in these

mice (Figure 3-16), reducing mTorc1 activity with Rapamycin might be effective for Raf1

mutant HCM. Rapamycin is approved for the pediatric population, and could be tested in NS

patients with HCM more rapidly than a MEK inhibitor. In order to test this hypothesis, the same

general design was used as in our original MEK inhibitor treatment study (464), and Rapamycin

(2 mg/kg BW/day), a dose that reverses HCM in LS mice (404), was used to treat L613V/+ mice.

Our preliminary data showed that, at this dose, rapamycin treatment caused significant growth

retardation and weight loss in both L613V/+ and WT control mice (Figure 5-4A and B), which

forced us to terminate the treatment. It could not rescue the cardiac hypertrophy after 4 weeks of

treatment (Figure 5-4C). These data suggest that excess S6 phosphorylation is not a major part of

HCM pathogenesis in NS. Alternatively, many Erk-dependent pathways might have to be

reversed to ameliorate Raf1 mutant HCM.

5.2.2 Downstream target(s) of the RAS/ERK signaling in NS and HCM

Many pathways are implicated in HCM pathogenesis (Figure 1-14), in addtition to the

RAS/ERK pathway demonstrated by our studies, yet the key targets of excessive Erk activation

in HCM pathogenesis remain to be determined. Specific transcription factors can mediate cardiac

hypertrophy. For example, Gata4 activates hypertrophy-associated genes and causes

cardiomyopathy, in part via Nfat family members, themselves key transducers of the

hypertrophic response (392). Gata4 interacts with Nfat2 and Nfat3 to strengthen DNA binding,

thereby synergistically activating endothelin-1 (475) and BNP (476) transcription. Notably,

Gata4 is an Erk1/2 target (419, 477). Therefore, these transcription factors are attractive

candidates for mediating the hypertrophic effects of NS-associated Raf1. To test the possible

involvement of Gata4 in Raf1 mutant HCM, we can assess phosphorylation at its Erk site, S105,

by immunoblotting. We also can cross L613V/+ mice to knock-in mice expressing Gata4S105A,

which lack the Erk-dependent phosphorylation site, and assess whether HCM can be attenuated

in these mice.

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Figure 5-3. Increased pS6 (S235/236) in cardiomyocytes and cardiac fibroblasts from

L613V/+ mice.

(A) Neonatal cardiomyocytes were starved for 24 hr, and then stimulated as indicated with AngII

or IL-6. (B) Neonatal cardiac fibroblasts were starved for 16 hr, and then stimulated as indicated

with EGF or IGF-I. ERK2 serves as a loading control.

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Figure 5-4. Rapamycin treatment does not rescue growth defect and cardiac hypertrophy

in L613V/+ mice.

Mice were injected IP daily with Rapamycin (Rapa; 2mg/kg) or vehicle, starting at 4 weeks of

age and for the succeeding 4 weeks. Body length (A) and body weight (B) were measured

weekly. # p<0.05, ### p<0.0001, two-way repeated-measures ANOVA; * p<0.05, ** p<0.005,

*** p<0.0001, Bonferroni post-test when ANOVA was significant (black symbols, WT Rapa vs.

WT control; red symbols, L613V/+ Rapa vs. L613V/+ control). (C) Heart weight/body weight

(HW/BW) ratio was measured by the end of treatment. ns, not significant. *** p<0.0001,

Bonferroni post-test when ANOVA was significant. n= 11 (WT); 10 (WT Rapa); 13 (L613V/+);

12 (L613V/+ Rapa).

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5.2.3 Cell(s)-of-origin for Raf1 mutant-induced HCM

Global Raf1L613V expression causes eccentric cardiac hypertrophy with enhanced cardiac

contractility and cardiac fetal gene re-expression (464). Under basal conditions, fibrosis is

minimal, but TAC causes massive fibrosis in L613V/+ hearts and functional decompensation.

Recent Langendorf preparations (data not shown) reveal increased contractility in the L613V/+

heart, indicating a cardiac-intrinsic process, but the detailed physiological effects of mutant Raf1

at the single cell level are unclear. Furthermore, both cardiomyocytes (CM) and cardiac

fibroblasts (CF) from L613V/+ mice show increased Mek/Erk activation in response to multiple

agonists, including growth factors, cytokines, and GPCR agonists (464), so either cell type might

contribute to HCM pathogenesis in NS. Indeed, CF are found throughout the heart and account

for up to two-thirds of the cells in adult hearts (478). Moreover, recent studies indicate that CF

play key roles in myocardial development and function (457, 458). Embryonic CF induce

myocyte proliferation (457), whereas adult CF promote myocyte hypertrophy (457) and evoke

pathological hypertrophy and fibrosis in response to disease stimuli (451, 452). Notably,

enhanced Ras/Erk activation in CF is critical for pathological hypertrophy and fibrosis caused by

overexpression of the adrenergic receptor in CM (452). In a neonatal rat cell co-culture model

(479), Ang II stimulates cardiac myocyte hypertrophy via paracrine release of TGF-β1 and

endothelin-1 from CF. Studying the cell(s)-of-origin for RAF1 mutant-induced HCM would

advance our knowledge on the pathogenesis of HCM, and also provide important information for

therapeutic investigations (e.g., drug screening on the relevant cell type).

In order to ask whether CM and/or CF expression of mutant Raf1 is/are important for the

cardiac phenotype in L613V/+ mice, I have begun to induce Raf1L613V expression selectively in

CM or CF by crossing inducible L613Vfl/+ mice with tissue-specific Cre lines, and monitor

effects on basal and pressure overload-induced cardiac hypertrophy. For CM-specific expression,

L613Vfl/+ mice were first crossed to Mlc2v-Cre mice, in which Cre is expressed beginning at

~E9.5 (480, 481), to generate L613Vfl/+:Mlc2vCre/+ mice. The morphology of the

L613Vfl/+:Mlc2vCre/+ and control hearts was analyzed by echocardiography at 10 weeks of age,

at what time L613V/+ mice already have a significant cardiac hypertrophy and chamber

dilatation. However, L613Vfl/+:Mlc2vCre/+ mice did not show the same cardiac phenotypes as

L613V/+ mice. Instead, they exhibited a trend towards, but not a significant, increase in LVPWd

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Figure 5-5. Cardiomycyte-specific expression of Raf1L613V does not cause significant

cardiac hypertrophy.

LVPWd (A) and LV chamber dimension (B), as measured by echocardiography at 10 weeks of

age. ns, not significant. * p<0.05, Bonferroni post-test when ANOVA was significant. n= 9

(WT); 11 (L613Vfl/+); 9 (Mlc2vCre/+); 11 (L613Vfl/+:Mlc2vCre/+).

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at 10 weeks of age (Figure 5-5A). There was no chamber dilatation in these mice, although they

showed a significant decrease in LVIDs (Figure 5-5B), which was consistent with a mild

increase in cardiac function (Figure 5-6) and contractility (Figure 5-7). In total,

L613Vfl/+:Mlc2vCre/+ mice showed a mild concentric cardiac hypertrophy instead of the

eccentric cardiac hypertrophy observed in L613V/+ mice. However, we cannot conclude that

CMs are not the only cell-of-origin for the HCM phenotype because our preliminary data showed

that expression of Mlc2v-Cre was not universal in all ventricular CMs at E12.5, although studies

from other groups demonstrate that Mlc2v-Cre efficiently targets the ventricular myocardium

(481, 482). The big variation in the LVPWd measurement (Figure 5-5A) could reflect

differential Mlc2v-Cre expression and Raf1L613V induction within the ventricle. The cardiac

phenotype of these L613Vfl/+:Mlc2vCre/+ mice should be monitored at a later stage (e.g., 4

months of age) to see whether they would progress to more severe cardiac hypertrophy or even

chamber dilatation. Alternatively, another CM-specific Cre line (e.g., α-MHC-MerCreMer (483))

might be needed to determine the contribution of CM in Raf1 mutant-induced HCM.

To date, there is no Cre line for specifically targeting cardiac fibroblasts. However,

several Cre lines targeting the general fibroblast population have been used to study the function

of CF. Considering that studies have shown that CF are heterogeneous (45-49), different CF-

specific Cre lines, such as Cre expressed under the control of fibroblast-specific protein 1

promoter (Fsp1-Cre) (484-486) and the Col1a2 promoter (Col12a-Cre) (487, 488), should be

used to monitor the effects of CF-specific Raf1L613V expression.

There are several potential outcomes to these experiments. HCM might be mediated

entirely via CM or CF; alternatively, both might contribute. In the latter case, CM or CF might

be particularly important for different aspects of the HCM phenotype (e.g., effects on wall

thickness vs. contractility). Our preliminary data above for CM-specific expression of NS-

associated Raf1 mutant suggests that CM might not be the only cell-of-origin for RAF1 mutant-

induced HCM. We strongly suspect that CF will be important (if not required) for the severe

fibrosis post-TAC found in L613V/+ mice, but also expect effects on basal function. If CF-

specific expression of the Raf1 mutant does not cause detectable HCM, it would imply a

complex interaction between CM and CF in HCM pathogenesis. We would then evaluate mice

expressing mutant Raf1 in both cell types by generating compound CF/CM-Cre:L613Vfl/+

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Figure 5-6. Cardiomycyte-specific expression of Raf1L613V causes mild increase in cardiac

function.

Echocardiographic parameters at 10 weeks of age. ns, not significant. * p<0.05, ** p<0.005,

Bonferroni post-test when ANOVA was significant. n= 9 (WT); 11 (L613Vfl/+); 9

(Mlc2vCre/+); 11 (L613Vfl/+:Mlc2vCre/+).

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Figure 5-7. Cardiomycyte-specific expression of Raf1L613V causes mild increase in cardiac

contractility.

Cardiac contractility (A) and relaxation (B) of 4-month-old hearts as measured by invasive

hemodynamic analysis. ns, not significant. * p<0.05, Bonferroni post-test when ANOVA was

significant. n= 3 (WT); 3 (L613Vfl/+); 5 (Mlc2vCre/+); 6 (L613Vfl/+:Mlc2vCre/+).

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mice. In the unlikely event that even the latter does not result in HCM, we would explore the

possibility that other cell types contribute to the HCM phenotype, such as endothelial cells (Tie2-

Cre) (93). Surprisingly, recent preliminary data indicate that L613Vfl/+:Tie2-Cre/+ mice showed

a significant cardiac hypertrophy (data not shown), although whether it was primary or

secondary to valve defects needs to be further investigated.

5.2.4 The roles of specific Erk isoforms in Raf1-induced HCM

I found that post-natal MEK inhibition normalizes the growth, facial and cardiac defects

in L613V/+ mice, indicating that elevated MEK/ERK activity is critical for causing HCM and

other NS phenotypes. To further investigate the role of specific Erks in HCM pathogenesis, I

took a genetic approach by knocking-out Erk1 and/or Erk2 in L613V/+ mice and monitoring

their cardiac phenotypes.

In initial experiments, I analyzed L613V/+:Erk1+/- and L613V/+:Erk1-/- mice (Figure

5-8, Figure 5-9, and Figure 5-10). Remarkably, Erk1 deficiency normalized the increased

contractility in L613V/+ mice, but did not rescue the cardiac hypertrophy. On the other hand,

preliminary data showed that knocking-out one allele of Erk2 in L613V/+ mice could not rescue

the cardiac hypertrophy, and might even cause more severe hypertrophy (Figure 5-11). The

contractility of L613V/+:Erk2+/- hearts needs to be analyzed further.

There are several potential outcomes of these experiments. If reducing Erk2 levels can

rescue cardiac contractility as does Erk1 deficiency, the total Erk activity likely is critical for

inducing cardiac phenotypes in NS, and there may be a different threshold of the level of Erk

activation for regulating cardiac contractility vs. hypertrophy. If knocking-out one allele of Erk2

does not alter the contractility in L613V/+ mice, there is likely to be a specific role for Erk1 in

regulating cardiac contractility, considering that Erk2 levels are higher than Erk1 in both CM and

CF (464, 489). To further investigate the specific roles of Erk1/2 in cardiac contractility vs.

hypertrophy conclusively, complete deletion of Erk2 needs to be carried out. Due to the

embryonic lethality caused by full Erk2 depletion, cardiac-specific depletion of Erk2 by Nkx2.5-

Cre line (490), or knocking-out of Erk2 in the relevant cell type(s), which are the cell(s)-of-

origin for Raf1 mutant-induced HCM, needs to be done. These mouse models with specific

depletion of Erk1 or Erk2, or both, also will help us to investigate the downstream target(s) of

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Figure 5-8. Reducing Erk1 levels in L613V/+ mice does not rescue the cardiac hypertrophy

and chamber dilatation.

HW/BW ratio (A), LVPWd (B) and LV chamber dimension (C and D) at 4 month of age. ns, not

significant. * p<0.05, ** p<0.005, Bonferroni post-test when ANOVA was significant. n= 12

(WT); 15 (Erk1+/-); 11 (Erk1-/-); 12 (L613V/+); 15 (L613V/+:Erk1+/-); 8 (L613V/+:Erk1-/-).

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Figure 5-9. Reducing Erk1 levels in L613V/+ mice does not rescue the cardiac function.

Echocardiographic parameters at 4 month of age. ns, not significant. # p<0.05, 1-tailed Student’s

t test; ** p<0.005, *** p<0.0001, Bonferroni post-test when ANOVA was significant. n= 12

(WT); 15 (Erk1+/-); 11 (Erk1-/-); 12 (L613V/+); 15 (L613V/+:Erk1+/-); 8 (L613V/+:Erk1-/-).

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Figure 5-10. Reducing Erk1 levels in L613V/+ mice rescues the cardiac contractility.

Cardiac contractility (A) and relaxation (B) of 4-month-old hearts as measured by invasive

hemodynamic analysis. ns, not significant. # p<0.05, ## p<0.005, 1-tailed Student’s t test; *

p<0.05, ** p<0.005, *** p<0.0001, Bonferroni post-test when ANOVA was significant. n= 14

(WT); 18 (Erk1+/-); 8 (Erk1-/-); 14 (L613V/+); 13 (L613V/+:Erk1+/-); 6 (L613V/+:Erk1-/-).

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Figure 5-11. Reducing Erk2 levels in L613V/+ mice does not rescue the cardiac

hypertrophy.

HW/BW ratio at 4 month of age. * p<0.05, *** p<0.0001, Bonferroni post-test when ANOVA

was significant. n= 4 for each genotype.

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RAS/ERK signaling in the pathogenesis of NS and HCM, and in the regulation of cardiac

contractility vs. hypertrophy.

5.2.5 Genetic modifiers in D486N/D486N mice

The incompletely penetrant growth/facial dysmorphia/viability phenotype of

D486N/D486N mice on a 129Sv × C57BL/6 mixed background suggests the existence of a

modifier gene(s) that vary between the 129Sv and C57BL/6 strains. To screen genetic loci linked

to the s-D486N/D486N phenotypes, genomic DNA from s-D486N/D486N and n-D486N/D486N

mice was collected (Figure 5-12) and subjected to linkage analysis using the Illumina Mouse

Medium Density linkage panel. This panel contains 1449 SNP markers, among which 877 are

informative (different) between 129Sv and C57BL/6 mice. LOD (logarithm of odds) scores were

calculated by R/qtl (491) for mapping quantitative trait loci (QTLs) in experimental populations

derived from inbred lines (Figure 5-13). The trait was also binarized using a cutoff of body

weight at 10g (Figure 5-14 and Figure 5-15).

These preliminary mapping studies identified a 129Sv locus on mouse chromosome 8

that is strongly linked (LOD score ~15 for quantitative trait; LOD score ~9 for binary trait) to the

s-D486N/D486N phenotype (Figure 5-14, Figure 5-15 and Figure 5-16). However, this region

covers more than 30 megabase pairs (Mbps), and contains numerous genes. The Mouse SNP

Query from Mouse Genome Informatics (MGI) reveals that there are hundreds of SNPs, varying

between 129Sv and C57BL/6 strains, within this region, the majority of which are coding-

synonymous or located within introns or non-coding sequences. Most likely, the modifier(s) are

differentially regulated transcriptionally or translationally between 129Sv and C57BL/6 mice.

Further studies need to be performed in order to find the critical genetic modifier(s)

linked to the s-D486N/D486N phenotypes. More genomic DNA samples from s-D486N/D486N

and n-D486N/D486N mice could be collected and thereby increase the sample size of the

screening in order to narrow the region. Alternatively, this can be achieved by genotype

imputation without including more samples. The number of SNPs assessed in Illumina Mouse

Medium Density genotyping panels provides very low genomic coverage. Statistically, it is

therefore more likely for phenotype-associated SNPs to be in linkage disequilibrium (LD) with

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Figure 5-12. Distributions of the body weight data for all the 56 samples.

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0

5

10

15

Chromosome

LOD

Sco

re

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 1819 X

Figure 5-13. Genome-wide LOD score plot of the quantitative trait.

Produced by Pingzhao Hu

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0

2

4

6

8

Chromosome

LOD

Sco

re

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 1819 X

Figure 5-14. Genome-wide LOD score plot of the binary trait.

The trait is binarized using a cutoff of body weight: 10g. Produced by Pingzhao Hu

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Figure 5-15. LOD score plot of the binary trait on Chromosome 8.

The trait is binarized using a cutoff of body weight: 10g. Black dots indicate the individual SNPs

tested in the analysis.

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Figure 5-16. Boxplots of body weight for mice with different genotypes.

rs6237645 and rs3662808 are the two SNPs with highest LOD score from the array.

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causal variants than to be causal themselves (492). Therefore, a list of all SNPs that are in strong

LD with each SNP strongly linked to the s-D486N/D486N phenotype could be generated using

data from the laboratory mouse haplotype map (493). Through this imputation approach, we

could extend the coverage of phenotype-associated SNPs to include a more comprehensive list of

putative functional SNPs. On the other hand, gene expression profiles for the genes coded within

this region could be compared in available databases or be analyzed by qPCR in the relevant

tissues (e.g., pituitary, hypothalamus and liver for growth retardation) between the two mouse

strains. The gene(s) that is(are) strongly down-regulated on the 129/Sv background, could be the

critical genetic modifier(s) linked to the s-D486N/D486N phenotypes.

5.3 Concluding Remarks

In this thesis, I have carried out detailed studies leading to a better understanding of the

molecular pathogenesis of NS-associated RAF1 mutations. Using knock-in mouse models and a

variety of biochemical and molecular biology approaches, I have demonstrated that elevated

MEK/ERK activation is critical for causing HCM and other NS phenotypes. My work also

indicates that increased BRAF heterodimerization is the common theme in the pathogenesis of

NS-associated RAF1 mutations. The role of RAS/ERK signaling in regulating pathological

cardiac hypertrophy revealed in this study resolves a prior controversy in the field. I further

identified MEK inhibitors as potential therapeutic agents for the treatment of RAF1 mutant-

associated NS. In comparison with other RASopathy mouse models, I propose that different

RASopathies lead to distinct alterations in signaling, and a personalized, mutant gene-specific

strategy may be needed for treating the RASopathies. Further studies are required to clarify the

cell(s)-of-origin underlying HCM in NS, the role of specific Erk family members and modifier

gene(s) in Raf1 mutant NS.

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