RNA RECOGNITION BY THE PATTERN RECOGNITION RECEPTOR RIG-I:
ROLES OF RNA BINDING, MULTIMERIZATION, AND RNA-DEPENDENT
ATPASE ACTIVITY
By
ELIZABETH E. DELANEY
Submitted in partial fulfillment of the requirements for the degree of Doctor of
Philosophy
Dissertation Advisor: Dr. Eckhard Jankowsky
Department of Biochemistry
CASE WESTERN RESERVE UNIVERSITY
August 2014
CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve this thesis/dissertation of
Elizabeth E. DeLaney .
Candidate for the Doctor of Philososphy degree*
(signed) David Samols . (chair of the committee)
Eckhard Jankowsky .
Derek Abbott .
Blanton Tolbert .
Jonatha Gott .
(date) March 28, 2014 .
*We also certify that written approval has been obtained for any
proprietary material contained therein.
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Table of Contents
List of Tables…………………………………………………………………………….ix
List of Figures…………………………………………………………………………….x
Acknowledgements>……………………………………………………………………xiv
List of Abbreviations……………………………………………………………….…..xvi
Abstract…………………………………………………………………………………xix
Chapter 1: Pathogen detection by the innate immune system
1.1 Innate immunity: detection of conserved molecular patterns…………………...1
1.2 Types of PRRs……………………………………………………………………2
1.2.1 Toll-like receptors……………………………………………………….2
1.2.2 C-type lectin receptors………………………………………………...... 5
1.2.3 Nod-like receptors………………………………………………………7
1.2.4 RIG-I-like receptors……………………………………………………..9
1.2.4.1 Identification and function of RIGI……………………………...10
1.2.4.2 Identification and function of MDA5 and LGP2………………...11
1.3 RIG-I signaling pathway…………...…………………………………………...13
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1.4 RIG-I and MDA5 activate antiviral signaling in response to distinct viruses…17
1.4.1 RIG-I recognizes a diverse range of viruses………………………….17
1.4.2 MDA5 is primarily activated by picornaviruses……………………...20
1.5 RIG-I domain structure and function…………………………………………..20
1.5.1 Overall RIG-I domain structure………………………………………20
1.5.2 CARD domains in RIG-I are necessary for signal transduction………23
1.5.2.1 Structure and function of RIG-I CARD domains………………..23
1.5.2.2 Ubiquitination of the CARD domains regulates RIG-I signal
transduction………………………………………………………26
1.5.3 RIG-I helicase/ATPase domain………………………………………..30
1.5.4 RIG-I CTD functions in ligand binding and multimerization…………37
1.5.5 Molecular mechanism of RIG-I substrate recognition…………….…..41
1.6 RIG-I recognizes multiple structural features of viral RNA……………………45
Chapter 2: Development of L21 ribozyme as a method to generate RIG-I RNA
substrates……………………………………………………………………51
2.1 Rationale for using the L-21 ribozyme for producing RIG-I substrates in
vitro...... 51
2.2 Characterization of L-21 ribozyme activity on RIG-I substrates……….……...54
2.3 Discussion……………………………………………………………………..60
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Chapter 3: RIG-I tightly binds dsRNA………………………………………………....61
3.1 Introduction……………………………………………………………………61
3.2 Results…………………………………………………………………………62
3.2.1 Purification of wild-type and mutant RIG-I………………………………62
3.2.2 RIG-I binds ssRNA with low affinity despite presence of a
5’-triphosphate……………………………………………………………63
3.2.3 5’-triphosphate does not impact RIG-I binding to dsRNA……………….64
3.2.4 RIG-I binds dsRNA tightly regardless of duplex length………………….68
3.2.5 RIG-I binding to RNA duplexes at least 16 bp involves two species or
RNA-protein complexes………………………………………..…………70
3.2.6 RIG-I multimerization is dependent upon RNA duplex length…….…....72
3.2.7 RIG-I deletion mutant demonstrate major RNA binding site is the CTD..77
3.3 Discussion……………………………………………………………………...84
3.3.1 RNA 5’-end structure has no significant effect on RIG-I RNA binding….84
3.3.2 RIG-I RIG-I binds dsRNA cooperatively and multimerization is dependent
on duplex length…………………………………………………………..87
3.3.3 Major RNA binding site is in the RIG-I CTD…………………………….88
Chapter 4: RIG-I ATPase activity recognizes the presence of dsRNA……..………….90
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4.1 Introduction………………………………………………………..…………90
4.2 Results…………………………………………..……………………………91
4.2.1 Nucleotide has no effect on RIG-I-RNA affinity, but promotes complex
dissociation………………………………………………….…………………91
4.2.2 ATP enhances dissociation of RIG-I from RNA………………………..96
4.2.3 RIG-I RNA-dependent ATPase activity is independent of RNA duplex
length…………………………………………………………………...100
4.3 Discussion…………………………………………………………………….102
Chapter 5: RIG-I proofreads RNA duplexes for blunt ends with its ATPase activity…106
5.1 Introduction…………………………………………………………………...106
5.2 Results…………………………………………………………………………107
5.2.1 RIG-I efficiently binds RNA duplexes with blocked ends………………107
5.2.2 RIG-I ATPase activity proofreads RNA duplexes for blunt ends……….109
5.3 Discussion…………………………………………………………………….112
5.3.1 RIG-I RNA-dependent ATPase activity identifies blunt duplex ends…...112
5.3.2 Model of RIG-I substrate recognition……………………………………113
Chapter 6: Future directions……………………………………………………………116
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6.1
Introduction……………………………………………………………………...116
6.2 Effects of RNA 5’-end structure on RIG-I RNA recognition in vivo……….117
6.3 Activation of antiviral signaling in vivo by frayed end substrates…………..118
6.4 Effect of ubiquitination of RIG-I on RIG-I-RNA binding and ATP
hydrolysis…………………...... 119
Chapter 7: Materials and Methods…………………………………………………….124
7.1 Materials………………………………………………………………………124
7.1.1 Plasmids…………………………………………………………………124
7.1.2 Proteins………………………………………………………………….126
7.1.3 Oligonucleotides………………………………………………………...127
7.1.4 Miscellaneous reagents………………………………………………….130
7.2 Methods………………………………………………….……………………131
7.2.1 Radiolabeling and gel purification of RNA and DNA oligonucleotides..131
7.2.2 Characterization of L-21 ribozyme activity on RIG-I substrates………..132
7.2.3 End-labeling of ssRNA with L-21 ribozyme…………………………….132
7.2.4 RIG-I binding under equilibrium conditions…………………………….133
7.2.5 Derivation of equations for coupled equilibrium………………………..134
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7.2.6 Dissociation kinetics of RIG-I-RNA complexes………………………...136
7.2.7 Glutaraldehyde crosslinking of RIG-I-RNA complexes…………………137
7.2.8 Measurement of RIG-I ATPase activity with thin layer chromatography.138
Bibliography……………………………………………………………………………139
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List of Tables
Table 1.1. Viruses recognized by RIG-I………………………………………………...19
Table 2.1. L-21 ribozyme RNA precursor substrate sequences…………………………55
Table 3.1. Wild-type RIG-I binding parameters calculated from coupled equilibrium…71
Table 3.2. Stimulation of RIG-I ATPase activity by 36 nt ssRNAs and 36 bp duplexes
with and without a 5’-triphosphate……………………………………...…..79
Table 3.3. RIG-IΔCARD binding parameters calculated from coupled equilibrium...…82
Table 4.1. RIG-I binding to 36 bp duplex with a 5’-triphosphate in the presence of
nucleotide…………………………………………………………………...92
Table 4.2. Off rates of RIG-I from indicated RNA duplexes in presence and absence of
ATP…………………………………………………………………………..99
Table 5.1. Dissociation rate constants and kinetic parameters of RIG-I in presence of
end-blocked RNA substrates………………………………………………111
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List of Figures
Figure 1.1. Schematic of the major classes of cellular PRRs……………………………4
Figure 1.2. Schematic of RIG-I domain structure……………………………………….10
Figure 1.3. Schematic of MDA5 and LGP2 domain structures…………………………13
Figure 1.4. Schematic of RLR signaling pathway………………………………………15
Figure 1.5. Sequence alignment of RIG-I, MDA5, and LGP2………………………….21
Figure 1.6. Schematic showing the Greek key motif present in CARD domains………24
Figure 1.7 Structure of MAVS CARD domain…………………………………………24
Figure 1.8 Crystal structure of duck RIG-I in the absence of RNA and NTP………….25
Figure 1.9. TRIM25 interacts with T55 in CARD1 to ubiquitinate K172 in CARD2…27
Figure 1.10. Schematic of RIG-I domain structure with residues ubiquitinated by
Riplet………………………………………………………………………28
Figure 1.11. Schematic of RIG-I domain structure with residue important for interaction
with TRIM25 shown and the interaction of the second CARD domain with
K63-linked polyubiquitin chains…………………….……………………29
Figure 1.12. Schematics of conserved RNA helicase domain motifs…………………..32
Figure 1.13. Crystal structure of ligand bound human RIG-I………………………...... 37
Figure 1.14. Schematic and crystal structure of RIG-I CTD……………………………40
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Figure 1.15 RIG-I undergoes a conformational change upon RNA binding…...……….43
Figure 2.1. Tetrahymena ribozyme crystal structure and schematic of mechanism……54
Figure 2.2. L-21 ribozyme efficiently cleaves R13L21PBot precursor RNA………...... 57
Figure 2.3. L-21 ribozyme efficiently cleaves R13L21P and R10L21P precursor
RNAs………………………………………………………………………..58
Figure 2.4. L-21 ribozyme efficiently cleaves and 84 nt precursor in the presence of GTP
And analogs……………………………………….………………………59
Figure 3.1. Schematic of wild-type RIG-I and RIG-I deletion mutants……………….62
Figure 3.2. Purified WT and mutant RIG-I proteins……………………………….…..63
Figure 3.3. RIG-I binds ssRNA weakly, and 5’-triphosphate enhances binding
affinity…………………………………………………………………….64
Figure 3.4. RIG-I binds a 36 bp dsRNA tightly, regardless of presence of a
5’-triphosphate……………………………………………………………..65
Figure 3.5. RIG-I binds a 10 bp dsRNA tightly, regardless of presence of a
5’-triphosphate...... 67
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Figure 3.6. RIG-I binds a 13 bp dsRNA tightly, regardless of presence of a
5’-triphosphate...... 68
Figure 3.7. RIG-I binds dsRNA tightly regardless of duplex length…………………....70
Figure 3.8. RIG-I forms multiple RNA-protein complexes with a thirty-six base pair
RNA duplex………………………………………………………………...72
Figure 3.9. Schematic of stoichiometric binding reactions……………………………...73
Figure 3.10. RIG-I binds a thirteen base pair duplex as a dimer………………………..74
Figure 3.11. RIG-I binds a nineteen base pair duplex as a dimer, and a thirty-six base
pair duplex as a trimer……………………………………………………76
Figure 3.12. RIG-I binds a thirty-six base pair duplex with a 5’-triphosphate as a
dimer……………………………………………………………………….77
Figure 3.13. RIG-IΔCARD protein binds RNA very similarly to wild-type RIG-I…….81
Figure 3.14. Multiple RNA-protein complexes form when RIG-IΔCARD binds a 36 bp
RNA duplex……………………………………………………………….82
Figure 3.15. RIG-I deletion mutants lacking the RD are deficient in RNA binding……83
Figure 4.1. Schematic of glutaraldehyde crosslinking rections…………………………93
Figure 4.2. RIG-I-RNA complexes dissociate in the presence of ATP…………...…….94
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Figure 4.3. Schematic of reactions for measuring RIG-I off rate……………………….97
Figure 4.4. ATP enhances dissociation of RIG-I from RNA duplexes…………………98
Figure 4.5. Schematic of reactions for measuring RIG-I RNA-dependent ATPase activity
under conditions of saturating RNA……………………………………….101
Figure 4.6. RIG-I RNA-dependent ATPase activity is independent of RNA duplex length
and presence of 5’-triphosphate…………………………...………………102
Figure 5.1. Schematics of end-blocked RNA substrates……………………………….106
Figure 5.2. Single ssRNA overhangs on a nineteen bp RNA duplex does not affect RIG-I
RNA binding………………………………………………………………107
Figure 5.3. RIG-I tightly binds an RNA duplex with one end blocked………………..108
Figure 5.4. RIG-I tightly binds an RNA duplex with both ends blocked……………...109
Figure 5.5. An exposed blunt end is necessary for ATPase activity…………………...110
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Acknowledgements
I would like to thank my advisor, Dr. Eckhard Jankowsky, for the opportunity to
perform this work and for the training I received. I would like to thank my committee
members, Dr. David Samols, Dr. Derek Abbott, and Dr. Jonatha Gott for their scientific
and technical insight while I performed this work. I would like to thank Dr. Blanton
Tolbert for participating in my thesis defense committee. Additionally, I would like to
thank Dr. Derek Abbott for providing guidance regarding the expression of RIG-I in
insect cells.
I would like to thank Dr. Mark Caprara for serving on my committee prior to his
departure from the university. He initially suggested using the L-21 ribozyme to produce
5’-triphosphate 5’-end labeled RNA, and provided the construct encoding the L-21
ribozyme. Dr. Dominique Garcin provided the constructs necessary to express wild-type
and mutant RIG-I in E. coli. In addition, I thank Dr. Erik Andrulis for the use of his
tissue culture hood and incubator while I was propagating insect cells. I also would like
to thank Dr. William Merrick and Diane Baus for providing technical support.
I also am grateful for the training I received in Dr. Cheng-Ming Chiang’s lab prior
to my work with Dr. Jankowsky. My former colleagues, Dr. Shwu-Yuan Wu, Dr. Wei-
Ming Wang, and Dr. A-Young Lee provided valuable scientific and technical support.
I am very lucky to have worked with a fantastic group of colleagues in the
Jankowsky lab, both past and present, whose support has been a crucial factor in the
completion of my studies. Dr. Heath Bowers trained me in the Jankowsky lab method of
protein purification and Dr. Nicholas Kaye trained me to perform the gel shift assays that
xiv
have proven so important in my work. Dr. Fei Liu provided the equations for the coupled
equilibrium. Andrea Putnam taught me enzyme kinetics, and both she and Dr. Ulf-Peter
Guenther were always willing to help when needed, whether it was simply removing a
gel from the gel dryer for me so I could catch my bus or for discussing both theoretical
and technical issues that arose. The other current lab members, Frank Tedeschi, Armend
Axhemi, Sukanya Srinivasan, Xuan Ye, and Zhaofeng Gao were also always willing to
provide scientific or technical assistance.
xv
List of Abbreviations
5’-UTR, 5’-untranslated region
ADP, adenosine diphosphate
ADPNP, adenosine imidodiphosphate
ATP, adenosine triphosphate
BIR, Baculovirus inhibitor of apoptosis repeat
CARD, Caspase activation and recruitment domain
CARDIF, CARD adaptor inducing interferon-β
CLR, C-type lectin receptor
CRD, Carbohydrate recognition domain
CTD, C-terminal domain
CYLD, Cylindramatosis
DD, Death domain
DED, Death effector domain
ERIS, Endoplasmic reticulum interferon stimulator
FADD, Fas-associated death domain
GDP, Guanosine diphosphate
GMP-PNP, Guanylyl imidodiphosphate
GTP, Guanosine triphosphate
HCV, Hepatitis C virus
IκB, Inhibitor of κB
IKK, Inhibitor of NF-κB kinase
IKKε, Inhibitor of NF-κB kinase subunit ε
IKKγ, Inhibitor of NF-κB kinase subunit γ
IPS-1, Interferon-β promoter stimulator
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IRF-3, Interferon regulatory factor 3
LGP2, Laboratory of genetics and physiology
LRR, Leucine-rich repeat
MAPK, mitogen activated protein kinase
MAVS, Mitochondrial antiviral signaling protein
MDA5, Melanoma differentiation-associated gene 5
MITA, Mediator of IRF-3 activation
MPYS, Membrane tetraspanner associated with MHC class II
NBS, Nucleotide binding site
NEMO, NF-κB essential modulator
NF-κB, Nuclear factor κB
NLR, Nod-like receptor
NLRX1, NLR family member X1
NS1, Nonstructural protein 1
PAMP, Pathogen associated molecular pattern
PKC, Protein kinase C polyI:C, polyinosinic polycytodylic acid
PRR, Pattern recognition receptor
PYD, Pyrin domain
RD, Regulatory domain
REUL, RIG-I E3 ubiquitin ligase
RIG-I, Retinoic acid inducible gene one
RIP1, receptor interacting protein 1
RLR, RIG-I-like receptor
RNF125, Ring finger protein 125
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RNF135, Ring finger protein 135
siRNA, small interfering RNA
SF1, Superfamily 1
SF2, Superfamily 2
STING, Stimulator of interferon genes
TBK1, TANK binding kinase 1
TIR, Toll/IL-1R homology
TLR, Toll-like receptor
TRADD, TNF receptor type 1-associated DEATH domain protein
TRAFs, Tumor necrosis factors
TRIM25, Tripartite motif 25
VISA, Virus-induced signaling adaptor
VSV, Vesicular stomatitis virus
xviii
RNA Recognition by the Pattern Recognition Receptor RIG-I: Roles of RNA Binding,
Multimerization, and RNA-dependent ATPase Activity
Abstract
By
ELIZABETH E. DELANEY
Recognition of viral RNA by mammalian cells is critical for the activation of the
innate immune system. Viral RNA is recognized by several pathogen recognition
receptors, including retinoic acid inducible gene I, or RIG-I. RIG-I consists of two N-
terminal tandem caspase activation and recruitment domains, a central helicase/ATPase
domain, and a C-terminal regulatory domain. Following RNA binding, RIG-I undergoes
a conformational change, ubiquitination, and dimerization, all of which are necessary for
interaction with the adaptor protein mitochondrial antiviral signaling (MAVS). Binding
to MAVS triggers signaling cascades that induce the transcription of antiviral peptides.
RIG-I has been shown to be activated by both dsRNA and dsRNA containing 5’-
triphosphates in vivo, and its ATPase activity is critical for activation. A significant body
of work has been published regarding the cellular role of RIG-I, but how RIG-I
distinguishes viral RNAs from cellular RNAs remains unclear. To understand how RIG-I
distinguishes between different substrates, we performed a biochemical analysis of RIG-I
RNA binding, ATPase activity, and oligomerization. We used purified RIG-I to
quantitatively analyze how RIG-I interacts with various model RNAs. We show that
RIG-I binds tightly to dsRNA regardless of the presence of a 5’- triphosphate.
xix
Dissociation of RIG-I from RNA is enhanced by ATP. RIG-I ATPase activity is
stimulated by RNA duplexes as short as 10 bp, and a RIG-I monomer is sufficient for
ATPase activity. RIG-I binds to RNA duplexes with and without blunt ends, however
ATPase activity is only activated by RNA duplexes containing at least one blunt end.
Collectively, these data suggest that duplex structure and nucleotide binding play a
critical role in RIG-I binding and activation. Our data suggest a model in which
distinguishing self from non-self RNA requires the recognition of multiple features in a
single RNA by RIG-I.
xx
Chapter 1
Introduction
Pathogen Detection by the Innate Immune System
1.1 Innate Immunity: Detection of conserved molecular patterns
Detection of invading pathogens requires an organism to distinguish between its own (self) RNA and DNA and foreign (non-self) nucleic acid (Akira et al, 2006).
Vertebrates have two different forms of immunity to protect against invading microorganisms, innate immunity and adaptive immunity (Dempsey et al, 2003). The innate immune system is the first line of defense and can mount a response to a pathogen within hours, slowing down the infection to provide time for the adaptive immune response to develop (Akira et al, 2006). Innate immunity is primarily mediated by macrophages, dendritic cells, and neutrophils (Dempsey et al, 2003, Akira et al, 2006, and Kumar et al, 2011). Stimulation of the innate immune systems also activates adaptive immunity (Dempsey et al, 2003). Antigen presenting cells (APCs), such as macrophages and dendritic cells, internalize and degrade extracellular pathogens or virus- infected cells. They then present pathogen-associated proteins to T-cells. Adaptive immunity results in the generation of antigen-specific receptors functions to both clear the initial infection with a particular pathogen and to prevent future infection with the same pathogen (Dempsey et al, 2003 and Kumar et al, 2011). Innate immune responses are not restricted to vertebrates and are the only immune responses present in both invertebrates and plants (Akira et al, 2006). Adaptive immunity arose less than 500
1 million years ago and is only present in vertebrates (Dempsey et al, 2003 and Akira et al,
2006).
Adaptive immunity responds specifically to pathogens by generating antigen- specific receptors from somatic recombination of germline-encoded immunoglobulin gene segments (Gonzalez et al, 2007 and Iwasaki and Medzhitov, 2010). The innate immune system detects pathogens through a more general mechanism (Figure 1.1).
Innate immunity depends on several classes of proteins known as pattern recognition receptors (PRRs) that detect the presence of pathogens in the cell. PRRs are germline- encoded and are constitutively expressed in many tissue types (Akira et al, 2006 and
Iwasaki and Medzhitov, 2010). PRRs are able to identify pathogens via the recognition of pathogen associated molecular patterns (PAMPs). PAMPs are conserved molecular features of a pathogen that are frequently necessary for its survival or reproduction (Akira et al, 2006). Examples of PAMPs are bacterial cell wall components such as lipopolysaccharide, the primary protein component of bacterial flagellum (flagellin), and double-stranded RNA (dsRNA) viral replication intermediates. Upon activation of the innate immune system, infected cells produce proinflammatory cytokines and type-I interferons (Figure 1.1) (Akira et al, 2006 and Kumar et al, 2011).
1.2 Types of PRRs
1.2.1 Toll-like Receptors
The innate immune system employs several major classes of PRRs. The Toll-like receptors (TLRs) are evolutionarily conserved from the nematode Caenorhabditis
2 elegans to mammals (Figure 1.1) (Akira et al, 2006). The founding member Toll was initially identified in Drosophila melanogaster as a protein necessary for the development of embryonic dorsoventral polarity (Hashimoto et al, 1988), and later shown to be critical for the fly antifungal response (Lemaitre et al, 1996). Mammals express twelve different TLR proteins, and subgroups of the TLR family recognize distinct types of PAMPs (Akira et al, 2006 and Kumar et al, 2011). TLR signaling is activated in response to bacterial, viral, fungal, and protozoan infection of cells (Akira et al, 2006).
TLR1, TLR2, and TLR6 recognize lipids present in bacterial and fungal cell walls (e.g. diacyl and triacyl lipopeptides and phospholipomannan) (Alexopoulou et al, 2002,
Ozinsky et al, 2000, Takeuchi et al, 2001, Thoma-Uszynski et al, 2001, Takeuchi et al,
2002, Gilleron et al, 2003, Krutzik et al, 2003, and Travassos et al, 2004,). TLR3, TLR7,
TLR8, and TLR9 recognize viral double-stranded and single-stranded RNA, and DNA
(Hemmi et al, 2000, Krieg, 2002, Hochrein et al, 2004, Krug et al 2004a, Krug et al,
2004b, Lund et al, 2003, Tabeta et al, 2004, Hemmi et al, 2002, Heil et al, 2004, Diebold et al, 2004). The remaining TLRs recognize various proteins present in viral envelopes
(e.g. haemagglutinin), many proteins and polysaccharides present in bacterial and fungal structures or cell walls (e.g. flagellin, lipopolysaccharide, and mannan), and also host proteins produced during the inflammatory response (e.g. heat shock proteins and fibrinogen) (Poltorak et al, 1998, Shimazu et al, 1999, Hayashi et al, 2001, Netea et al,
2004). TLRs are expressed in various immune cells, including macrophages and dendritic cells, and also in some non-immune cells such as epithelial cells and fibroblasts
(Akira et al, 2006).
3
4 All TLRs are type I integral membrane glycoproteins and are present either on the
cell surface or in internal endsomal compartments (Figure 1.1) (Palsson-McDermott and
O’Neill, 2007). TLR extracellular domains consist of varying numbers of leucine-rich
repeat motifs (LRRs) that are domains are responsible for ligand sensing (Bell et al,
2005). Their intracellular domains are a cytoplasmic signaling domain known as the
Toll/IL-1R homology (TIR) domain. The TIR domain is responsible for homotypic
protein interactions with downstream adaptors (Bowie and O’Neill, 2000). Following stimulation of a TLR by a PAMP, an adaptor protein containing a TIR domain is recruited to the receptor. Signaling cascades are initiated that result in the activation of nuclear factor κB (NF-κB), mitogen activated protein kinase (MAPK), and interferon regulatory factor 3 (IRF-3) pathways (Figure 1.1)(Akira et al, 2006). The attack of the invading pathogen is then countered by the cell producing proinflammatory cytokines and type I interferons (Akira et al, 2006 and Palsson-McDermott and O’Neill, 2007).
1.2.2 C-type Lectin Receptors
The C-type lectin receptors (CLRs) are a diverse group of proteins with both immune and non-immune functions. CLRs were initially found to recognize endogenous ligands involved in cell adhesion, but recently several CLRs have been found to function as PRRs (Geitjenbeek et al, 2004). CLRs are either soluble proteins secreted into the extracellular matrix, or are transmembrane proteins that contain an extracellular carbohydrate recognition domain (CRD) and a variety of intracellular signaling domains
(Figure 1.1) (Palsson-McDermott, 2007). CLRs are classified as either classical or
5 nonclassical based on their CRD domains. The CRD domain of classical CLRs contains
conserved residues that form calcium binding sites and carbohydrate binding motifs.
Classical CLRs bind either mannose- or galactose-type carbohydrates. The CRD of
nonclassical CLRs does not contain these conserved residues, and these proteins are capable of binding a diverse repertoire of ligands (Geitjenbeek et al, 2004 and Graham
and Brown, 2009). For example, the CLRs Dectin-1 and Dectin-2 recognize α-mannans and β-glucans respectively, which are carbohydrates found in the cells walls of fungi such as Saccharomyces cerevisiae and Candida albicans (Brown et al, 2003 and
Drummond et al, 2011). Mincle, another CLR involved in pathogen recognition, was shown to recognize fungal α-mannose and the mycobacterial glycolipid, trehalose-6’6’- dimycolate (Yamasaki et al, 2009 and Ishikawa et al, 2009).
The CLRs identified as PRRs are primarily encoded in the natural killer gene complex (NKC), located on human chromosome 12, and consist of two groups: the
Dectin-1 and Dectin-2 clusters. These CLRs are expressed on myeloid cells (neutrophils, macrophages, and dendritic cells), and recognition of pathogens by CLRs results in the autophagy, degradation, and antigen presentation of the pathogen (Geitjenbeek et al,
2004). CLRs from the Dectin-1 group all have an extracellular CRD, a transmembrane domain, and one of two intracellular signaling domains: either immunoreceptor tyrosine- based activation motif (ITAMs) or immunoreceptor tyrosine-based inhibition motif
(ITIMs) (Figure 1.1) (Palsson-McDermott and O’Neill, 2007). Interaction of signaling molecules with ITAMs and ITIMs either activate or inhibit transcription factors, respectively (Palsson-McDermott and O’Neill, 2007, Geitjenbeek et al, 2004 and Graham and Brown, 2009). CLRs from the Dectin-2 group have a similar domain structure as the
6 Dectin-1 group, but their intracellular domains are very short and lack signaling
capability. To mediate downstream signaling following ligand recognition, these CLRs
pair with an additional receptor containing an ITAM. CLRs have been shown to induce
proinflammatory cytokines through the activation of NF-κB, and some have putative
roles in autoimmune disorders (Graham and Brown, 2009).
1.2.3 Nod-like Receptors
The Nod-like receptors (NLRs) are multi-domain cytoplasmic proteins implicated
in the recognition of bacterial, viral, and parasitic PAMPs. Thus far more than twenty
human NLRs have been identified (Palsson-McDermott and O’Neill, 2007). The
defining feature of NLRs is their nucleotide binding site (NBS) domain, which is a
member of the AAA+ superfamily of nucleotide binding and oligmerization domains.
This domain mediates ATP-dependent multimerization, which is necessary for the
activation of signaling following PAMP detection (Palsson-McDermott and O’Neill,
2007). The founding members of the NLR family, Nod1 and Nod2, recognize distinct
motifs in the bacterial cell wall component peptidoglycan (Chamaillard et al, 2003 and
Girardin et al, 2003).
Several subclasses of the NLRs form inflammasomes upon PAMP recognition.
Inflammasomes are multiprotein complexes formed by self-oligomerizing scaffold
proteins that are responsible for the activation of inflammatory caspases (Martinon et al,
2002). The NLRP1 inflammasome is activated in response to Bacillus anthrasis lethal toxin (Boyden and Dietrich, 2006), while the NLRP3 inflammasome has been shown to
7 be activated by pathogens including S. cerevisiae, C. albicans (Gross et al, 2009), toxins produced by Listeria monocytogenes and Staphylococcus aureus (alpha-toxin)
(Mariathasan et al, 2006 and Kanneganti et al, 2006), and both DNA (adenovirus) and
RNA viruses (Sendai virus and influenza virus) (Muruve et al, 2008). Additionally, mutations in NLR proteins have been shown to play significant roles in the development of various autoimmune diseases, with extensive work done on Nod2 and Crohn’s disease
(Ogura et al, 2001, Hugot et al, 2001, and Miceli-Richard et al, 2001).
Like TLRs, NLRs sense the presence of pathogens with varying numbers of LRRs
((Palsson-McDermott and O’Neill, 2007). The NLR family is divided into subfamilies depending on their N-terminal domain (Bryant and Fitzgerald, 2009). NLRs contain one of three domains involved in either the stimulation or inhibition of apoptosis: a caspase activation and recruitment domain (CARD), a pyrin domain (PYD), or a baculovirus inhibitor of apoptosis repeat domain (BIR). These domains are responsible for homotypic protein interactions with downstream effector proteins. All of the domains of
NLRs play specific roles in activating signaling in cells (Bryant and Fitzgerald 2009 and
Palsson and McDermott, 2007). Upon sensing a PAMP via its LRR domains, an NLR then undergoes ATP-dependent multimerization and a conformational change to expose the N-terminal domain. This CARD, PYD, or BIR domain then interacts with downstream effector proteins, initiating inflammatory signaling pathways such as the NF-
κB and MAPK pathways (Figure 1.1) (Bryant and Fitzgerald, 2009).
8 1.2.4 RIG-I-like Receptors
The RIG-I-like receptor (RLR) family members contain a conserved tripartite
domain structure and are localized in the cytoplasm. They initiate proinflammatory signaling in response to viral infections (Akira et al, 2006). RLRs contain a C-terminal
ligand sensing domain, a central helicase/ATPase domain, and tandem N-terminal protein
interaction domains responsible for interaction with downstream adaptor proteins (Akira
et al, 2006, Kang et al, 2002, and Yoneyama et al, 2004). Unlike NLRs, which contain
several different N-terminal domains responsible for signal transduction, the protein
interaction domain of RLRs is the CARD domain (Yoneyama et al, 2004). RLRs are
broadly expressed in both immune and non-immune cells. In uninfected cells, RLR
protein levels are very low. Protein levels increase dramatically in the presence of viral
infection and interferon production (Yoneyama et al, 2004).
The RLR family consists of only three proteins: RIG-I (retinoic acid inducible
gene I, annotated as DDX58), MDA5 (melanoma differentiation associated gene 5,
IFIH1), and LGP2 (laboratory of genetics and physiology 2, DHX58). They are
superfamily 2 (SF2) RNA helicases (Yoneyama et al, 2004, Kang et al, 2002, and Cui et al, 2001). Like NLRs, RLR binding to a PAMP stimulates an ATP-dependent conformational change and multimerization that exposes the signal transducing CARD
domains (Saito et al, 2007 and Cui et al, 2008). RLR signaling is potentiated by
interaction with a downstream adaptor protein that initiates the NF-κB and IRF-3
signaling pathways (Figure 1.1) (Seth et al, 2005, Xu et al, 2005, Meylan et al, 2005, and
Kawai et al, 2005).
9 1.2.4.1 Identification and Function of RIG-I
RIG-I was initially isolated as a retinoic acid-inducible gene involved in the differentiation of acute promyelocitic leukemia cells (1997, GenBank: AF038963). In
2000, the connection between RIG-I and viral infection was suggested by its porcine homolog, RHIV-1. RHIV-1 was discovered to be a helicase that was induced upon viral infection (Zhang et al, 2000). Yoneyama et al (2004) isolated Hhman RIG-I from a cDNA library screen for genes that can activate an IRF reporter gene in response to transfection with the double-stranded RNA mimic, polyinosinic-polycytidylic acid
(polyI:C), which is a strong inducer of interferon-β in mammalian cells. The N-terminal
CARD domains of RIG-I were identified as a positive regulator of type I interferons, since a RIG-I construct lacking the helicase or CTD domains was constitutively active for signaling in cells (Yoneyama et al, 2004). Furthermore, ATPase activity was shown to be necessary for the induction of interferon-β. A mutation to the nucleotide binding site in RIG-I’s helicase/ATPase domain significantly diminished RIG-I’s ability to induce interferon-β (Yoneyama et al, 2004).
RIG-I knockout mice provided additional evidence for RIG-I’s critical role in innate immunity. Disruption of RIG-I in mice resulted primarily in embryonic lethality, with only a few RIG-I-/- mice surviving for several weeks. However, mouse embryonic fibroblasts (MEFs) derived from these mice showed a significant defect in interferon-β
10 production after being transfected with in vitro transcribed dsRNA (Kato et al, 2006).
Utilizing a different approach, Wang et al (2007) was able to generate viable and fertile
RIG-I-/- mice. MEFs derived from these mice also demonstrated a compromised
immune response, similar to Kato et al (2006) (Wang et al, 2007). The identification and initial characterization of RIG-I demonstrate its role as a PRR responsible for the detection of viruses in the mammalian innate immune system.
1.2.4.2 Identification and Function of MDA5 and LGP2
MDA5 was identified in 2002 in a screen to identify genes involved in the interferon-β‐ and a protein kinase C (PKC) agonist-mediated terminal differentiation of
melanoma cells by Kang et al (2002). Sequence analysis of MDA5 revealed that it
contained helicase/ATPase and CARD domains, and its expression was found to be
upregulated by treatment of a melanoma cell line or human skin fibroblasts with
interferons or tumor necrosis factor-α (TNFα). Finally, MDA5 was shown to have
ATPase activity in the presence of polyI:C (Kang et al, 2002). MDA5 was also
independently identified in 2004 as a binding target of the inhibitory V proteins of
paramyxoviruses (Andrejeva et al, 2004). These lines of evidence suggested a possible
role for MDA5 in the cellular response to virus infections.
Following the identification of RIG-I as a PRR, a database search revealed the
sequence homology between RIG-I, MDA5, and LGP2, resulting in the annotation of the
RLR family of SF2 RNA helicases. MDA5 exhibits 23% sequence similarity to RIG-I’s
CARD domains and 35% sequence similarity to its helicase domain (Figure 1.4 and
Yoneyama et al, 2005). Yoneyama et al (2005) demonstrated that MDA5 participates in
11 the induction of the interferon response in the presence of polyI:C. Additionally, MDA5 knockout mice became extremely susceptible to picornavirus infection (Kato et al, 2006).
These lines of evidence suggested that like RIG-I, MDA5 functions as a PRR in mammalian cells.
Cui et al (2001) initially identified LGP2 in a screen to identify genes involved in normal and neoplastic mammary tissue development. Sequence analysis demonstrated that LGP2 contained a helicase/ATPase domain, though the protein’s function at the time was unknown (Cui et al, 2001). LGP2 lacks the CARD domains shared by its other two family members, but does share 31% sequence similarity with RIG-I and 41% sequence similarity with MDA5 (Figure 1.4 and Yoneyama et al, 2005). The role of LGP2 in the regulation of antiviral signaling is less clear. Lacking the signal transducing CARD domains, LGP2 is unable to activate antiviral signaling pathways. Like RIG-I and
MDA5, its expression is upregulated by interferons and it has demonstrated the ability to bind dsRNA (Komuro and Horvath, 2006). However, several different groups have demonstrated that LGP2 can act to either inhibit or enhance RLR signaling in cell culture
(Rothenfusser et al 2005, Saito et al, 2007, Venkataraman et al, 2007, and Murali et al,
2008). LGP2’s disparate domain structure suggests a different role than that of RIG-I or
MDA5, but its role in the modulation of RLR signaling remains unclear (Komuro and
Horvath, 2006).
12
1.3 RIG-I Signaling Pathway
In a resting cell, RIG-I and MDA5 are thought to exist in an auto-inhibited conformation (Luo et al, 2011 and Kowalinski et al, 2011). When an RNA virus invades a cell, RLRs recognize the genomic RNA and replication intermediates of the virus
(Akira et al, 2006). RLRs bind the viral RNA, triggering a conformational change in the protein that allows interaction with their downstream adaptor, mitochondrial antiviral signaling protein (MAVS). (Saito et al, 2008, Cui et al, 2008, Jiang et al, 2011,
Kowalinski et al, 2011, and Luo et al, 2011). MAVS was simultaneously discovered by four groups in 2005 (Seth et al, 2005), and is also known as virus-induced signaling adaptor (VISA) (Xu et al, 2005), interferon-β promoter stimulator 1 (IPS-1) (Kawai et al,
2005), and CARD adaptor inducing interferon-β (CARDIF) (Meyland et al, 2005).
MAVS is a 62 kDa protein that consists of an N-terminal CARD domain, a proline rich motif, and a transmembrane domain and is localized to the outer mitochondrial membrane (Kawai et al, 2005, Xu et al , 2005, Seth et al, 2005, and Meylan et al, 2005).
RLR interaction with MAVS results in the recruitment of various adaptor proteins involved in inflammatory and apoptotic signaling. Identified adaptors thus far include tumor necrosis factor (TNF) associated factors (TRAFs) 2, 3, and 6 (Saha et al,
2006), NF-κB essential modulator (NEMO)/inhibitor of NF-κB kinase subunit γ(Ikkγ)
13 (Zhao et al, 2007), Fas-associated death domain (FADD) (Balachandran et al, 2004),
receptor interacting protein-1 (RIP1) (Rajput et al, 2011), TNF receptor type 1-associated
DEATH domain protein (TRADD) (Michallet et al, 2008), and Caspases 8 and 10
(Takahashi et al, 2006). Several different signaling pathways resulting in the
transcription of proinflammatory cytokines are activated by MAVS interaction with
varying combinations of these proteins (Figure 1.4). For example, a signaling pathway
consisting of TRAF3 and NEMO/Ikkγ results in the activation of the serine/threonine
kinases TRAF family member-associated NFκB activator (TANK) binding kinase 1
(TBK-1) and inhibitor of κB (IκB) kinase ε (IKKε) (Yoneyama and Fujita, 2009). TBK-
1 and IKKε phosphorylate IRF3, which then dimerizes and moves into the nucleus where
it activates the transcription of type I interferons (interferon-α and interferon-β).
Interaction of MAVS with TRAF2/6, FADD, RIP1, and NEMO/Ikkγ results in the activation of the inhibitor of NF-kappaB (IkappaB) kinase (IKK) complex which phosphorylates the inhibitor of κB (IκB) family of molecules, marking these inhibitory proteins for proteosomal degradation (Zhao et al, 2007, Michaellet et al, 2008, Yoshida et al, 2008, Saha et al, 2008, and Chen, 2005). Free NF-κB dimerizes and translocates to the nucleus where it activates the transcription of target genes for proinflammatory and immunoregulatory molecules (Yoneyama and Fujita, 2009).
14
15 Several additional proteins have been shown to regulate RIG-I signaling by interaction with MAVS. Stimulator of interferon genes (STING) (Ishikawa and Barber,
2008), also known as mediator of IRF3 activation (MITA) (Zhong et al, 2008), endoplasmic reticulum (ER) interferon stimulator (ERIS) (Sun et al, 2009), or membrane tetraspanner associated with MHC class II (MPYS) (Jin et al, 2008), was demonstrated to interact with both MAVS and RIG-I. Interestingly, STING has no apparent effect on
MDA5-mediated antiviral signaling (Ishikawa and Barber, 2008). STING’s subcellular localization is under some dispute, as it has been reported to be localized on the outer mitochondrial membrane (Zhong et al, 2008) or the ER (Ishikawa and Barber, 2008 and
Sun et al, 2009). STING-deficient mice are highly susceptible both RNA and DNA virus infection (Ishikawa and Barber, 2008, Zhong et al, 2008, and Sun et al, 2009). Zhong et al (2008) demonstrated that STING interacts with TBK-1, IKKε, and IRF-3, suggesting that STING functions as a scaffolding protein for the recruitment of signaling complexes in RIG-I antiviral signaling.
A member of the NLR family of PRRs has also been reported to regulate RLR antiviral signaling. NLR family member X1 (NLRX1) was identified in 2008 by Moore et al in a screen to identify NLRs located in the mitochondria. While it contains a central
NBS domain and C-terminal LRR motifs, its N-terminal domain is atypical of the NLR family and was found to contain a mitochondrial targeting targeting sequence (Moore et al, 2008). Moore et al (2008) demonstrated that NLRX1 is localized on the outer mitochondrial membrane, and interacts with the CARD domain of MAVS. Interestingly, overexpression of NLRX1 repressed expression of interferon-β in reporter gene assays, and knockdown of endogenous NLRX1 in cell culture resulted in significantly increased
16 interferon-β levels in Sendai virus infected cells (Moore et al, 2008). Moore et al (2008) proposed that NLRX1 acts as a negative regulator of RIG-I antiviral signaling, but the mechanism for this inhibition is more complicated than simple binding competition. In the absence of activated RIG-I or exogenous MAVS, knockdown of NLRX1 doesn’t lead to increased signaling, suggesting that MAVS requires additional steps following the disruption of the interaction with NLRX1 to be competent for signaling (Moore et al,
2008).
1.4 RIG-I and MDA5 Activate Antiviral Signaling in Response to Distinct Viruses
1.4.1 RIG-I Recognizes a Diverse Range of Viruses
RIG-I has been shown to be activated in response to infection with a wide range of both positive- and negative-strand RNA viruses (Habjan et al, 2008). RIG-I detects the presence of many viruses that have a significant impact on human health, such as influenza and hepatitis C viruses (Table 1.1) (Pichlmair et al,2006 and Saito et al, 2008).
Upon its discovery, RIG-I was shown to promote IRF3 dimerization and IFN-β production with stimulation by either the dsRNA analog poly(I:C) or by infection of cells with negative-strand RNA viruses such as Newcastle disease virus (Yoneyama et al,
2004). RIG-I also has been shown to recognize a DNA virus, Epstein Barr virus. RIG-I is thought to recognize 5’-triphosphate RNA intermediates produced by RNA polymerase
III from AT-rich DNA in the viral genome (Samanta et al, 2008, Ablasser et al, 2009 and
Choi et al, 2009). In addition to viruses, RIG-I also activates an immune response in the presence of bacteria. Abdullah et al (2012) demonstrated that RIG-I senses nucleic acids
17 secreted from Listeria monocytogenes. RNA secreted by L. monocytogenes during its exponential growth phase was purified and found to be capable of inducing RIG-I- dependent interferon-β expression in macrophages, and the treatment of this RNA with calf intestinal alkaline phosphatase prior to transfection diminished interferon-β induction
(Abdullah et al, 2012). Additionally, Abdullah et al (2012) demonstrated that RIG-I also recognizes L. monocytogenes through the RNA polymerase III pathway described for the
Epstein Barr virus (Samanta et al, 2008, Ablasser et al, 2009 and Choi et al, 2009).
In 2006, 5’-triphosphate containing ssRNA generated by cells infected with the influenza virus was identified as a likely RIG-I PAMP (Pichlmair et al, 2006 and
Hornung et al, 2006). Saito et al (2008) identified the first positive-strand virus known to be recognized by RIG-I, hepatitis C virus (HCV). Furthermore, they determined that
RIG-I was activated by HCV in a sequence-dependent manner. They found that RIG-I activation was dependent upon the presence of a poly-U-rich region in the 5’-untranslated region (UTR) (Saito et al, 2008). However, many questions remain as to which RNA features are necessary for RIG-I to recognize viral RNA.
18 Virus Family Virus Name Reference Rhabdoviridae Vesicular stomatitis virus Yoneyama et al, 2005 (VSV) Kato et al, 2005 Rabies virus Hornung et al, 2006 Paramyxoviridae Sendai virus Yoneyama et al, 2005 Kato et al, 2005 Baum et al, 2010 New Castle Disease virus Kato et al, 2005 Respiratory syncytical Loo et al, 2008 virus (RSV) Measles virus Plumet et al, 2007 Nipah virus Habjan et al, 2008 Orthomyxoviridae Influenza A virus Kato et al, 2006 Influenza B virus Loo et al, 2008 Filoviridae Ebola virus Habjan et al, 2008 Arenaviridae Lassa virus Habjan et al, 2008 Bunyaviridae Rift Valley Fever virus Habjan et al, 2008 Hantaan virus Habjan et al, 2008 Crimean Congo Habjan et al, 2008 Hemorrhagic Fever virus Flaviviridae Hepatitis C virus (HCV) Saito et al, 2007 Japanese Encephalitis virus Kato et al, 2006 Dengue virus* Loo et al, 2008 Kato et al, 2011 West Nile virus* Loo et al, 2008 Kato et al, 2011 Reoviridae Orthoreovirus Loo et al, 2008 Bornaviridae Borna Disease virus Habjan et al, 2008 Table 1.1. Viruses recognized by RIG-I. * indicates viruses also recognized by MDA5.
19 1.4.2 MDA5 is Primarily Activated by Picornaviruses
Much less is known about the recognition of viral RNA by MDA5. Overall,
MDA5 appears to be activated by long cytosolic RNA duplexes produced during viral infections, while RIG-I has been shown to activated by shorter RNA duplexes that may contain an exposed 5’-triphsphate (Kato et al, 2006, Hornung et al, 2006, and Pichlmair et al, 2006). MDA5 has been shown to recognize members of the Picornavirus family, including encephalomyelocarditis virus (EMCV), Mengo’s virus, and Theiler’s virus
(Kato et al, 2005 and Gitlin et al, 2006). Additionally, several murine pathogens have been shown to be recognized by MDA5, including the Calicivirus norovirus-1 and the
Coronavirus murine hepatitis virus (McCartney et al, 2008 and Roth-Cross et al, 2008).
1.5 RIG-I Domain Structure and Function
1.5.1 Overall RIG-I Domain Structure
RIG-I is responsible for detecting the presence of viral RNA in mammalian cells, and stimulating a downstream signaling pathway that results in a subsequent antiviral response. As such, the structure and function of its individual domains reflects its critical role in innate immunity. RIG-I contains tandem N-terminal CARD domains, a central
RNA helicase/ATPase domain, and a C-terminal domain (Yoneyama et al, 2004 and
Kang et al, 2002). As described in the following sections, these domains perform distinct functions in the recognition of viral RNA and in the activation of antiviral signaling.
20
21
22 1.5.2 CARD Domains in RIG-I are Necessary for Signal Transduction
1.5.2.1 Structure and Function of RIG-I CARD Domains
RIG-I contains tandem N-terminal CARD domains (Yoneyama et al, 2004). This domain is a member of the death domain (DD) superfamily. The death domain (DD) superfamily includes four families: the death domain, death effector domain (DED), pyrin domain (PYD), and CARD domain (Bouchier-Hayes and Martin, 2002). The DD superfamily members play pivotal roles in the assembly of oligomeric complexes in inflammatory and apoptotic signaling responses (Bouchier-Hayes and Martin, 2002).
Apoptosis is a vital process for all multicellular organisms. It influences embryogenesis
and development, homeostasis, and the function of the immune system (Elmore, 2007).
Each of member of the DD superfamily interacts with other proteins exclusively through
homotypic protein-protein interactions (e.g. CARD-CARD, DD-DD, PYD-PYD, and
DED-DED) (Bouchier-Hayes and Martin, 2002).
The DD superfamily members all exhibit a characteristic structural topology.
They consist of six antiparallel α-helices arranged in a Greek-key motif (Figure 1.5). The
major structural differences are in the orientation of the individual helices. In CARD domains all six helices are almost parallel to each other (Bouchier-Hayes and Martin,
2002). The structure of the human MAVS CARD domain has been determined (Figure
1.7), as has the structure of full-length avian RIG-I, which includes the structure of both
CARD domains (Figure 1.8). The CARD domains of RIG-I and MDA5 share about 20%
sequence homology with that of MAVS and each other (Potter et al, 2008, Jiang et al,
2011, Kowalinski et al, 2011, and Luo et al, 2011) .
23
The CARD domains of MAVS (Potter et al, 2008), RIG-I (Luo et al, 2011, and
Kowalinski et al, 2011), and MDA5 (Potter et al, 2008) all have the canonical CARD fold
(Figures 1.7 and 1.8). The surface charge distributions of all three domains are asymmetric with positive and negative areas on opposite sides of the protein, which is characteristic of CARD domains and most likely involved in electrostatic interactions with other CARD-containing proteins (Potter et al, 2008). However, compared to
MAVS, the surface charge distributions of the first CARD domains of RIG-I and MDA5 differ significantly. This suggests that the CARD domains of the two RLRs may interact very differently with the MAVS CARD compared to one another (Potter et al, 2008).
24 The duplication of the CARD domains in RIG-I and MDA5 is unique to cytosolic PRRs, with the exception of the NLR Nod2 which also has tandem CARD domains (Yoneyama et al, 2004 and Bryant and Fitzgerald, 2009). The structure of full- length avian RIG-I showed that the tandem CARD domains form a head-to-tail unit
(Figure 1.8 and Luo et al, 2011 and Kowalinski et al, 2011). This forms a rigid functional unit of which both CARD domains are required for activation of antiviral signaling (Jiang et al, 2011, Kowalinski et al, 2011, and Luo et al, 2011). RIG-I lacking the CARD domains and a splice variant lacking 44 residues of the first CARD domain are dominant negative in vivo (Yoneyama et al, 2004). The ligand free structure of RIG-I demonstrated that an interaction between the second CARD domain and the helicase/ATPase domain is critical for the auto-inhibition of the protein in the absence of
RNA (Jiang et al, 2011, Kowalinski et al, 2011, and Luo et al, 2011).
25 1.5.2.2 Ubiquitination of the CARD Domains Regulates RIG-I Signal Transduction
Post-translational modification of the CARD domains has been demonstrated to
play a critical role in regulating RIG-I antiviral signaling (Oshiumi et al, 2012). Gack et
al (2007) suggested that upon detection of viral RNA, the first CARD domain becomes
accessible for interaction with the E3 ubiquitin ligase tripartite motif 25 (TRIM25).
TRIM25 was first reported to ubiquitinate RIG-I at lysine 172 and be necessary for the
activation of RIG-I antiviral signaling in 2007 (Gack et al, 2007). Further work
demonstrated that threonine 55 is necessary for this interaction; a mutation to isoleucine
at this residue in the hepatocyte cell line Huh7.5 confers an increased permissiveness to
hepatits C virus (HCV) replication (Sumpter et al, 2005 and Gack et al, 2008). TRIM25
binds to the first CARD of RIG-I, which leads to K63-linked polyubiquitination of lysine
172 in the second CARD domain (Gack et al, 2007). A point mutant at this residue with a severe defect in ubiquitination also exhibited significantly decreased binding to MAVS, while mutating threonine 55 to isoleucine abolished TRIM25 binding, ubiquitination of
RIG-I at K172, and MAVS binding (Gack et al, 2008). Overexpression of the T55I RIG-
I mutant in HEK293T cells failed to induce expression of either an interferon-β or NF-κB reporter construct (Gack et al, 2007 and Gack et al, 2008). This evidence suggests that the permissiveness of Huh7.5 cells to HCV replication is due to the disabling of the RIG-
I signaling pathway (Sumpter et al, 2005, Gack et al, 2007, and Gack et al, 2008). The importance of TRIM25 in the regulation of RIG-I antiviral signaling is further underscored by the mechanism of inhibition employed by the influenza A virus to suppress RIG-I signal transduction. The influenza A virus nonstructural protein 1 (NS1)
26 interacts with TRIM25, blocking its multimerization and ubiquitin ligase activities
(Garcia-Sastre et al, 1998 and Gack et al, 2009).
27 Two groups independently identified RIG-I as a substrate for a novel E3 ubiquitin ligase, Riplet, also known as RIG-I E3 ubiquitin ligase (REUL) or RNF135
(Oshiumi et al, 2009 and Gao et al, 2009). Riplet has been shown to interact with and ubiquitinate the RIG-I CTD and the second CARD domain of RIG-I (Oshiumi et al,
2009, Oshiumi et al, 2010, and Gao et al, 2009). K63-linked polyubiquitination of lysine residues 849, 851, 888, 901, and 907 were reported to be necessary for the activation of
RIG-I signaling (Oshiumi et al, 2009 and Oshiumi et al, 2010). In contrast to the findings of Oshiumi et al (2009), Gao et al (2009) demonstrated that Riplet-mediated ubiquitination of the second CARD domain resulted in a significant increase in RIG-I signal transduction. Oshiumi et al (2010) further demonstrated the importance of ubiquitination of the CTD with Riplet knockout mice. Riplet knockout mice demonstrate increased susceptibility to viral infection, decreased interferon production, and abrogated
RIG-I activation and ubiquitination (Oshiumi et al, 2010).
28 An alternative mechanism of ubiquitin-mediated regulation of the RIG-I signaling pathway was uncovered by Zeng et al, 2010. Utilizing an in vitro reconstituted RIG-I signaling system, they demonstrated that RIG-I activation was dependent upon RIG-I binding both dsRNA and unanchored K63-linked polyubiquitin chains, and that the polyubiquitin chains bound specifically to the CARD domains of RIG-I. Gack et al,
2008 concluded that the abrogation of interferon-β production in cells expressing the
RIG-I point mutant T55I was due to the inability of TRIM25 to bind and ubiquitinate
RIG-I. However, Zeng et al, 2010 demonstrated that the T55I mutant impaired the ability of RIG-I to bind polyubiquitin chains, while the ubiquitination-deficient K172R mutant was still capable of binding polyubiquitin chains and activating RIG-I signaling in reporter gene assays. This evidence suggested that contrary to previous reports,
TRIM25-mediated ubiquitination of the second RIG-I CARD domain is not necessary for
RIG-I antiviral signaling, and instead RIG-I’s association with unanchored K63-linked polyubiquitin chains was necessary for the activation of RIG-I signaling both in vitro and in vivo (Zeng et al, 2010).
29 RIG-I ubiquitination has also been shown to suppress RIG-I signaling. The E3 ubiquitin ligase Homo sapiens ring-finger protein 125 (RNF125) was shown to regulate cellular levels of RIG-I through K48-linked ubiquitination and subsequent proteosomal degradation. This functions as a negative feedback loop for the regulation of RIG-I, since
RNF125 expression is increases with the production of type I interferon (Arimoto et al,
2007). Friedman et al (2008) demonstrated that the deubiquitinase CYLD is also involved in the negative regulation of RIG-I antiviral signaling. CYLD is specific for
K63-linked ubiquitin chains and was shown to interact with RIG-I’s CARD domains and remove K63-linked polyubiquitin chains from the protein, thereby inhibiting downstream signaling (Friedman et al, 2008). This suggests CYLD regulates RIG-I signaling activity by the modulation of K63-linked ubiquitination of RIG-I.
1.5.3 RIG-I Helicase/ATPase Domain
RIG-I is an SF2 RNA helicase. SF2 RNA helicases are the largest class of enzymes in eukaryotic RNA metabolism. Proteins in this superfamily are involved in remodeling RNA structure or altering the components of ribonucleoprotein complexes in an ATP-dependent manner (Fairman-Williams et al, 2010). They are found in all kingdoms of life (Anantharaman, et al, 2002). RNA and DNA helicases are closely related and are classified into six superfamilies. The members of superfamilies 3 to 6 form oligomeric rings, while the non-ring forming helicases are in superfamilies 1 and 2
(Gorbalenya and Koonin, 1993 and Singleton et al, 2007). SF1 and SF2 are further divided into helicase families based on their sequence, structure, and function (Fairman-
Williams et al, 2010).
30 SF1 and SF2 helicases contain a structurally conserved helicase core formed by
two tandem RecA-like helicase domains. This helicase core is conserved among all SF1
and SF2 RNA helicases, and contains up to twelve characteristic sequence motifs. The
level of sequence conservation is high between family members, but decreases when
comparing helicases from different families (Fairman-Williams et al, 2010). These
sequence motifs function in nucleoside triphosphate (NTP) binding, coordination
between nucleic acid and NTP binding sites, and nucleic acid binding sites. The first
RecA-like domain in RIG-I contains the motifs Q, I (Walker A), Ia, Ib, Ic, II (Walker B),
IIa, and III. The second RecA-like domain contains motifs IV, V, and VI. Motifs Q, I,
and II in RIG-I are responsible for NTP binding, motifs Ia, Ib, Ic, IV, and V are RNA
binding sites, and motif III is involved in coordinating the RNA and NTP binding sites
(Jankowsky, 2011). In nearly all helicases, the helicase core domain is surrounded by N-
and C-terminal domains which have specific functions, such as nucleic acid binding,
protein binding, nuclease activity, and multimerization (Zhang et al, 2004, He et al, 2010,
Cui et al, 2008, and Klostermeier et al, 2009).
31
Helicases have been shown to demonstrate one of two mechanisms of nucleic acid duplex unwinding. The first is canonical duplex unwinding, which is utilized by most
DNA helicases and several viral RNA helicases. In this mechanism, the helicase binds to an adjacent single-stranded region, and translocates along the bound strand to displace the complementary strand (Singleton et al, 2007 and Pyle, 2008). RNA helicases in the
DEAD-box family use a distinct unwinding mechanism called local strand separation.
The helicase is directly loaded to the duplex region, and the strands are separated in an
32 ATP-dependent manner (Yang et al, 2007, Yang et al, 2006, Bizebard et al, 2004, and
Tijerina et al, 2006).
RNA helicases participate in nearly every aspect of RNA metabolism in a cell.
As a result they contribute to many disparate cellular functions, such as ribosome
biogenesis, pre-mRNA splicing, RNA export, and translation. Despite showing no
significant substrate specificity when studied in vitro, most RNA helicases whose
function is known only act in one specific process in vivo (Fairman and Jankowsky,
2010). In addition to unwinding RNA duplexes, RNA helicases have been shown to
translocate on dsRNA and facilitate strand annealing (Myong et al, 2009 and Fairman and
Jankowsky, 2010). All of these activities are postulated to play a role in RNA and RNP
structure conversions necessary for RNA metabolism in vivo (Jankowsky, 2011).
Not all proteins defined as helicases by their sequence motifs actually function
enzymatically to unwind nucleic acid duplexes in an ATP-dependent fashion. RIG-I was
shown by Takahasi et al (2008) to unwind RNA duplexes with a 3’-overhang of at least
five nucleotides. No other group has been able to replicate these results. Under the
published conditions, strand exchange independent of RIG-I could have occurred
(Takahasi et al, 2008). Currently, RIG-I is considered to have no RNA unwinding
activity. In 2009, Myong et al used single molecule fluorescence microscopy to
demonstrate that RIG-I was capable of translocating on a dsRNA without unwinding the
duplex. This translocation was ATP-dependent, and RIG-I was also able to translocate
on an RNA/DNA hybrid duplex, suggesting that RIG-I tracks along a single RNA strand
in the duplex. These results suggest that translocation could serve to allow RIG-I to scan
33 a long viral RNA looking for structural motifs that are RIG-I PAMPs (Myong et al,
2009).
RIG-I has RNA-dependent ATPase activity that has been shown to be critical for
the induction of antiviral signaling (Yoneyama et al, 2004). Gee et al, 2008 demonstrated
that RIG-I selectively hydrolyzes adenine nucleotide triphosphates, as ATP and dATP
were efficiently hydrolyzed in the presence of RNA. The K270A mutant of RIG-I has a
point mutation in the NTP binding motif I (Walker A motif). This mutant has been
shown to be incapable of signal transduction in vivo, yet retains RNA binding activity in vitro (Saito et al, 2008). While RIG-I’s RNA-dependent ATPase activity is clearly important for the activation of signaling, its precise role in the mechanism of RIG-I substrate recognition is not clear.
Sequence alignment of the RIG-I helicase/ATPase domain with other SF2 RNA helicases revealed that RIG-I’s helicase domain contains a unique insertion domain in between the two RecA-like domains (Fairman and Jankowsky, 2010). Other RLR family members also contain this domain (Yoneyama et al, 2005). The organization of the RIG-
I helicase domain is most similar to that of the Pyrococcus furiosus Hef helicase domain.
In the Hef helicase, this insertion domain plays a role in recognizing nucleic acid structures, leading to the suggestion that in the RLRs this domain may contribute to preferential substrate binding (Nishino et al, 2005).
Four groups simultaneously published the structure of RIG-I in 2011, including the structures of full-length Anas platyrhynchos (mallard duck) RIG-I (Kowalinski et al,
2011), the helicase domain of mouse RIG-I (Civril et al, 2011), and human RIG-
34 IΔCARD in the presence of RNA and NTP (Jiang et al, 2011, and Luo et al, 2011).
Avian RIG-I shares 53% sequence homology with human RIG-I, and its enhanced
thermal stability relative to human RIG-I is likely the reason that crystallization of full-
length human RIG-I has been unsuccessful thus far (Kowalinski et al, 2011). These
structures shed significant information on the conformational changes in the RIG-I
helicase domain that accompany ligand and NTP binding.
In the absence of ligand, the RIG-I helicase domain is in an open conformation in
which the first RecA-like domain (Hel1, residues 242-456) is not in direct contact with
the second RecA-like domain (Hel2, residues 458-469 and 609-745) (Figure 1.8)
(Kowalinski et al, 2011 and Luo et al, 2011). Hel1 and Hel2 both consist of α-helices
and β-strands. The insertion domain (Hel2i, residues 470-608) is present in the Hel2
domain and is comprised of five long, antiparallel α-helices (Luo et al, 2011). In this open conformation, the Hel1 and Hel2 domains are not able to engage to form a functional ATPase site (Luo et al, 2011 and Kowalinski et al, 2011). A motif truly unique to the RIG-I helicase domain is a V-shaped structure formed by two long α-
helices that connects Hel2 to the C-terminal domain (CTD) of RIG-I (Luo et al, 2011,
Jiang et al, 2011 and Kowalinski et al, 2011). This bridging or “pincer” domain, grips an
α-helix that protrudes from Hel1. The bridging domain provides a physical and
mechanical connection between RIG-I’s Hel1, Hel2, and CTD domains (Kowalinski et al,
2011 and Luo et al, 2011). Disruption of the bridging domain results in decreased
ATPase activity, RNA binding, and stimulation of an interferon-β/firefly luciferase
reporter gene (Luo et al, 2011). This evidence suggests a role for the bridging domain in
35 communication between the different domains of RIG-I (Jiang et al, 2011, Luo et al,
2011, and Kowalinski et al, 2011).
Upon dsRNA and NTP binding, the RIG-I helicase domain undergoes a conformational change in which all three subdomains move relative to one another to form a closed conformation (Figure 1.13 and Luo et al, 2011, Jiang et al, 2011, and
Kowalinski et al, 2011). The structure becomes more similar to those of other RNA helicases such as Dbp5 bound to ligand and NTP (Kowalinski et al, 2011). The structures generated by Kowalinski et al (2011) and Luo et al (2011) showed that RNA binding stimulates direct contact of the two RecA-like domains, creating a functional
ATPase site. The RIG-I helicase domain binds at the extreme blunt end of the RNA, with the three subdomains wrapping around the helix (Luo et al, 2011, Jiang et al, 2011, and
Kowalinski et al, 2011). The helicase domain contacts the sugar phosphate backbone of the RNA, particularly with the strand containing the 3’-terminus (Kowalinksi et al, 2011).
This is consistent with the electrostatic charge distribution of RIG-I, which shows that the
RNA is bound in a highly basic channel formed by the RIG-I helicase domain (Jiang et al, 2011 and Kowalinski et al, 2011). The structures of both the ligand free (Figure 1.8) and ligand bound (Figure 1.13) RIG-I helicase domain were consistent with the observation that RIG-I is incapable of hydrolyzing ATP without bound RNA (Jiang et al,
2011, Luo et al, 2011, and Kowalinski et al, 2011).
36
1.5.4 RIG-I C-Terminal Domain Functions in Ligand Binding and Multimerization
RIG-I contains a C-terminal extension with little sequence homology to other
proteins (Saito et al, 2007). This extension is called the C-terminal domain (CTD) or
regulatory domain (RD). The CTD was initially identified by Saito et al, 2007 as a
domain responsible for repressing the activation of antiviral signaling. They found that
deletion of the CTD resulted in constitutively active RIG-I in reporter gene assays in the
absence of viral infection (Saito et al, 2007). Saito et al (2007) expressed the isolated
CTD in Sendai virus infected cells and found it exhibited dominant-negative control of
RIG-I signaling. Furthermore, they found that either virus infection in cells or transfection of high levels of RIG-I promoted self-association of the protein (Saito et al,
2007). Based on this evidence they suggested that the CTD autoregulates RIG-I through internal interactions that favor a conformation where the CARD domains are masked and unable to signal. During virus infection, RIG-I dsRNA binding and conformational
37 changes occur that allow RIG-I to both multimerize and unmask the CARD domains to allow signal transduction (Saito et al, 2007).
Cui et al, 2008 examined the RNA-dependent ATPase activity of wild-type RIG-I and several deletion mutants to analyze the contributions of RIG-I’s domains to RNA- stimulated RIG-I activation. RIG-IΔRD, without the CTD, did not show RNA-dependent
ATPase activity on all substrates tested, suggesting a role for the CTD in substrate binding (Cui et al, 2008). A structural characteristic of viral RNA and a putative PAMP is an exposed 5’-triphosphate (Hornung et al, 2006 and Pichlmair et al, 2006). This moiety would be unique to nonself RNA in mammalian cells, since cellular mRNAs contain a 5’-7-methylguanosine cap and the uncapped RNAs produced by RNA polymerase III such as transfer RNAs and micro RNAs generally undergo post- transcriptional processing that modifies the 5’-end (Hopper, 2013). In 2006 RIG-I was shown to be activated in reporter gene assays by 5’-triphosphate containing RNA by several groups (Hornung et al, 2006 and Pichlmair et al, 2006). To examine the role of the RIG-I CTD in ligand binding, Cui et al, 2008 tested the RNA binding activity of the isolated RIG-I CTD on both a dsRNA and a 5’-triphosphate containing RNA and found the isolated CTD preferentially bound to an RNA containing a 5’-triphosphate.
Furthermore, competition of the ATPase activity of a RIG-I mutant lacking the CARD domains (RIG-IΔCARD) with an excess of RIG-I CTD inhibited the ATPase activity of
RIG-IΔCARD. This evidence suggested that the CTD of RIG-I contains an RNA binding site that preferentially binds to the 5’-triphosphate of RNA (Cui et al, 2008).
The final designation of the RIG-I CTD as a major RNA binding site in the protein was confirmed by the structure of the isolated domain (Cui et al, 2008 and
38 Takahasi et al, 2008). The structure of the RIG-I CTD was determined in 2008 by two groups with X-ray crystallography (Cui et al, 2008) and NMR spectroscopy (Takahasi et al, 2008), respectively. Both structures show that the CTD is a flat domain with a concave and convex side. It consists of two four-stranded and one two-stranded antiparallel β-sheets connected by small helical turns and organized in three leaves.
Analysis of the charge distribution and sequence conservation revealed that the concave side of the RD contained a highly positively charged groove with significant sequence conservation (Cui et al, 2008 and Takahasi et al, 2008). Mutation of several conserved residues in the positively charged groove reduced the binding affinity of the RIG-I CTD for 5’-triphosphate containing RNA in vitro (Cui et al, 2008). These same mutants also showed diminished signaling in interferon-β/firefly luciferase reporter gene assays in cell culture (Cui et al, 2008). The structural data suggests that RIG-I contains a critical RNA binding site in its CTD, and that this site may play a role in the recognition of the 5’- triphosphate of viral RNA.
The RIG-I CTD contains a tetrahedral zinc coordination site, which is formed by two protruding loops that laterally connect the two four-stranded β-sheets (Cui et al,
2008). These loops each contain two highly conserved cysteine residues to coordinate the zinc ion. Cysteine residues are characteristic of structural zinc coordination sites, and the zinc ion functions to stabilize the conformation of the protein (Alberts et al, 1998).
Cui et al (2008) demonstrated that mutation of any of these residues to arginine abrogated the RIG-I response to vesicular stomatitis virus (VSV) infection and found that a mutant with all of the conserved cysteine residues mutated to arginine was also unable to bind
RNA with a 5’-triphosphate in a pulldown assay. Taken together, this evidence suggests
39 that the zinc coordination site is a key structural motif and its integrity is necessary for
the binding of viral RNA (Cui et al, 2008).
RIG-I was first reported to form higher order oligomers in cells in 2007 (Saito et al, 2007). Cui et al, 2008 examined RIG-I multimerization further by performing gel filtration experiments with wild-type RIG-I and deletion mutants to determine if RIG-I bound a 5’-triphosphate containing RNA as an oligomer, and if so which domains were important for RIG-I multimerization. They demonstrated that wild-type RIG-I was capable of multimerization, but that the deletion mutant lacking the CTD eluted only as a monomer regardless of the presence of RNA suggesting that the CTD was required for
RIG-I multimerization (Cui et al, 2008). Performing the same experiment with the isolated CTD, they showed that the RIG-I CTD is capable of multimerization (Cui et al,
40 2008). Collectively, the RIG-I CTD has been shown to contain a major RNA binding
site and possibly contribute to RIG-I multimerization.
1.5.5 Molecular Mechanism of RIG-I Substrate Recognition
Ligand free RIG-I exists in an open conformation (Luo et al, 2011 and
Kowalinski et al, 2011). In this state RIG-I has low affinity for RNA (Luo et al, 2011).
The structural studies revealed that the autoinhibition of RIG-I is likely due to the
interaction between the second CARD domain and the Hel2i domain (Luo et al, 2011 and
Kowalinski et al, 2011). The CARD domains form a rigid functional unit and are linked head to tail. CARD1 is solvent exposed and is directly linked to the N-terminal helix of
CARD2 (Kowalinski et al, 2011). CARD2 plays a critical role in autoinhibition through its interaction with the Hel2i domain (Luo et al, 2011 and Kowalinski et al, 2011). This sequestering of the CARD2 domain by the Hel2i domain serves two functions. First, the
CARD2 domain is inaccessible for interaction with both ubiquitination enzymes and polyubiquitin chains, both of which have been shown to be necessary for the activation of
RIG-I signaling. Second, the interaction of the CARD2 with the Hel2i domain is
mediated by salt bridges and hydrophobic interactions (Luo et al, 2011). This interaction
is energetically stable and likely to sterically hinder the binding of RNA to RIG-I (Figure
1.15A) (Luo et al, 2011 and Kowalinski et al, 2011).
In the absence of RNA, RIG-I is unable to hydrolyze ATP due to a misalignment
of the helicase domains (Luo et al, 2011 and Kowalinski et al, 2011). A conformational
change is necessary to align the Hel1 and Hel2 domains to form an active site competent
41 for ATP hydrolysis and RNA binding (Luo et al, 2011, Kowalinski et al, 2011, and Jiang et al, 2011). Finally, the CTD is connected to the rest of the protein via a flexible linker
(Figure 1.15A) (Luo et al, 2011 and Kowalinski et al, 2011). It remains available to sense the presence of and bind to viral RNA in the cytoplasm (Cui et al, 2008 and
Takahasi et al, 2008). The auto-inhibition of RIG-I is crucial for the survival of cells, as inappropriate activation of inflammatory signaling pathways would be disadvantageous for a healthy cell (Akira et al, 2006). As such, the individual domains of RIG-I function to both recognize and respond to viral infection and to regulate the protein’s function to prevent inappropriate activation of antiviral signaling.
Upon RNA binding, a conformational change ejects the CARD domains from the helicase domain, allowing the helicase domain to interact with the dsRNA and inducing the active, closed conformation of RIG-I (Luo et al, 2011 and Kowalinski et al, 2011).
Luo et al (2011) and Kowalinski et al (2011) proposed that the CARDs are now available in solution for interaction with modifying enzymes and the adaptor protein MAVS
(Figure 1.15). The bridging domain may be involved in this conformational change
(Kowalinski et al, 2011, Luo et al, 2011, and Jiang et al, 2011). Through modeling of full-length RIG-I bound to dsRNA, Jiang et al, 2011 proposed that the bridging domain is located in close proximity to the CARD2 domain. This suggests that upon RNA binding, the bridging domain may function as a hinge for CTD movement and RIG-I activation
(Jiang et al, 2011).
42
The helicase domain primarily contacts the sugar phosphate backbone of the 3’-
strand, while the CTD primarily contacts the 5’-terminus of the RNA (Luo et al, 2011
and Jiang et al, 2011). The terminal end of the RNA duplex is capped by the RIG-I CTD, while the Hel1, Hel2, and Hel2i domains completely surround the duplex, occupying about ten base pairs (Luo et al, 2011, Jiang et al, 2011 and Kowalinski et al, 2011). All of the structures of RIG-I published thus far show RIG-I binding to RNA substrates as a
43 monomer (Kowalinski et al, 2011, Luo et al, 2011, and Jiang et al, 2011). The structures of free and ligand bound RIG-I provided a clear picture of the mechanism of RNA recognition. All of RIG-I’s functional domains play discrete roles in the discrimination of self from nonself RNA.
Multimerization plays a crucial role in the RLR antiviral signaling pathway. The downstream adaptor MAVS has been shown to aggregate on the mitochondrial membrane upon activation of antiviral signaling, and it has been suggested that this propagates the innate immune respone (Hou et al, 2011). A possible role for RIG-I multimerization was suggested by work done on the RLR family member MDA5 (Berke et al, 2012a and Berke et al, 2012b). In two separate papers, Berke et al (2012a and
2012b) used electron microscopy (EM) to demonstrate that MDA5 forms ATP-sensitive helical filaments on RNA duplexes, and that the CTD of MDA5 is critical for the cooperative assembly of these filaments. Jiang et al (2012) demonstrated that the binding of K63-linked polyubiquitin chains was necessary for the multimerization of both RIG-I and MDA5. Finally, RIG-I was shown to multimerize on viral dsRNA in a length- dependent manner by Patel et al (2013). They also suggested that the multimerization of
RIG-I was dependent upon the presence of ATP, and that RIG-I bound as a monomer in the absence of ATP (Patel et al, 2013). Thus far, the role of multimerization in the recognition of viral RNA by RIG-I remains unclear.
44 1.6 RIG-I Recognizes Multiple Structural Features of Viral RNA
Once RIG-I was identified as a PRR, work began in earnest to identify the RIG-I
PAMP. Definition of the RIG-I PAMP is thought to provide information crucial for the development of vaccines, adjuvants, and antiviral drugs. The first clues as to what viral
RNA features are recognized by RIG-I were uncovered by groups that noted nonspecific effects in cells transfected with small interfering RNAs (siRNA) (Kim et al, 2004 and
Marques et al, 2006). Kim et al (2004) noted that the interferon response was not activated by synthetically produced siRNAs, but interferon induction was observed in the presence of siRNAs produced with in vitro transcription. The RNAs produced by in vitro transcription contained a 5’-triphosphate because phage polymerase transcription is template-dependent and occurs primer independent from the 5’- to the 3’-end of the RNA
(Kochetkov et al, 1998). Furthermore, removal of the 5’-triphosphate from the siRNAs eliminated the expression of interferon (Kim et al, 2004). Meanwhile, Marques et al,
2006 observed that the induction of interferon expression as a result of transfection with siRNA could be abrogated if the siRNAs contained two nucleotide 3’-overhangs, similar to endogenous Dicer products. These findings suggest that RIG-I is using several RNA features to distinguish between self and non-self RNA.
Further evidence for the importance of the 5’-triphosphate was published by two groups in 2006. Hornung et al (2006) screened several types of RNA molecules for their ability to induce IFN-α in human primary monocytes in order to identify putative RIG-I ligands. They demonstrated that RNAs containing a 5’-triphosphate induced IFN-α, and that removal of the 5’-triphosphate abolished the expression of IFN-α (Hornung et al,
2006). They also showed that neither a 5’-monophosphate or diphosphate was sufficient
45 to induce IFN-α to the same levels as the 5’-triphosphate (Hornung et al, 2006). This
finding was further supported by the work of Pichlmair et al (2006) and Cui et al (2008).
The detection of influenza virus RNA by RIG-I was dependent upon the RNA containing
at least a 5’-monophosphate, and dsRNA was found to be dispensible for this recognition
(Pichlmair et al, 2006). Cui et al (2008) found that RIG-I RNA binding and ATPase
stimulation in vitro was significantly enhanced for ssRNA with a 5’-triphosphate compared to a blunt dsRNA.
There is minimal evidence for sequence-dependent recognition of viral RNA by
RIG-I. Saito et al (2008) showed that compared to a transcript of the full-length genome
of HCV, a transcript in which the 3’-NTR was deleted was non-immunostimulatory. The
3’-NTR of HCV contains a poly-U/UC rich region that is the putative RIG-I PAMP
(Saito et al, 2008). In addition to U-rich sequences, the authors also showed similar levels of interferon induction for A-rich sequences (Saito et al, 2008). Davis et al (2012) discovered similar evidence for the influenza virus, in which a U/A rich region in the 3’-
NTR was shown to induce interferon-β independent of the presence of a 5’-triphosphate when transfected into human lung epithelial cells.
The role of RNA secondary structure in RIG-I RNA recognition was initially unclear. Marques et al (2006) proposed that synthetic blunt-ended dsRNA is recognized by RIG-I. This result was only seen in a glioblastoma cell line (T98G), and was not found in either HeLa or HT1080 cells (Marques et al, 2006). However, Pichlmair et al
(2006) found that the presence of dsRNA was dispensible for the recognition of influenza virus RNA and Cui et al (2008) found that dsRNA was not sufficient for the activation of
RIG-I ATPase activity in vitro.
46 In 2009, two groups demonstrated that dsRNA is necessary for the recognition of
viral RNA by RIG-I, and also demonstrated why it appeared that ssRNA was recognized by RIG-I. They showed that activation of RIG-I signaling required dsRNA with a base-
paired 5’-triphosphate. Furthermore, they discovered that the activation of RIG-I signaling seen in the presence of ssRNA was due to the source of these substrates (Schlee et al, 2009 and Schmidt et al, 2009). They were all produced with in vitro transcription
(Hornung et al, 2006, Pichlmair et al, 2006, and Cui et al, 2008).
Schlee et al (2009) demonstrated that a chemically synthesized ssRNA with a 5’- triphosphate was incapable of activating RIG-I signaling, while the same RNA produced with in vitro transcription was stimulatory. They found that the RNA produced with in
vitro transcription contained double-stranded hairpin species and complementary
sequences from RNA template RNA transcription by the phage polymerase (Cazenave
and Uhlenbeck, 1994, Triana-Alonso et al, 1995, and Schlee et al, 2009). They postulate
that the RIG-I PAMP for negative-strand RNA viruses such as influenza is the panhandle
region of the ssRNA viral genome (Schlee et al, 2009 and Schmidt et al, 2009). The
panhandle consists of a region with highly complementary 5’- and 3’-sequences that
forms a short region of dsRNA with a blunt end (Hsu et al, 1987). The role of viral
panhandles as RIG-I PAMPs was further emphasized by work from Marq et al. Marq et
al 2010a demonstrated that the panhandles of Arenavirus and Bunyavirus genomes are
not recognized by RIG-I due to a unique replication mechanism that results in a single
nucleotide overhang, leaving the 5’-triphosphate unpaired. They further showed that this
panhandle structure can act as a RIG-I decoy, competitively inhibiting the stimulation of
47 RIG-I signaling when co-transfected with a blunt dsRNA with a 5’-triphosphate (Marq et
al, 2010b).
Despite these initial findings, in the field the role of the 5’-triphosphate in RIG-I
RNA recognition remains controversial. In the course of this thesis work, several groups
have addressed this question. Jiang et al (2011) found that the presence of a 5’-
triphosphate had no significant effect on either RIG-I RNA binding. Meanwhile, work
from Anna Marie Pyle’s lab shows a significant effect by the 5’-triphosphate. Vela et al
(2012) demonstrated that the presence of a 5’-triphosphate resulted in an increase in affinity of RIG-I for dsRNA of nearly two orders of magnitude compared to the same
RNA duplex with a 5’-hydroxyl. They obtained similar results in 2013, where their data showed that the presence of a 5’-triphosphate does enhance the affinity of RIG-I for dsRNA, but they did not see any enhancement of RIG-I RNA-dependent ATPase activity being mediated by the 5’-triphosphate (Kohlway et al, 2013).
Many questions remain regarding the role of RIG-I RNA binding, multimerization, and RNA-dependent ATPase activity in the recognition of the RIG-I
PAMP. Several features of the PAMP have been identified, including a dependence on
RNA secondary structure (dsRNA) and the presence of a 5’-triphosphate (Schlee et al,
2009 and Schmidt et al, 2009). However, the mechanism by which RIG-I utilizes its different activities to distinguish between self and non-self RNA remains to be elucidated. In this thesis, I investigated the contributions of RIG-I RNA binding, multimerization, and ATPase activity to RNA recognition in vitro. I characterized RIG-I
RNA binding, multimerizataion, and ATPase activity with a series of model substrates to
48 determine how RNA secondary structure and 5’-end structure to contribute to substrate recognition by RIG-I.
Initially, the work of Hornung et al (2006) and Pichlmair et al (2006) suggested that the minimal RIG-I PAMP was ssRNA containing a 5’-triphosphate. Later, Schlee et al (2009) and Schmidt et al (2009) conclusively demonstrated that these results were likely a consequence of the use of phage polymerase in vitro transcribed RNA containing short regions of dsRNA that were being recognized by RIG-I. This finding highlighted the limitations of phage polymerase in vitro transcription, and underscored the need for additional methods of generating 5’-triphosphate RNA to serve as RIG-I ligands. In chapter 2, I describe a novel method of generating 5’-triphosphate containing ssRNA from commercially purchased RNA oligos with the Tetrahymena L21 ribozyme. With this method, 5’-triphosphate 5’-end labeled RNA is easily and reliably produced, and the use of the L21 ribozyme also will allow for the production of RNAs with additional 5’- end structures which could be used to fully investigate the contribution of RNA 5’-end structure to RIG-I RNA recognition.
Chapters 3 and 4 of this thesis quantitatively analyze the interaction of RIG-I with a series of model substrates to determine the contributions made by RNA secondary structure, RNA duplex length, and the 5’-triphosphate to RIG-I RNA recognition. I demonstrate that RNA binding and RNA-dependent ATPase activity are dependent upon the presence of dsRNA, and both are independent of RNA duplex length and the presence of a 5’-triphosphate. Additionally, in the presence of ATP the dissociation rate constant for RIG-I dramatically increases, suggesting that in the presence of an activating ligand,
RIG-I’s interaction with the RNA changes. Collectively, these findings suggest that the
49 minimum requirement for RNA binding by RIG-I is dsRNA, and that once bound to the
RNA duplex RIG-I is then able to recognize additional RNA features such as the 5’-
triphosphate, which is indispensable for the activation of antiviral signaling (Schlee et al,
2009 and Schmidt et al, 2009) and utilize its RNA-dependent ATPase activity, which is a
sign the protein has adopted a signaling competent conformation.
In chapter 5, I quantitatively analyzed the effect of available blunt duplex ends on
RIG-I RNA binding and RNA-dependent ATPase activity. Structural studies of ligand- bound RIG-I suggested that it requires a blunt duplex end for efficient interaction with an
RNA (Luo et al, 2011, Kowalinski et al, 2011, and Jiang et al, 2011). RIG-I tightly bound to all RNA duplexes tested, even one with no available blunt duplex ends.
Strikingly, stimulation of RIG-I RNA-dependent ATPase activity required a blunt duplex
end. The abolishment of RNA-dependent ATPase activity by ssRNA overhangs suggests
that RIG-I distinguishes between viral and cellular RNA by proofreading RNA duplexes
with its ATPase activity in order to interrogate the presence of a blunt end. Overall, my
findings suggest a model where RIG-I is sampling dsRNA, and using its RNA binding,
multimerization, and RNA-dependent ATPase activities to examine the RNA and identify
multiple features before recognizing a viral RNA as a PAMP.
50 Chapter 2
Development of L21-ScaI Ribozyme as a Method to Generate RIG-I RNA
Substrates
2.1 Rationale for using the L21 Ribozyme for Producing RIG-I Substrates in vitro
The minimal RIG-I PAMP has been identified as a dsRNA of at least seven base pairs, with a base paired 5’-triphosphate (Schlee et al, 2009, Schmidt et al, 2009, Marq et al, 2010a, and Marq et al, 2010b). The initial identification of this PAMP was stymied by researchers unwittingly producing contaminating dsRNA while in vitro transcribing RIG-
I ligands with phage polymerase (Schlee et al, 2009 and Schmidt et al, 2009). Since this limitation of in vitro transcription was identified in 2009 by Schlee et al and Schmidt et al, the field has been searching for methods to produce 5’-triphosphate RNA that is not contaminated by dsRNA due to RNA template RNA dependent transcription by phage polymerase (Cazenave and Uhlenbeck, 1994). In the literature, 5’-triphosphate containing RNA has been generated by in vitro transcription (Cui et al, 2008), chemical synthesis (Schlee et al, 2009 and Schmidt et al, 2009), or single-site RNase T1 cleavage
(Vela et al, 2012). These methods have significant drawbacks. The generation of short ssRNAs with T7 RNA polymerase in vitro is limited by the low yield produced by conventional methods (Ambion). Vela et al (2012) produced a fourteen nucleotide 5’- triphosphate containing RNA through in vitro transcription of a longer RNA followed by selective cleavage of the longer transcript with RNase T1. Unfortunately, this limits the sequence composition of the desired RNA, since RNase T1 cleaves selectively after
51 guanine residues (Vela et al, 2012). Additionally, the polymerase can inadvertently
produce RNAs with short double-stranded regions (Schlee et al, 2009 and Schmidt et al,
2009). While chemically synthesized 5’-triphosphate RNA is commercially available, it
is prohibitively expensive and unavailable to be purchased as individual oligos of any
desired sequence as of the writing of this thesis.
To generate 5’ end-labeled 5’-triphosphate RNA I used the Tetrahymena
thermophilia L-21 ribozyme. The L-21 ribozyme cleaves RNA after a specific sequence,
and covalently links that attacking guanosine to the 5’-end of the 3’-end of the substrate
(Zaug et al, 1986b). When this reaction is performed in the presence of α-32P-GTP, the
5’-end of the substrate produced is α-32P-GTP. The L-21 ribozyme is derived from the
400 nucleotide self-splicing intron in the gene encoding the large subunit (26S) of the
Tetrahymena ribosome (Zaug and Cech, 1986b). The first evidence of the self-splicing
activity of the intron was provided in 1982 by Kruger et al. They demonstrated that an in
vitro transcribed fragment of the T. thermophilia pre-rRNA underwent processing to
remove the intron and ligate the flanking exons in the absence of any protein. This
reaction only requires the presence of GTP or other guanosine containing analog, Mg2+,
and salt to occur (Cech et al, 1981 and Kruger et al, 1982).
Shortened linear forms of the intron (e.g. L-19 and L-21 ribozymes) have been
shown to process exogenous RNA molecules. The L-19 and L-21 transcripts of the ribozyme lack the first nineteen and twenty-one nucleotides of the intron, respectively, and both perform a sequence-specific endonuclease reaction analogous to the first step of
splicing (Cech et al, 1981, Kruger et al, 1982, and Zaug et al, 1986b). Ribozymes
derived from the self-splicing intron cleave RNA very specifically after the sequence
52 CUCU in the presence of Mg2+ and exogenous guanosine. The products of this reaction are the 5’ end of the RNA ending with a 3’-hydroxyl group and the 3’ end of the substrate with the attacking guanosine covalently linked at the 5’ end (Zaug and Cech, 1986b)
(figure 2.1). In addition to site-specific endonuclease activity, the ribozyme has also phosphotransferase, nuclease, and nucleotidyl transferase activities under specific reaction conditions (Zaug and Cech, 1986a and Zaug and Cech, 1986b). With this system, I could create 5’ end-labeled 5’-triphosphate RNA substrates by including α32P-
GTP, which also resulted in the cleavage product being end-labeled. Additionally, since
the ribozyme requires only guanosine for its enzymatic activity, it may be possible to
create 3’-end labeled RNAs with different 5’ end structures and examine how the 5’ end
structure affects RIG-I RNA binding in great detail.
A number of guanosine analogs have already been shown to be enzymatically active for the self-splicing of the pre-RNA in vitro (Bass and Cech, 1984). This investigation determined that the intron exhibits significant specificity for guanosine as a substrate and that guanosine is binding to a specific site on the RNA. Their work indicated that modification of the 2’- or 3’-hydoxyl of the ribose sugar and modification of the exocyclic substituents of the pyrimidine ring abrogated the intron’s splicing activity. Conversely, substitution at the 5’-hydroxyl or modifications to the imidazole ring had very little effect on the splicing activity (Bass and Cech, 1984). This work suggested that it may be possible create RNAs with different 5’ end structures using the
L-21 ribozyme and guanosine analogs that are tolerated by the preRNA intron.
53
2.2 Characterization of L21-ScaI ribozyme activity on RIG-I substrates
To examine the activity of the ribozyme on RIG-I substrates, I designed several
RNAs containing a sequence complementary to the ribozyme’s internal guide sequence
54 (IGS) (highlighted in yellow in table 2.1 and shown by the red line in figure 2.1, step 1), to allow the ribozyme to bind to the substrate (figure 2.1, step 2). The ribozyme was first activated by incubation with salt and Mg2+ to allow for proper folding of the RNA. The cleavage reaction was initiated by the addition of precursor RNA and GTP (figure 2.1, step 1). The IGS in the ribozyme bound to the recognition sequence in the precursor
RNA, and GTP bound to the guanosine binding site in the ribozyme. The precursor RNA was cleaved immediately following the recognition site, and the ribozyme released two products: a product analogous to the 5’-exon that contains the recognition sequence and a downstream product that has the exogenous GTP covalently added to its 5’-end with a phosphodiester bond (figure 2.1, steps 2 and 3).
RNA Name RNA Sequence
R10L21P 5’ CCC UCU AGC ACC GUA 3’ R13L21P 5’ CCC UCU AGC ACC GUA AAG 3’ R13L21PBot2 5’ CCC UCU ACU UUA CGG UGC 3’ R84-L21 5’ GGG AGA CAA AGU AUA GCC CUC UAA CAC AAA CGC GUC AUG AUU GAC ACU UUA CGG UGC CCA GCA UCA AUG ACA UCA GCA UCU AC 3’ Table 2.1. L21-ScaI RNA Precursor Substrate Sequences. Yellow highlighting indicates the sequence complementary to the ribozyme’s IGS.
Prior to performing a large-scale reaction to generate 5’-triphosphate 5’-end labeled RNA, I characterized the activity of the ribozyme on the RIG-I substrates to demonstrate efficient cleavage of these particular substrates. The precursor substrates were incubated with the L-21 ribozyme, guanosine triphosphate (GTP), excess Mg2+, and salt. A trace amount of radiolabeled (5’-end labeled) RNA precursor and a trace amount
55 of α-32P-GTP were included in the reactions to be able to track both cleavage products
with autoradiography (figure 2.2).
First, I examined the dependence of the reaction on both GTP and the ribozyme,
to ensure that the product being generated in the reactions was the desired thirteen
nucleotide 5’-triphosphate 5’-end labeled product. The ribozyme’s catalytic activity was
dependent upon the presence of exogenous GTP or other guanosine analog and Mg2+
(Zaug et al, 1986b). In the absence of either ribozyme or GTP, I saw no accumulation of the thirteen nucleotide product, and the eighteen nucleotide precursor RNA did not disappear after incubation for sixty minutes (figure 2.2, left two panels). The shorter bands with weak signal present on all the gels are due to degradation of the 5’-end labeled precursor RNA. The reaction conditions are extremely harsh due to the presence of high concentrations of Mg2+ and the high temperature.
In the presence of the ribozyme and GTP, I saw efficient cleavage of the eighteen
nucleotide (R13L21PBot) precursor RNA over a sixty minute time course. The eighteen
nucleotide precursor substrate disappeared while the thirteen nucleotide 5’-triphosphate
and 5’-end labeled product accumulated (figure 2.2, right two panels). In reactions
containing a 5’-end labeled precursor, I also saw the accumulation of the five nucleotide
upstream cleavage fragment, while in the absence of 5’-end labeled precursor RNA, I see
the appearance of a labeled thirteen nucleotide product starting at thirty minutes (figure
2.2, right two panels). These data indicate that the L-21 ribozyme efficiently cleaves
RNAs containing its recognition sequence, and that it is adding a α-32P-GTP nucleotide to
the 5’-end of the downstream cleavage product.
56
I also examined L21 cleavage of the fifteen nucleotide RNA precursor (R10L21P) and the other eighteen nucleotide RNA precursor (R13L21P). The downstream sequence of the two eighteen nucleotide precursor RNAs is different (Table 2.1). In the presence of the ribozyme and GTP, I saw cleavage of the 5’-end labeled substrates (figure 2.3).
Accumulation of the 5’-triphosphate 5’-end labeled thirteen nucleotide product was also seen for the additional eighteen nucleotide substrate (figure 2.3A). This product (ten nucleotides) was not observed for the fifteen nucleotide substrate despite efficient cleavage of the precursor (figure 2.3B). However, under the large-scale cleavage conditions, a correctly sized end-labeled product is formed (figure 2.3C). There are several possibilities to explain the lack of accumulation of the ten nucleotide product in the characterization reactions: (i) The 5’-triphosphate 5’-end labeled product was being
57 degraded in the harsh reaction conditions. (ii) The ribozyme was catalyzing the
hydrolysis of the substrate. The phosphodiester bonds in both the splice sites and the
circularization sites in the T. thermophilia self-splicing intron have been shown to be unusually susceptible to hydrolysis (Zaug et al, 1984, Zaug et al 1985, and Inoue et al,
1986). Zaug et al (1984) postulated that the local secondary and tertiary structure of the
RNA at these cleavage sites in the intron can increase the reactivity of the phosphate that
is attacked by the nucleophile to facilitate cleavage at a particular site in the RNA.
58 Finally, I examined if guanosine analogs could be substituted in place of GTP.
For this data set, I used an eighty-four nucleotide precursor RNA R84-L21. Like the eighteen and fifteen nucleotide precursor RNAs described previously, L-21 ribozyme cleavage of the eighty-four nucleotide precursor did not occur in the absence of either the ribozyme or GTP (figure 2.4, right panels). Consistent with the results with the shorter precursor substrates, in the presence of GTP, the eighty-four nucleotide substrate was efficiently cleaved within sixty minutes (figure 2.4, center panel). I chose to use GDP and the non-hydrolyzable analog GMP-PNP. GDP was previously shown to have very little effect on the splicing activity of the T. thermophilia intron by Bass and Cech (1984).
The ribozyme efficiently cleaved the precursor RNA in the presence of both GDP and
GMP-PNP within sixty minutes (figure 2.4, right two panels).
59 2.3 Discussion
In this chapter, the data presented demonstrate that the L-21 ribozyme derived from the T. thermophilia self-splicing intron efficiently generates RNAs containing a 5’- end labeled 5’-triphosphate that can serve as RIG-I substrates in vitro and possibly in vivo. Utilizing several different RNA precursor substrates, I characterized the L-21 ribozyme’s activity in the presence of GTP (figures 2.2, 2.3, and 2.4). The 5’- triphosphate containing 5’-end labeled RNAs produced with this method were used in
Chapter 3 to produce short RNA duplexes to act as RIG-I substrates for RNA binding.
This method allowed the generation of short 5’-triphoshate containing RNAs that can not be generated by in vitro transcription alone, and removes the sequence restrictions necessary for further site-specific processing with RNases. Furthermore, it can be used with commercially available RNA oligos of nearly any final sequence desired by the user.
As suggested by the work of Bass and Cech (1984), the ribozyme also efficiently cleaved RNAs in the presence of the GTP analogs GDP and GMP-PNP. Since the modifications to these nucleotides were both to the 5’-hydroxyl of ribose, the L-21 ribozyme was still capable of binding these analogs and utilizing them in the cleavage reaction (figure 2.5). This data suggests that different guanosine analogs could be included in the ribozyme cleavage reactions in order to generate RNAs with different 5’- end structures. This has potential uses not only for producing substrates to use in investigating the interaction of RIG-I with RNA, but for examining the effects of 5’-end structure on any other RNA binding protein that is interacting with the 5’-end.
60 Chapter 3
RIG-I Binding to dsRNA
3.1 Introduction
RIG-I RNA binding is the first step of substrate recognition. The first RIG-I
PAMP identified was the exposed 5’-triphosphate that is present on viral ssRNA
(Pichlmair et al, 2006 and Hornung et al, 2006). The ideal RIG-I substrate for the activation of antiviral signaling in cell culture was then further defined in 2009 as an
RNA duplex of at least seven base pairs containing a base-paired 5’-triphosphate (Sclee et al, 2009, Schmidt et al, 2009, Marq et al, 2010a). Additionally, these published reports suggested that the stimulation of RIG-I signaling in vivo seen with in vitro transcribed ssRNA containing a 5’-triphosphate was due to contaminating dsRNA produced by the
T7 RNA polymerase (Schlee et al, 2009, Schmidt et al, 2009, Marq et al, 2010a, and
Marq et al, 2010b). This data raised questions surrounding which RNA features are necessary for RIG-I recognition and suggested that RIG-I may identify more than one
RNA feature when distinguishing between self and non-self RNA in cells.
In order to address this question, I performed a comprehensive analysis of RIG-I
RNA binding to define how RNA binding by RIG-I contributes to the mechanism of
RNA recognition. I examined wild-type RIG-I protein binding to a series of RNA model substrates to determine how RNA secondary structure and 5’-end structure affect RIG-I-
RNA complex formation (Table 7.1 in section 7.1). I measured the binding affinities of
RIG-I for a ssRNA substrate and a series of dsRNA substrates ranging from ten to thirty- six base pairs in length. I also examined if the 5’-triphosphate contributed to RIG-I RNA
61 binding. To determine the contributions of RIG-I’s individual domains to its RNA binding activity, I examined the RNA binding activity of a series of RIG-I deletion mutants (Figure 3.1).
RIG-I CARD CARD Helicase/ATPase RD 1-925 aa
∆CARD Helicase/ATPase RD 242-925 aa
∆RD CARD CARD Helicase/ATPase 1-796 aa
∆CARD∆RD Helicase/ATPase 242-796 aa
Figure 3.1. Schematic of wild-type RIG-I and RIG-I deletion mutants.
3.2 Results
3.2.1 Purification of wild-type and mutant RIG-I
I purified wild-type and mutant recombinant RIG-I proteins (figure 3.2). The protein concentrations were determined with the Bradford assay. The active protein concentrations were determined by normalizing the RNA-dependent ATPase activity of individual preps. RIG-IΔCARD should retain its RNA binding activity due to the presence of the CTD. Initial characterization of the RIG-I CTD demonstrated it is critical for RIG-I RNA binding and activation of its ATPase activity (Cui et al, 2008 and
Takahasi et al, 2008). However, a different group demonstrated this mutant to have constitutive ATPase activity (Gee et al, 2008). RIG-IΔCTD and RIG-IΔCARDΔCTD
(the isolated helicase domain) have been shown to be deficient in ssRNA binding and
62 have significantly impaired ATPase activity (Cui et al, 2008 and Luo et al, 2011). I purified wild-type RIG-I and all of the deletion mutants using the multi-step approach described in section 6.2. The final purified proteins are shown in figure 3.2.
3.2.2 RIG-I binds ssRNA with low affinity despite presence of a 5’-triphosphate
I measured the binding affinity of RIG-I for a thirty-six nucleotide ssRNA both with and without a 5’-triphosphate using a gel shift assay performed under equilibrium conditions as described in section 7.4. The establishment of equilibrium was confirmed by incubating the low concentrations of RIG-I with RNA over a time course ranging from one minute to one hour to confirm that complex formation was consistent over time.
Overall, RIG-I bound weakly to the ssRNA, regardless of whether it had a 5’-hydroxyl or a 5’-triphosphate, with dissociation constants of K1/2 = 129.8 ± 3.8 nM and K1/2 = 57.1 ±
1.9 nM, respectively (figure 3.3). The results indicated a significant enhancement of binding to ssRNA containing a 5’-triphosphate, with RIG-I binding the ssRNA with a 5’- triphosphate with an affinity about two-fold higher than that of the 5’-hydroxyl. The
63 weak binding of RIG-I to ssRNA was consistent with the lack of stimulation of antiviral signaling of ssRNA seen by Schlee et al (2009) and Schmidt et al (2009). This suggests that while RIG-I is capable of binding to ssRNA, it is unable to adopt a signaling competent conformation. The structures of ligand-bound RIG-I by Luo et al (2011),
Kowalinski et al (2011), and Jiang et al (2011) showed that upon binding an RNA duplex the subdomains of RIG-I’s helicase domain formed an ATP hydrolysis competent conformation that requires contacts with both strands of the RNA duplex.
3.2.3 5’-triphosphate does not impact RIG-I binding to dsRNA
I next examined RIG-I’s affinity for dsRNA with and without a 5’-triphosphate to establish whether the presence of the 5’-triphosphate has any impact on RIG-I’s RNA binding. As in section 3.2.3, I determined the dissociation constants of RIG-I for a thirty- six base pair duplex both with and without a single 5’-triphosphate under equilibrium
64 conditions. RIG-I bound the thirty-six base pair dsRNA lacking a 5’-triphosphate with a
K1/2 = 11.4 ± 1.1 nM. The results for the same duplex containing a single 5’-triphosphate were similar. RIG-I bound the duplex containing a 5’-triphosphate with a K1/2 = 9.7 ± 3.1
nM. Unlike binding to ssRNA, in which the presence of a 5’-triphosphate resulted in a
small enhancement in RIG-I’s affinity, the presence of a single 5’-triphosphate on a
thirty-six base pair RNA duplex had no significant effect on RIG-I RNA binding (figure
3.4).
This result is consistent with some published reports (Jiang et al, 2011), however other
groups have shown that the presence of a 5’-triphosphate on a dsRNA results in
significantly tighter binding compared to an RNA duplex lacking a 5’-triphosphate
(Schlee et al, 2009 and Vela et al, 2012). To confirm this result, I measured the affinity
of RIG-I for both ten and thirteen base pair RNA duplexes with and without a single 5’-
65 triphosphate. These duplexes differ from the thirty-six base pair duplex in the source of the 5’-triphosphate. The 5’-triphosphate-containing RNA in the thirty-six base pair duplex was produced with in vitro transcription. The 5’-triphosphate-containing strand for the ten and thirteen base pair duplexes was produced with the cleavage of a precursor substrate by the L-21 ribozyme as described in chapter 2. The use of these substrates allowed me to exclude any effects of in vitro transcription (Schlee et al, 2009 and
Schmidt et al, 2009).
I measured the dissociation constants for RIG-I binding to the ten base pair duplex with and without a 5’-triphosphate using gel shift assays under equilibrium conditions. The results were similar to those for the thirty-six base pair duplex, in that the presence of a 5’-triphosphate had no effect on RIG-I’s affinity for the RNA duplex
(figure 3.5). However, despite the similar dissociation constants, RIG-I binding to the ten base pair duplex lacking a 5’-triphosphate exhibited diminished binding amplitude (figure
3.5B). This is likely due to decreased duplex stability under RIG-I reaction conditions, which would diminish the amount of radiolabeled duplex available for RIG-I binding and increase the amount of radiolabeled ssRNA in the unbound fraction.
66
I next examined RIG-I binding to a thirteen base pair duplex both with and
without a 5’-triphosphate. The slightly longer duplex is more stable under the reaction
conditions, removing the possibility of significant dissociation of the radiolabeled duplex impacting the overall binding amplitude. The results were consistent with those of both
the thirty-six and ten base pair duplexes, in that the dissociation constants of RIG-I for
the 5’-hydroxyl and 5’-triphosphate-containing RNA duplexes differed by less than two-
fold, suggesting the 5’-triphosphate does not significantly contribute to RIG-I RNA
binding (5’-hydroxyl: K1/2 = 10.5 ± 1.2 nM and 5’-triphosphate: K1/2 = 20.5 ± 1.1 nM)
(figure 3.6).
67
3.2.4 RIG-I binds dsRNA tightly regardless of RNA duplex length
One intriguing observation from section 3.2.3 was that RIG-I formed multiple
RNA-protein complexes on the thirty-six bases pair duplex, but primarily formed one
RNA-protein complex on the ten and thirteen base pair duplexes (figures 3.4, 3.5, and
3.6). To further investigate the role of RNA duplex length in RIG-I RNA binding I analyzed RIG-I RNA binding to a series of RNA duplexes ranging from ten to thirty-six base pairs in length, none of which contained a 5’-triphosphate. Using gel shift assays performed under equilibrium conditions, I measured the dissociation constants of RIG-I for ten, thirteen, sixteen, nineteen, and thirty-six base pair RNA duplexes.
As shown in figure 3.7, RIG-I’s affinity for dsRNA was not impacted by duplex length. RIG-I has an affinity for RNA duplexes between ten and thirty-six base pairs
68 ranging from K1/2 = 10 to K1/2 = 20 nM, indicating very tight binding (figure 3.7A).
However, including the sixteen and nineteen base pair duplexes revealed multiple RIG-I-
RNA complexes with increasing duplex length (figures 3.4, 3.5, 3.6, and 3.7B). The
smaller duplexes (10 and 13 bp) formed primarily one RIG-I-RNA complex, while the
longest (36 bp) formed at least two RIG-I-RNA complexes. These results suggest that
multimerization of RIG-I is correlated with RNA duplex length. Additionally, RIG-I
multimerization was first apparent on the thirteen and sixteen base pair duplexes,
suggesting that the minimal binding site size is between six to eight base pairs. This is
consistent with the work of Marq et al (2010a), who showed that a minimum of seven
continuous base pairs was necessary for stimulation of RIG-I-mediated antiviral
signaling.
As this work was being performed, a number of other groups published work suggesting
RIG-I RNA binding is dependent on RNA duplex length. First, the structures of ligand bound and ligand free RIG-I were published (Kowalinski et al, 2011, Jiang et al, 2011,
Luo et al, 2011, and Civril et al, 2011). The structures of ligand bound RIG-I
demonstrated that the minimal binding site size is about ten base pairs (Jiang et al, 2011 and Luo et al, 2011). In 2013, Patel et al demonstrated that RIG-I multimerization on a
Sendai virus defective interfering RNA was dependent on the length of a stem present in the predicted stem-loop structure. These findings are consistent with the observation of
RIG-I-RNA complex formation being length dependent in my gel shift assays (figures
3.4, 3.5, 3.6, and 3.7B).
69
3.2.5 RIG-I binding to RNA duplexes of at least 16 bp involves two species of RNA- protein complexes
RIG-I has been shown to bind activating RNA substrates as a dimer (Cui et al,
2008 and Luo et al, 2011). My gel shift assays with the thirty-six base pair duplexes show two discrete protein-RNA complexes forming in the binding titration, with the lower molecular weight complex forming early and disappearing as the higher molecular weight complex appeared (figure 3.8A). Using a model for a coupled equilibrium, I calculated the binding parameters for each individual complex (figure 3.8B). For wild-
70 type RIG-I binding to the thirty-six base pair duplex with a 5’-triphosphate, formation of
the first complex (K1/2 = 56.7 ± 17.2 nM) is much weaker than formation of the second complex (K1/2 = 2.21 ± 0.9 nM) (table 3.1 and figure 3.8). This indicates cooperative binding, in which the binding of the first unit of RIG-I significantly increases the affinity of the subsequent unit of RIG-I for the RNA. This finding held true for a thirty-six base pair duplex lacking a 5’-triphosphate, in which formation of the lower molecular weight complex (K1/2 = 76.1 ± 23 nM) was again much weaker than the formation of the higher
molecular weight complex (K1/2 = 2.07 ± 0.9 nM) (table 3.1). Formation of RIG-I
multimers is energetically favored, and RIG-I binds dsRNA cooperatively regardless of
the presence of a 5’-triphosphate. The role of cooperative binding in RIG-I substrate
recognition and antiviral signaling remains unclear, since Kohlway et al (2013)
demonstrated that a RIG-I monomer is sufficient to induce an interferon-β/firefly
luciferase reporter gene.
71
Section 3.2.6 RIG-I multimerization depends on RNA duplex length
Sections 3.2.4 and 3.2.5 demonstrated that the stoichiometry of RIG-I-RNA complexes formed under equilibrium conditions appears to correlate with RNA duplex length. In order to confirm the stoichiometry of the complexes formed on the different
72 length RNA duplexes, I performed stoichiometric binding titrations. A known amount of
RNA (29 nM unlabeled and 1 nM labeled) was incubated with increasing amounts of
RIG-I. At four points throughout the RIG-I titration, the amount of RIG-I was either
equimolar with the total RNA concentration, or represented multiples (2X, 3X, and 4X)
of the total RNA concentration. RNA-protein complexes were resolved with native
PAGE (figure 3.9).
The resulting binding curves indicate the stoichiometry of the complex (figure
3.10). Two straight lines are fit to the binding curve, the first line marks the concentration range where the fraction bound is steadily increasing. The second line marks the concentration range where RIG-I has saturated the RNA. The intersection point of these two lines indicates the concentration of RIG-I at which binding to the RNA is saturating. The stoichiometry of the complex can then be determined by dividing the saturating RIG-I concentration by the RNA concentration.
I quantitatively examined the stoichiometry of RIG-I-RNA complexes with the thirteen, nineteen, and thirty-six base pair RNA duplexes. The results indicate that
73 despite the formation of primarily one apparent RNA-protein complex on the gel under
equilibrium conditions (figure 3.6, right gel), RIG-I is binding the thirteen base pair
duplex as a dimer (figure 3.10). This result was somewhat surprising, since the structures
of ligand-bound RIG-I identified the minimal binding site size as nine to ten base pairs
(Kowalinski et al, 2011, Luo et al, 2011, and Jiang et al, 2011). However, Marq et al
(2010b) demonstrated that only seven continuous base pairs were required to activate
RIG-I signaling in vivo and in section 3.2.4 my data showed that RIG-I is binding a sixteen base pair duplex as a multimer. Collectively, these findings suggest the true minimal binding site size of RIG-I may be between seven and eight base pairs. The dimer forming on the thirteen base pair duplex may be unstable in the gel shift assays, but still forming due to the duplex length being nearly two minimal binding site sizes.
74 RIG-I bound the nineteen base pair duplex as a dimer and the thirty-six base pair duplex as a trimer, both of which are consistent with the nine to ten base pair minimal binding site size identified by Luo et al (2011), Kowalinski et al (2011), and Jiang et al
(2011) (figure 3.11). This result is also consistent with a shorter seven or eight base pair minimal binding site size, as the nineteen base pair duplex would consist of 2 to 2.5 RIG-
I binding sites and the thirty-six base pair duplex would consist of 4 to 5 RIG-I binding sites. However, only two of the potential binding sites in the thirty-six base pair duplex would occupy the duplex ends, and RIG-I binding to interduplex regions may be weaker, since the CTD would not be able to interact with a duplex end. This discrepancy in affinity for the two types of binding sites in RIG-I may result in the lag seen in the initial points in the titration for the thirty-six base pair duplex. Higher order multimers of RIG-I are unlikely to be resolved on the gel shifts due to their size precluding migration of the
RNA-protein complex into the gel.
75
Interestingly, the presence of the 5’-triphosphate on the thirty-six base pair RNA changed the stoichiometry. In the presence of a 5’-triphosphate, RIG-I bound the thirty- six base pair duplex as a dimer (figure 3.12). This is inconsistent with the trimer formed on the thirty-six base pair duplex with a 5’-hydroxyl, demonstrating that the 5’- triphosphate influences RIG-I multimerization (figure 3.12). This finding suggests that
76 while the 5’-triphosphate does not impact the affinity of RIG-I for an RNA, that recognition of the 5’-triphosphate by RIG-I may result in a conformational change to the protein that affects the binding of subsequent units to the RNA. Collectively, the stoichiometric binding results demonstrate that RIG-I multimerization is dependent upon
RNA duplex length, and that the presence of a 5’-triphosphate on dsRNA changes the mode of RIG-I RNA binding.
3.2.7 RIG-I deletion mutants demonstrate that the CTD is the major RNA binding site
In addition to wild-type RIG-I, I also purified two point mutants that are deficient in ATPase activity and three deletion mutants (Figure 3.1). The RIG-I CTD has been
77 shown to contain a strong RNA binding site (Cui et al, 2008 and Takahasi et al, 2008), which binds to the 5’-end of a blunt RNA duplex (Luo et al, 2011, Kowalinski et al,
2011, and Jiang et al, 2011). Initial characterization of the RIG-I CTD demonstrated that it is critical for RIG-I RNA binding and activation of its ATPase activity (Takahasi et al,
2008 and Cui et al, 2008). Additionally, there are RNA binding sites in the
helicase/ATPase domain, which have been shown to make important contacts to the
phosphate backbone of RNA (Kowalinski et al, 2011, Luo et al, 2011, and Jiang et al,
2011).
Since the RIG-IΔCARD mutant retains all of the RNA binding sites and the
CARD domains have not been shown to function in RIG-I RNA binding, this protein might be expected to retain its RNA binding. Furthermore, it has been shown to have intact and constitutively active ATPase activity (Gee et al, 2008). RIG-IΔCTD and RIG-
IΔCARDΔCTD (the isolated helicase domain) have been shown to be deficient in RNA binding and have significantly impaired ATPase activity (Gee et al, 2008 and Cui et al,
2008). In section 3.2.1 I have shown all of the purified deletion mutants (figure 3.2). I examined the RNA binding and ATPase activity of the RIG-I deletion mutants to help define the role of the helicase domain in substrate recognition.
I first examined the RNA-dependent ATPase activity of the RIG-I deletion mutants to determine the contribution of the different functional domains to RIG-I ATP hydrolysis. I qualitatively tested RNA-dependent the RNA-dependent ATPase activity of the RIG-I mutants under conditions of excess RNA (1 µM) and ATP (1 mM). ATPase stimulation was only seen for wild-type RIG-I and RIG-IΔCARD in the presence of double-stranded RNA (table 2). This finding is consistent with published data for wild-
78 type RIG-I (Schlee et al, 2009 and Schmidt et al, 2009). The mutants lacking the CTD,
RIG-IΔCTD and RIG-IΔCARDΔCTD both exhibited no stimulation of ATPase activity under the conditions tested (table 2). This finding is consistent with the purported role of the CTD in functional RNA binding, and suggests that the lack of ATPase stimulation is a result of the mutant proteins having significantly diminished RNA binding or being unable to form an ATP hydrolysis competent conformation (Cui et al, 2008, Takahasi et al, 2008, Luo et al, 2011, Kowalinski et al, 2011, and Jiang et al, 2011).
79 I next performed gel shift assays to examine RIG-IΔCARD RNA binding to ssRNA and dsRNA. I used the thirty-six nucleotide ssRNAs with and without a 5’- triphosphate, and the thirty-six base pair duplexes with and without a 5’-triphosphate so that I could directly compare the binding of the mutant protein with that of the wild-type.
Like wild-type RIG-I, RIG-IΔCARD binds more weakly to ssRNA, and the presence of a
5’-triphosphate has no significant effect on the affinity of RIG-IΔCARD for ssRNA
(figure 3.13A). RIG-IΔCARD binds dsRNA with an affinity that is three- to four-fold higher than that of ssRNA (figure 3.13). The RNA binding results suggest that the lack of ATPase stimulation of RIG-IΔCARD by ssRNA may be due to the diminished RNA binding (table 2 and figure 3.13). This finding was further supported when work published by Vela et al (2012) during the preparation of this thesis project demonstrated that a similar RIG-I mutant lacking the CARD domains exhibited no significant differences compared to wild-type RIG-I in its ability to bind ssRNA and dsRNA.
Gel shift assays with RIG-IΔCARD also show two discrete protein-RNA complexes forming in the gel, with the lower molecular weight complex forming early and disappearing as the higher molecular weight complex appeared. Like wild-type RIG-
I, using a model for a coupled equilibrium, I calculated the binding parameters for each individual complex (figure 3.14). For RIG-IΔCARD, formation of the first complex is much weaker than formation of the second complex (table 2). This suggests that like wild-type RIG-I, RIG-IΔCARD also binds dsRNA cooperatively
80
81
The CTD of RIG-I is thought to be necessary for both its RNA binding and
ATPase activities (Cui et al, 2008, Takahasi et al, 2008, and Gee et al, 2008). I performed gel shift assays with RIG-IΔCTD and RIG-IΔCARDΔCTD (isolated helicase domain) binding to a thirty-six base pair RNA duplex containing a 5’-triphosphate. RIG-
82 IΔCTD does not bind RNA at the concentrations tested, and the isolated helicase domain
binds very weakly to RNA, with a dissociation constant in excess of 1 µM (figure 3.15).
These results are consistent with the lack of ATPase activity seen in the presence of
RNA, and suggest that lack of ATPase correlates with diminished RNA binding.
Collectively, the ATPase and RNA binding results suggest that the RNA binding sites in the helicase domain appear to make a small energetic contribution to overall RNA binding, providing additional evidence that the CTD of RIG-I contains the major RNA binding site. This finding is consistent with work published during the course of this thesis project, when the isolated helicase domain of RIG-I was shown to bind to a fourteen base pair dsRNA with a 5’-triphosphate (KD = 760 ± 20 nM) (Vela et al, 2012).
As expected, the CARD domains of RIG-I do not participate in RNA binding
directly, as wild-type RIG-I and RIG-IΔCARD both bind dsRNA tightly and
83 cooperatively. The apparent lack of contribution the RNA binding sites in the helicase domain make to overall RIG-I RNA binding in vitro does not underscore their importance in the mechanism of RIG-I substrate recognition shown by the structural studies. It is possible that the deletion mutants are inactive due to improper folding of the shortened protein. However deletion of the CTD does result in a significant decrease in
RIG-I RNA binding, suggesting a critical role for this domain in RIG-I RNA recognition.
3.3 Discussion
3.3.1 RIG-I only binds dsRNA, and the 5’-triphosphate impacts RIG-I multimerization
In this section, I examined RIG-I RNA binding to a series of model substrates to determine the contributions made by RNA 5’-end structure and RNA secondary structure to RIG-I substrate recognition. One RNA feature recognized by RIG-I is dsRNA, since
RIG-I bound ssRNA significantly weaker than dsRNA (figures 3.3 and 3.4). This finding is consistent with studies in the literature demonstrating that ssRNA was incapable of activating antiviral signaling in vivo alone, and the presence of a ssRNA overhang on an
RNA duplex severely diminished or abrogated antiviral signaling activation (Schlee et al,
2009, Schmidt et al, 2009, Marq et al, 2010a, Marq et al, 2010b). Furthermore, as shown in figure 3.7, RIG-I tightly bound dsRNA regardless of RNA duplex length. Taking into account both the findings presented in sections 3.2.2, 3.2.3, and 3.2.4 and the published reports, the lack of signaling activation seen in the presence of ssRNA is likely due to
RIG-I being unable to adopt a signaling competent conformation when binding ssRNA.
84 The first viral RNA feature identified as a putative RIG-I PAMP was the 5’- triphosphate of viral RNA (Pichlmair et al, 2006 and Hornung et al, 2006). This moiety is a strong candidate due to its unlikely presence in a healthy cell, since cellular mRNAs have a 5’-methyl guanosine cap. It remains unknown why RIG-I doesn’t recognize the
5’-triphosphates of tRNAas and rRNAs (Pichlmair et al, 2006). My results demonstrate that the presence of a 5’-triphosphate on either a thirty-six nucleotide ssRNA or a thirty- six base pair dsRNA does not enhance RIG-I RNA binding (figures 3.3 and 3.4). Since conflicting reports have been published showing enhancement of binding mediated by the
5’-triphosphate, I further examined the effect of the 5’-triphosphate on two shorter RNA duplexes and found no significant effect of the 5’-triphosphate on the affinity of RIG-I for dsRNA (figures 3.5 and 3.6) (Luo et al, 2011 and Vela et al, 2012). The role of the
5’-triphosphate in RIG-I RNA binding thus remains controversial.
While several groups have demonstrated that the presence of a 5’-triphosphate enhances either RNA binding (Vela et al, 2012 and Kohlway et al, 2013) or RNA- dependent ATPase activity in vitro (Cui et al, 2008 and Schlee et al, 2009), my data and that of other groups suggest that the 5’-triphosphate makes no significant contribution to
RIG-I RNA binding (Jiang et al, 2011). The conflicting data may be due to sequence effects that either amplify or dampen the phosphate effect. RIG-I has been shown to have some sequence preferences in the 3’-NTR of HCV and influenza virus RNAs (Saito et al,
2008 and Davis et al, 2012). Davis et al (2012) reported that activation of RIG-I signaling by an A/U rich portion of the 3’-NTR was independent of the presence of a 5’- triphosphate.
85 Varying sequences of synthetic RNAs have been used in the literature to
investigate RIG-I. The RNA sequences used in this thesis consist of a heterogeneous mix of all four nucleotides in a random sequence (table 7.2). Vela et al (2012) showed over a
100-fold increase in the affinity of RIG-I for a duplex with a 5’-triphosphate, however the
5’-triphosphate containing RNA was created by single-site RNase T1 cleavage which
limited the sequence of the remainder of the oligo. The RNA was C-rich, and consisted
primarily of G-C base pairs (Vela et al, 2012). Kohlway et al (2013) also showed
enhancement of RIG-I RNA binding in the presence of a 5’-triphosphate with a sequence
consisting of GC repeats ranging from ten to thirty base pairs in length. They saw a ten
to twenty-five fold increase in the affinity of RIG-I for the same duplex with a 5’-
triphosphate (Kohlway et al, 2013). Interestingly, they also measured the binding affinity
of RIG-I for hairpins with a heterogeneous sequence containing a 5’-triphosphate and
obtained dissociation constants similar to those presented in chapter 3 but did not
measure the affinity of RIG-I to the same hairpins in the absence of a 5’-triphosphate
(Kohlway et al, 2013). Jiang et al (2011), who saw no significant difference in the
affinity of RIG-I for the same RNA substrate with and without a 5’-triphosphate, also
used an RNA duplex with a random, heterogeneous sequence. The discrepancies suggest
in that in addition to interrogating the RNA for the presence of dsRNA and possibly a 5’-
triphosphate, RIG-I may also be searching for certain sequence elements in order to
discriminate between cellular and viral RNAs.
86 3.3.2 RIG-I binds dsRNA cooperatively and multimerization is dependent on duplex length
In section 3.2.4, I demonstrated that RIG-I formed higher molecular weight
complexes with RNA duplexes at least sixteen base pairs long (figure 3.7). I confirmed
the stoichiometry of the complexes with stoichiometric binding titrations, which showed that RIG-I multimerization was dependent upon RNA duplex length. The results were consistent with the published minimal binding site size of about ten base pairs for RIG-I,
as RIG-I bound the nineteen base pair duplex as a dimer and the thirty-six base pair
duplex as a trimer (figures 3.10 and 3.11) (Luo et al, 2011 and Jiang et al, 2011). Since
multimerization was most pronounced for the thirty-six base pair duplex, I further
analyzed the results from this duplex by fitting them to a model for a coupled equilibrium
(figure 3.8). The calculated binding parameters indicate that RIG-I is binding dsRNA
cooperatively, with formation of the ES complex being significantly weaker than that of the E2S complex (table 3.1).
The role of cooperative binding in RIG-I substrate recognition and antiviral signaling is unclear. Initially multimerization of RIG-I was shown to be necessary for the activation of antiviral signaling (Cui et al, 2008), however Kohlway et al (2013) demonstrated that a ten base pair RNA hairpin which RIG-I bound as a monomer was sufficient for the induction of an interferon-β/firefly luciferase reporter gene. Kohlway et al (2013) suggest a model of RIG-I antiviral signaling activation in which RIG-I monomers bind to the duplex termini, and then these 1:1 RIG-I:RNA complexes multimerize via their CARD domains. Patel et al (2013) postulated that RIG-I multimerization plays a role in signal amplification in vivo. Even though a RIG- I
87 monomer is sufficient to activate antiviral signaling, cooperative binding of multiple
RIG-I molecules to a long viral RNA could be responsible for signal propagation. RIG-I
molecules bound to interduplex regions are unlikely to adopt a signaling competent
conformation, but their ability to bind to the dsRNA regardless could increase molecular
crowding at the viral RNA and enhance the opportunity for binding to an exposed duplex
end and activating antiviral signaling.
3.3.3 Major RNA binding site is in RIG-I CTD
Measuring the RNA affinity and RNA-dependent ATPase activity of a panel of
RIG-I deletion mutants (figure 3.1) demonstrated that a major substrate binding site in
RIG-I is likely present in the CTD. RIG-I mutants lacking the CTD demonstrated significant defects in RNA binding (figure 3.15). A RIG-I mutant lacking the CARD
domains, which are not involved in RNA binding, had behavior very similar to wild-type
RIG-I (figures 3.13 and 3.14). Removal of the CTD also abrogated the stimulation of
RIG-I RNA-dependent ATPase activity by dsRNA (table 3.2). The lack of ATPase activity seen in the mutants lacking the CTD is likely due to diminished RNA binding, suggesting that the CTD plays a critical role in RIG-I RNA recognition. While it is difficult to exclude the possibility that the defects exhibited by the deletion mutants are due to defects in protein folding, work done by other groups supports the role of the CTD as a significant RNA binding site in RIG-I. The importance of the CTD in RNA recognition was further underscored by structural studies demonstrating that it contains a positively charged patch identified as a 5’-triphosphate binding site (Cui et al, 2008 and
88 Takahasi et al, 2008) and by structures of the ligand-bound full-length protein showing the CTD capping the 5’-strand of the dsRNA substrate (Luo et al, 2011, Jiang et al, 2011, and Kowalinski et al, 2011).
89 Chapter 4
RIG-I ATPase Activity Recognizes the Presence of dsRNA
4.1 Introduction
As an SF2 RNA helicase, RIG-I contains a structurally conserved helicase domain. Proteins in the SF2 superfamily are thought to be involved in remodeling RNA
structure or altering the components of ribonucleoprotein complexes in an ATP-
dependent manner, including RNA folding, the unwinding of RNA duplexes, and
displacement of proteins from RNA. RIG-I has demonstrated RNA-dependent ATPase
activity (Gee et al, 2008 and Cui et al, 2008), and has been shown to translocate along an
RNA duplex in vitro (Myong et al, 2009).
Many questions remain as to what role ATP hydrolysis plays in RIG-I antiviral signaling. Saito et al (2007) utilized a nucleotide binding deficient mutant of RIG-I,
RIG-I K270A, to show that the RNA-dependent ATPase activity of RIG-I is critical for the activation of signaling in vivo. However, this mutant RIG-I protein still retained the ability to bind RNA qualitatively in vitro (Saito et al, 2007). RIG-I ATPase stimulation
is dependent upon RNA binding, as it has very low basal ATPase activity (Gee et al,
2008). The structures of RNA-bound RIG-I demonstrate that RIG-I’s helicase domain
only forms an ATP hydrolysis competent conformation when it is bound to an RNA
duplex end, and that this conformational change also makes the CARD domains
accessible for activation of downstream signaling (Luo et al, 2011, Kowalinski et al,
2011, and Jiang et al, 2011). A blunt duplex end with a base paired 5’-triphosphate was
identified in vivo as a RIG-I PAMP by Schlee et al (2009) and Schmidt et al (2009).
90 Collectively these findings suggest that RIG-I ATPase activity plays a role in substrate
discrimination by serving as a marker of PAMP identification.
In order to investigate the role of RIG-I RNA-dependent ATPase activity in
substrate recognition, I first investigated the RNA binding activity of an ATPase-
deficient RIG-I mutant and wild-type RIG-I RNA binding and multimerization in the
presence of various nucleotides representing different stages of the ATP hydrolysis cycle.
Next I measured the dissociation off rates of RIG-I from dsRNA in the presence of
different nucleotides to examine the effect of nucleotide on RIG-I-RNA complex
formation. Finally I examined RIG-I’s RNA-dependent ATPase activity in response to a series of model substrates to determine how RNA secondary structure and 5’-end
structure affect RIG-I ATPase stimulation.
4.2 Results
4.2.1 Nucleotide has no effect on RIG-I RNA affinity, but promotes complex dissociation
The gel shift experiments shown in chapter 3 to measure the affinity of RIG-I for
various model RNA substrates were all performed in the absence of nucleotide. In order
to determine if the presence of ATP would impact RIG-I RNA binding, I examined the affinity of RIG-I for a thirty-six base pair RNA duplex with a single 5’-triphosphate in the presence of ATP and additional nucleotides. I performed RIG-I binding titrations under equilibrium conditions in the presence of ATP, the non-hydrolyzable analog
ADPNP, the pre-hydrolysis state analog ADP-NaF, the transition state analog ADP-AlF4,
and post hydrolysis ADP. RIG-I-RNA complexes were resolved with native PAGE.
91 Overall, RIG-I’s affinity for a thirty-six base pair dsRNA with a 5’-triphosphate was
unaffected by the presence of ATP, ADP, the nonhydrolyzable analog ADPNP, ADP-
AlFx ,a transition state analog, and ADP-NaF, which is present in ADP-AlFx and is
serving as a control (table 4.1).
Similar to the multiple RNA-protein complexes formed on the thirty-six base pair
duplex shown in section 3.2.5, RIG-I formed two discrete RNA-protein complexes in the
presence of all nucleotides described above. Using the model for the coupled equilibrium
described in section 3.2.5 I determined the binding parameters for both complexes
formed in the gels. Similar to RIG-I-RNA complex formation in the absence of
nucleotide, in the presence of all nucleotides tested the affinity of the higher molecular
weight complex for the RNA was significantly higher than that of the lower molecular
weight complex (table 4.1). In the presence of ADP-AlFx, which is a transition state analog, the formation of the first complex became significantly weaker compared to the
ADP-AlFx other nucleotides tested (K1/2 = 181 ± 153 nM vs. K1/2 = ~50 nM) (table 4.1). This
suggests that the affinity of RIG-I for RNA during the transition state of ATP hydrolysis
is significantly decreased.
92 However, one of the drawbacks of this approach is the possibility of the nucleotide diffusing out of the complexes as they run into the gel, since the reactions are performed under equilibrium conditions. This would mean that the complexes formed in the gel are only RIG-I and RNA, instead or RIG-I, RNA, and nucleotide, and that the binding parameters being measured are those of RIG-I-RNA complex formation in the absence of nucleotide.
I next utilized a different approach to examine qualitatively if RIG-I RNA binding and multimerization were affected by the presence of ATP. RIG-I-RNA complexes were chemically cross-linked with glutaraldehyde, and resolved with SDS-
PAGE (figure 4.1). I compared RIG-I binding to 13 and 36 bp duplexes with and without a 5’-triphosphate, in the presence and absence of ATP. As expected, ligand-free RIG-I is a monomer (figure 4.2, lane 2, Saito et al, 2008 and Cui et al, 2008).
Under the reaction conditions necessary to obtain visualization of RIG-I-RNA complexes with Coomassie blue staining, RIG-I bound the thirty-six base pair duplex without a 5’-triphosphate primarily as a dimer, with an additional higher molecular
93 weight complex forming that is likely the trimer (figure 4.2, lane 7). The calculated molecular weight of a RIG-I trimer and the thirty-six base pair RNA duplex exceeds the molecular weight standard used, making it impossible to distinguish between a trimer or additional higher order oligomers that may have formed. RIG-I also formed a dimer with the thirty-six base pair duplex with a 5’-triphosphate (figure 4.3, lane 9). These results are consistent with the complexes formed in the gel shift assays under equilibrium conditions in section 3.2.3, and with the quantitative stoichiometry determined in section
3.2.6.
RIG-I bound the thirteen base pair RNA duplex as a monomer, regardless of the presence of the 5’-triphosphate (figure 3.17, lanes 3 and 5). This result is consistent with
94 the complexes formed in the gel shift assays under equilibrium conditions, in which only
one RIG-I-RNA complex primarily forms. However, this result is inconsistent with the
quantitative stoichiometry determined in section 3.2.6, in which RIG-I binds the thirteen
base pair duplex as a dimer, despite only one complex forming in the gel shift assays.
This suggests that while RIG-I is capable of binding the thirteen base pair duplex as a
dimer, this dimer may be unstable and is inefficiently cross-linked to the RNA.
There is ample evidence in the literature that RIG-I multimer formation is likely
an intrinsic part of RIG-I-RNA binding, even though the structures show monomers (Luo
et al, 2011, Kowalinski et al, 2011, and Jiang et al, 2011) and Kohlway et al (2013)
demonstrated it is not required for the activation of antiviral signaling in vivo. In addition
to the data in section 3.2.6, other groups have also demonstrated length dependent multimerization of RIG-I on dsRNA (Binder et al, 2011, Luo et al, 2011, Kohlway et al,
2013, and Patel et al, 2013). The discrepancy between the stoichiometric binding data in section 3.2.6 and the chemical crosslinking data in this section can possibly be explained by distinguishing between functional, or ATP hydrolysis competent, and nonfunctional
RNA binding by RIG-I. When RIG-I binds to duplexes whose length is less than two minimal binding site sizes, there may be only enough available base pairs for one RIG-I molecule to adopt a functional, ATP hydrolysis competent complex. The functional binding of one RIG-I molecule to the duplex would not prevent a second RIG-I from interacting in a nonfunctional manner with the remaining exposed duplex. Nonfunctional binding of RIG-I to RNA, such as its weaker interaction with ssRNA, does not stimulate
RIG-I RNA-dependent ATPase activity and has been shown to be significantly weaker
(section 3.2.2). The formation of a dimer on the thirteen base pair duplex would be an
95 intermediate step in RIG-I-RNA complex formation, where the dimer consists of one
ATP hydrolysis competent RIG-I bound to the blunt duplex end, and one RIG-I molecule
engaged in a nonfunctional and less stable interaction with a partial binding site.
While the inclusion of ATP in RIG-I binding reactions under equilibrium conditions appeared to have no effect on the affinity of RIG-I for RNA duplexes (section
4.2), the glutaraldehyde cross-linking results suggested that the presence of ATP affected
RIG-I-RNA complex stability in the absence of a 5’-triphosphate. Upon the addition of
ATP, the protein dissociated from the thirteen and thirty-six base pair RNA duplexes lacking a 5’-triphosphate (figure 4.2, lanes 4 and 8). While the protein appeared to dissociate completely from the thirteen base pair duplex (compare lanes 3 and 4), the change seen in the thirty-six base pair duplex was the disappearance of the higher molecular weight complex formed (compare lanes 7 and 8). This result would be similar to MDA5, which exhibits enhanced dissociation from RNA in the presence of ATP
(Berke et al, 2012b). Taken together, the results from section 4.2 and this section suggest that while the presence of nucleotide has no significant effect on RIG-I RNA affinity, the presence of ATP does affect RIG-I-RNA complex stability and multimerization.
4.2.2 ATP accelerates dissociation of RIG-I from RNA
To further examine the cross-linking result that suggested ATP enhanced the dissociation of RIG-I from the 13 bp RNA duplex, I measured the dissociation rate constants of RIG-I from several of the RNA duplexes. To measure the dissociation rate constants, I formed RIG-I-RNA complexes by incubating excess RIG-I with a limiting
96 amount of radiolabeled RNA. Next, I added an excess of unlabeled trap RNA and
various nucleotides. Following the addition of the trap and nucleotide, samples were
taken over the course of five minutes and resolved with native PAGE (figure 4.4).
Dissociation of RIG-I-RNA complexes were plotted versus time, and fitted to a first order rate law to determine the dissociation rate constants for RIG-I in the presence of ATP,
ADPNP, and ADP. Additional controls were performed in the presence of Mg2+ alone
and UTP (figure 4.5 and table 4.2).
I measured the dissociation rate constants for wild-type RIG-I from the thirty-six
base pair duplexes with and without a 5’-triphosphate (table 4.2). In the absence of
nucleotide, the dissociation rate constant of RIG-I from the thirty-six base pair RNA
-1 duplex with a 5’-hydroxyl was kdiss = 0.14 ± 0.03 min . As a control, I measured the dissociation rate constant of RIG-I in the presence of excess magnesium ion, since all the reactions containing nucleotide also contain equimolar concentrations of magnesium ion.
The addition of magnesium chloride to the reaction slightly enhanced the dissociation
97 -1 rate constant, to kdiss = 0.44 ± 0.18 min . However, in the presence of ATP, the
-1 dissociation rate constant of RIG-I increased ten-fold to kdiss = 1.6 ± 0.4 min , indicating that like MDA5, wild-type RIG-I dissociation from RNA is enhanced in the presence of
ATP (figure 4.5) (Berke et al, 2012b). I performed an additional control in the presence of UTP, which is not utilized by RIG-I’s helicase/ATPase domain (Gee et al, 2008). In the presence of UTP, the dissociation of RIG-I from the complex was comparable to the
-1 dissociation of RIG-I with no nucleotide present (kdiss = 0.27 ± 0.09 min ). Furthermore,
I measured the dissociation rate constant of RIG-I in the presence of ADP and the non- hydrolyzable analog ADPNP, and found the neither of these nucleotides enhanced the
ADP -1 ADPNP dissociation rate constant like ATP (kdiss = 0.18 ± 0.06 min and kdiss = 0.40 ±
0.15 min-1) (table 4.2).
98 The presence of the 5’-triphosphate on the RNA duplex did not affect the ATP-
dependent dissociation of RIG-I. In the absence of nucleotide, the dissociation rate
-1 constant of RIG-I from the RNA was kdiss = 0.18 ± 0.06 min , and the addition of ATP to
-1 the reaction enhanced the dissociation rate constant to kdiss = 1.3 ± 0.6 min (table 4.2).
The dissociation rate constant of RIG-I from the thirty-six base pair duplex with a 5’- triphosphate was not enhanced by either magnesium chloride, UTP, ADP, or ADPNP, which is consistent with the results from the same duplex lacking a 5’-triphosphate.
Collectively, the results presented in this section imply that RIG-I dissociation from RNA is enhanced by ATP hydrolysis, since the nonhydrolyzable ATP analog ADPNP did not accelerate the dissociation rate constant to a similar degree (table 4.1).
99 4.2.3 RIG-I RNA-dependent ATPase activity is independent of RNA duplex length
RIG-I-RNA dependent ATPase activity plays a crucial but undefined role in the
activation of antiviral signaling (Saito et al, 2007). Also, a correlation exists between a
substrate’s ability to activate antiviral signaling in vivo and its ability to stimulate RIG-I
ATPase activity (Schlee et al, 2009 and Schmidt et al, 2009). A similar correlation is not present for RNA binding, and the link between RNA recognition and ATP utilization remain unclear. To probe this question, I examined the stimulation of RIG-I RNA- dependent ATPase activity with a series of blunt RNA duplexes ranging from ten to thirty-six base pairs. The reactions were performed under saturating RIG-I and RNA conditions (figure 4.6A). γ-32P-ATP and free inorganic phosphate were resolved with
thin layer chromatography, and the amount of inorganic phosphate increased over time as
RIG-I hydrolyzed the γ-32P-ATP. The fraction hydrolyzed was quantified with
densitometry, and the velocity was determined under various ATP concentrations. The
velocity was plotted versus ATP concentration, and kinetic parameters were determined
from the Michaelis-Menten equation (figure 4.6B).
100
The results indicate that the only requirement for RIG-I ATPase stimulation appears to be double-stranded RNA. Both the catalytic efficiency (kcat/Km) and the affinity of RIG-I for ATP are similar for all RNA duplexes tested (figure 4.6). For a blunt RNA duplex, the Km for ATP is about 250 µM, suggesting that when bound to viral
RNA, under physiological conditions RIG-I will be saturated with ATP. Furthermore, no significant change of RIG-I ATPase activity is seen for RNAs with and without a 5’- triphosphate (figure 4.7). This implies that despite the critical role of the 5’-triphosphate
101 in the activation of signal transduction in vivo, the 5’-triphosphate is dispensable for the
stimulation of the RNA-dependent ATPase activity.
4.3 Discussion
4.3.1 RIG-I ATP hydrolysis serves a signal for the identification of dsRNA
In section 4.2.1, I measured the affinity of RIG-I for the thirty-six base pair
duplex with a 5’-triphosphate in the presence of various nucleotides representing the
different intermediates of the ATP hydrolysis cycle. Under equilibrium conditions, I
found that the presence of nucleotide did not affect the overall affinity of RIG-I for
dsRNA, but when fit to the model for a coupled equilibrium the presence of the transition
state analog ADP-AlFx weakened the affinity of the formation of the ES complex (table
4.1). These findings confirm that ATP or other nucleotide is unnecessary for RIG-I RNA
102 binding to occur and suggest that the formation of the ES complex weakens when ADP-
Pi is bound by RIG-I during the ATP hydrolysis cycle.
In order to account for the limitations of measuring RNA binding with nucleotide
under equilibrium conditions, I analyzed chemically crosslinked RIG-I-RNA complexes
in the presence of ATP to examine if nucleotide had any effect on multimerization. This
data demonstrated that in the presence of ATP, RIG-I dissociated from the RNA (figure
4.2). I followed this further by measuring the dissociation rate constants of RIG-I
binding to the thirty-six base pair duplexes with and without a 5’-triphosphate, which
showed that the presence of ATP in the reaction enhances the dissociation rate constant
by over ten-fold (figure 4.5 and table 4.2).
I next measured RIG-I’s RNA-dependent ATPase activity with a series of model
substrates to determine the contributions of RNA duplex length and the presence of a 5’-
triphosphate. In chapter 3, I showed that RIG-I ATPase activity is only present when a
double-stranded RNA substrate is used. I further investigated this is section 4.2.4, and
measured ATPase stimulation of RIG-I for substrates ranging from ten to thirty-six base
pairs. The results showed that RIG-I ATPase activity is independent of RNA duplex
length (figure 4.7). This finding is consistent with the work of Marq et al (2010a), who demonstrated that a minimum of seven continuous base pairs was necessary for
Arenavirus RNA to stimulate RIG-I ATPase activity. It also provides further support for
the role of RNA secondary structure in RIG-I PAMP recognition.
I also compared the ATPase stimulation of RIG-I by a substrate with a 5’-
hydroxyl to a substrate with a 5’-triphosphate. Schlee et al (2009) demonstrated that the
103 presence of a 5’-triphosphate significantly enhanced RIG-I ATPase activity, while
Kohlway et al (2013) saw no significant change in ATP turnover in when comparing the
same RNA duplex both with and without a 5’-triphosphate. My results agree with those
of Kohlway et al (2013) and show that the presence of the 5’-triphosphate is dispensable
for the activation of RIG-I RNA-dependent ATPase activity. While the 5’-triphosphate is
clearly important for the activation of signaling in vivo, in vitro the 5’-triphosphate has
little effect on RIG-I RNA binding or ATPase stimulation.
The single order of magnitude increase in the dissociation rate constant in the
presence of ATP indicates that the association rate constant will increase to a similar
RNA degree. The association rate constant can be estimated by calculating kcat/K1/2 . The
9 -1 -1 calculated association rate constant is kass~ 10 M s . This value is approaching the diffusion limit, suggesting that in the presence of ATP RIG-I binds dsRNA at nearly every encounter of the two molecules. Comparison of the dissociation rate constant and rate of ATP turnover for RIG-I reveal that in the presence of ATP, RIG-I hydrolyzes about 400 molecules of ATP during each encounter with a molecule of dsRNA.
Collectively, these values and the observation that RIG-I RNA-dependent ATPase activity is independent of RNA duplex length and presence of a 5’-triphosphate suggest that ATPase activity is functioning to recognize the presence of dsRNA.
Combined with the structural evidence showing RIG-I bound to a blunt duplex
end (Luo et al, 2011, Kowalinski et al 2011, and Jiang et al, 2011) and evidence that a
RIG-I monomer is sufficient for RIG-I ATPase stimulation (Luo et al, 2011), my findings suggest a model where RIG-I ATPase activity is “proofreading” RNA to locate a blunt duplex end. Upon identification of a blunt duplex end, RIG-I can adopt an ATP
104 hydrolysis competent conformation that exposes the CARD domains (Luo et al, 2011 and
Kowalinski et al 2011). The acceleration of the dissociation rate constant and concomitant increase in the association rate constant in the presence of ATP indicate that upon binding an activating substrate, ATP hydrolysis increases the amount of opportunities for RIG-I to undergo this conformational change. This would allow for the maximum number of opportunities for the exposed CARD domains to interact with downstream adaptor proteins, ubiquitination enzymes, and free ubiquitin chains. These interactions and posttranslational modifications may be involved in stabilizing a signaling competent conformation of RIG-I in order to activate or potentiate downstream antiviral signaling pathways.
105 Chapter 5
RIG-I proofreads RNA duplexes for blunt ends with its ATPase activity
5.1 Introduction
The crystal structures of full-length RIG-I bound to dsRNA suggested that RIG-I
requires a blunt duplex end for efficient binding (Jiang et al, 2011, Luo et al, 2011, and
Kowalinski et al, 2011). Furthermore, the presence of ssRNA overhangs on an RNA
duplex have been shown to be sufficient to prevent the activation of RIG-I antiviral
signaling in vivo (Schlee et al, 2009, Schmidt et al, 2009, Marq et al, 2010a, and Marq et al, 2010b). To investigate the role of an exposed blunt duplex end on RIG-I activity in vitro, I examined the RNA binding and RNA-dependent ATPase stimulation of RIG-I with a series of model substrates containing ssRNA overhangs that blocked either one or both duplex ends (figure 5.1).
106 5.2 Results
5.2.1 RIG-I efficiently binds RNA duplexes with blocked ends
The series of end-blocked RNA duplex substrates were designed using the sequence for the nineteen base pair blunt duplex RIG-I binds this RNA with an affinity of K1/2 = 15.9 ± 3.6 nM (Section 3.2.4). I next examined RIG-I binding to nineteen base pair duplexes containing either a single 5’- or 3’-overhang. The published structures of
RNA-bound RIG-I suggested that RIG-I may tolerate a 3’-overhang better than a 5’- overhang, since the CTD of RIG-I caps the 5’-strand of the duplex (Jiang et al, 2011,
Kowalinski et al, 2011, and Luo et al, 2011). RIG-I bound both of these duplexes with a similar affinity to that of a blunt nineteen base pair duplex (figure 5.2). The lack of effect mediated by either the 5’- or 3’-overhangs may be due to the available blunt end of the other terminus on these duplexes.
107 I next investigated if fully blocking one blunt end of the RNA affected the affinity of RIG-I for the RNA. I performed gel shift experiments using a duplex containing one blunt end and one end blocked with a 5’- and a 3’-overhang. The affinity of RIG-I for a nineteen base pair RNA duplex with one end blocked by ssRNA overhangs was K1/2 =
10.3 ± 0.2 nM. The data indicates that RIG-I’s RNA affinity was unaffected by the
availability of only one duplex blunt end (figure 5.3).
I next examined RIG-I RNA binding to substrates with three or four ssRNA
overhangs. The substrates with three ssRNA overhangs have one end completely
blocked with both a 5’- and a 3’-overhang, while the other end has either a 5’- or a 3’-
overhang. The substrate with four overhangs has both 5’- and 3’-overhangs on both ends
of the duplex (figure 5.1). Surprisingly, despite both ends of these duplexes being
blocked with ssRNA overhangs, RIG-I’s affinity for the nineteen base pair duplexes
108 containing three ssRNA overhangs was comparable to that of the nineteen base pair blunt
duplex (table 5.1). RIG-I binding to the duplex with four ssRNA overhangs, in which
both duplex ends are completely blocked, was decreased only slightly compared to the nineteen base pair blunt duplex (figure 5.4). Collectively, these data indicate that RIG-I is still capable of binding RNA duplexes without an exposed blunt end, despite structural evidence implicating a blunt RNA end is necessary for effective RIG-I RNA binding.
5.2.2 RIG-I ATPase activity proofreads RNA duplexes for blunt ends
RNA binding is only the first step involved in RIG-I RNA recognition.
Previously I demonstrated that RIG-I RNA-dependent ATPase activity requires dsRNA,
and is independent of RNA duplex length. However, some evidence indicates that the
presence of ssRNA overhangs on RNA duplexes decreases RIG-I ATPase activity
(Schlee et al, 2009, Schmidt et al, 2009, Marq et al, 2010a, and Marq et al, 2010b). I next
109 investigated if blocking the RNA duplex ends would have an impact on RIG-I RNA- dependent ATPase activity. I measured the stimulation of RIG-I ATPase activity in the presence of the series of end-blocked model substrates described in the previous section
(figure 5.5)
The results are summarized in table 5.1, and demonstrate that the presence of a blunt RNA duplex end is absolutely necessary for the significant stimulation of RIG-I
RNA-dependent ATPase activity. ATP hydrolysis by RIG-I was unaffected by the blocking of one duplex end with 5’- and 3’-ssRNA overhangs compared to a nineteen base pair blunt duplex. However, despite the evidence indicating that RIG-I tightly binds
RNA duplexes with both ends blocked by ssRNA overhangs, this binding is insufficient for the stimulation of RNA-dependent ATPase activity. The kcat of RIG-I in the presence
110 of both substrates with three ssRNA overhangs was decreased by over two orders of magnitude compared to either the nineteen base pair blunt RNA duplex or the duplex with one end blocked (table 5.1). An even more significant effect was observed for the duplex with both ends completely blocked (four ssRNA overhangs). RIG-I’s kcat was decreased by over four orders of magnitude compared to that of a duplex with at least one available blunt end (table 5.1).
111 5.3 Discussion
5.3.1 RIG-I RNA-dependent ATPase activity proofread duplexes for blunt ends
These results demonstrate a role for RIG-I RNA-dependent ATPase activity in
proofreading RNA duplexes for blunt ends. RIG-I is still capable of binding RNA
duplexes lacking an available blunt end, but this binding is insufficient for the stimulation
of RNA-dependent ATPase activity (table 5.1). This result is consistent with published
results from Marq et al, who demonstrated that Arenavirus and Bunyavirus genomes
contain an unpaired 5’-triphosphate nucleotide that allows these viruses to evade
detection of RIG-I (2010a and 2010b). RNAs produced by these viruses are unable to
activate RIG-I antiviral signaling in vivo, and show reduced stimulation of RIG-I RNA- dependent ATPase activity in vitro.
The data presented in this chapter suggest a model of RIG-I activation in which
RIG-I binds to RNA duplexes with no regard for the 5’-end structure. However, the small decrease in affinity observed for RIG-I binding to an internal duplex region compared to a duplex end is unlikely to be enough to allow any significant discrimination between the two types of binding sites in vivo, suggesting that RIG-I needs additional means by which to distinguish between self and non-self RNA. The structures of RNA- bound RIG-I demonstrated that RIG-I only adopts an ATP hydrolysis competent conformation upon binding to a dsRNA (Luo et al, 2011, Kowalinski et al, 2011, and
Jiang et al, 2011). While RIG-I is capable of binding to internal duplex regions with high affinity (section 5.1), the lack of an exposed 5’-terminus for the CTD to interact with
112 likely results in the protein adopting a different conformation in which the two RecA-like
domains are not oriented properly to form a functional ATPase site.
In order for signaling activation to occur, RIG-I ATPase activity must be stimulated, and only responds to the presence of a blunt duplex end. This finding is consistent with the structures of RNA-bound RIG-I by Luo et al (2011), Kowalinski et al
(2011), and Jiang et al (2011) and with the work of Kohlway et al (2013), who demonstrated that an RNA hairpin with a blunt duplex end was sufficient for the stimulation of RIG-I ATPase activity and for the activation of antiviral signaling. The results in section 5.2 demonstrate clearly that the function of RIG-I’s ATPase activity is to proofread RNA duplexes to identify exposed blunt ends. This proofreading is necessary for the activation of RIG-I antiviral signaling, and is responsible for detecting one of the features of the RIG-I PAMP.
5.3.2 Model of RIG-I substrate recognition
Collectively, the data presented in this thesis provide further evidence for a model of RIG-I RNA recognition in which the protein is searching for multiple signals within the RNA to discriminate between cellular RNA and viral RNA. The high affinity RIG-I exhibits for dsRNA described in chapter 3 is necessary for it to bind to the rare molecules of viral RNA present in the early stages of a viral infection. RIG-I must discriminate between the vast amounts of cellular RNA (e.g. ribosomal, transfer, messenger, etc.) and the relatively few molecules of viral RNA. RIG-I is capable of binding to both blunt duplex ends and internal duplex regions, suggesting that RIG-I would likely be bound to the duplex regions of cellular RNAs. Since cellular RNAs would lack additional features
113 identified by RIG-I, such as an exposed blunt end or 5’-triphosphate, RIG-I binding to
these RNAs would not result in activation of the antiviral signaling pathway. The
requirement for RIG-I to recognize multiple features of viral RNA before activating the
signaling pathway underscores the critical need for strict regulation of RIG-I activity.
Inappropriate activation of antiviral signaling pathways would be deleterious for cells
(Akira et al, 2006).
Previous models of RIG-I RNA recognition and activation of antiviral signaling
have proposed that in an uninfected cell, RIG-I exists in an auto-inhibited conformation
that prevents interaction with cellular RNA. Based on the data I have presented in this
thesis, I propose a model for RIG-I RNA recognition where in a healthy cell, RIG-I is
constantly “sampling” RNA, binding to and dissociating from cellular RNA while
proofreading the RNA attempting to identify the features of viral RNAs. This is
suggested by the high affinity of RIG-I for dsRNA in the absence of ATP and the
abundance of RNA in cells. Upon viral infection, RIG-I binds to the viral RNA and in
the course of proofreading the RNA identifies the features of viral RNA (e.g. dsRNA, an
exposed blunt duplex end, 5’-triphosphate, sequence elements). In particular, RIG-I
utilizes its RNA-dependent ATPase activity to identify dsRNA, and only upon binding to
dsRNA and adopting the proper conformation can RIG-I hydrolyze ATP (chapter 4 and
Luo et al, 2011, Kowalinski et al, 2011, and Jiang et al, 2011).
In chapter 5 I identified a blunt duplex end as a critical RNA feature necessary for
RIG-I RNA-dependent ATPase activity. RIG-I undergoes a conformational change upon
binding to a blunt duplex end which positions the two RecA-like domains in an ATP hydrolysis competent conformation (Luo et al, 2011, Kowalinski et al, 2011, and Jiang et
114 al, 2011). The affinity of RIG-I for ATP increases dramatically in the presence of a blunt
duplex end (chapters 4 and 5 and Luo et al, 2011). This finding in combination with the
RNA-bound structures of RIG-I (Luo et al, 2011, Kowalinski et al, 2011, and Jiang et al,
2011) provides strong evidence for the role of RIG-I RNA-dependent ATPase activity in
RIG-I substrate recognition. Only upon binding to a blunt duplex end can RIG-I adopt the conformation necessary for ATP hydrolysis, which Saito et al (2007) demonstrated was necessary for signal transduction in vivo.
The conformational change that results in ATP hydrolysis competent RIG-I also ejects the CARD domains and exposes them to the cytosol (Luo et al, 2011 and
Kowalinski et al, 2011). The structure of ligand free RIG-I demonstrated that the CARD
domains are not accessible due to an interaction with the helicase domain (Kowalinski et
al, 2011). Once exposed to the cytosol, the CARD domains are able to interact with the
downstream adaptor protein MAVS to potentiate downstream signaling pathways, and
are available for posttranslational modification and interaction with free ubiquitin chains
that have been shown to be necessary for RIG-I signaling in vivo (Kowalinski et al, 2011 and Kohlway et al, 2013).
115 Chapter 6
Future Directions
6.1 Introduction
In this thesis, I described my findings regarding the roles of RIG-I RNA binding,
ATPase activation, and multimerization in substrate recognition. I determined that both
RNA binding and ATPase stimulation require an RNA duplex, and that they seem to be independent of RNA duplex length, while RIG-I multimerization is dependent on RNA duplex length. Intact RNA-dependent ATPase activity has been shown to be necessary for the activation of antiviral signaling in vivo (Saito et al, 2007). In chapter 5, I demonstrated that RNAs lacking an available blunt duplex end exhibited a significantly reduced ability to stimulate RIG-I ATPase activity.
Collectively, the data presented in this thesis suggest a model of where RIG-I
RNA recognition depends on the ability of RIG-I to proofread an RNA duplex in search
of an exposed blunt end. RNA binding by RIG-I is the first step of substrate recognition,
and the findings in chapter 3 and chapter 5 suggest that in cells RIG-I is likely interacting with cellular RNA, sampling any dsRNA it encounters. When RIG-I does bind to a viral
RNA, it adopts an ATP hydrolysis competent conformation. This conformational change exposes the CARD domains, allowing for the induction of antiviral signaling. In this chapter, I will present possible future investigations to extend the understanding of the mechanism of viral RNA recognition by RIG-I in cells.
116 6.2 Effects of RNA 5’-end structure on RIG-I RNA recognition in vivo
Multiple lines of evidence suggest that the 5’-end structure of an RNA duplex plays a role in the recognition of viral RNA by RIG-I. The data I presented in chapter 3 suggests that the 5’-triphosphate has no significant impact on RIG-I RNA binding, which is consistent with some published results and inconsistent with others (Pichlmair et al,
2006, Hornung et al, 2006, Cui et al, 2008, Schlee et al, 2009, Schmidt et al, 2009, Vela et al, 2012, and Kohlway et al, 2013). Overall, the literature suggests that despite the inconsistencies seen in vitro, in vivo RIG-I requires the presence of a base-paired 5’- triphosphate to activate antiviral signaling (Pichlmair et al, 2006, Hornung et al, 2006,
Cui et al, 2008, Schlee et al, 2009, Schmidt et al, 2009, and Marq et al, 2010b). Some groups have investigated if variations in the 5’-end structure of the RNA had any significant effect on stimulation of RIG-I reporter genes. Hornung et al (2006) found that
RNA with either a monophosphate or a diphosphate was incapable of inducing interferon levels to those seen in the presence of a 5’-triphosphate, while Pichlmair et al (2006) found that the presence of only one phosphate group on influenza virus RNA was necessary for the induction of RIG-I-dependent signaling. Furthermore, additional work has suggested that RIG-I will not recognize a 5’-monophosphorylated RNA, and that some viruses avoid detection by RIG-I by modifying their 5’-ends from 5’-triphosphates to 5’-monophosphates (Schlee et al, 2009, Schmidt et al, 2009, and Habjan et al, 2008).
In chaper 2, I described a novel method to generate 5’-triphosphate 5’-end labeled
RNAs to use as RIG-I substrates for in vitro studies. I further showed that in addition to
GTP, the ribozyme utilized other guanosine analogs to efficiently cleave the precursor substrates. This method could be used to generate RIG-I substrates with additional 5’-
117 end structures. RNAs containing a 5’-monophosphate, a 5’-disphosphate, and 5’-
imidophosphate could be easily produced. The guanosine analog 5’-guanylyl-
imidophosphate has an oxygen atom between the γ- and β-phosphate groups replaced
with an amine, rendering this analog unhydrolyzable.
RNAs with varying 5’-end structures, including the ones described above, could
be transfected into cell lines to examine the effect of 5’-end structure on the activation of
antiviral signaling. This would provide additional evidence for the role of the 5’-end structure on RIG-I RNA recognition. Additionally, the mechanism by which members of
the Bunyavirus family evade RIG-I detection could possibly be elucidated by using
RNAs with a 5’-imidophosphate. If the virus is using a phosphatase to process the
genome 5’-termini, then the transfection of RNA with a 5’-imidophosphate would be
resistant to dephosphorylation and would still be able to activate RIG-I-dependent
antiviral signaling. This would demonstrate the molecular mechanism of RIG-I evasion
employed by Bunyaviruses.
6.3 Activation of antiviral signaling in vivo by frayed end substrates
In addition to the 5’-end structure, the induction of RIG-I signaling in vivo is also
dependent upon the presence of a blunt duplex end (Schlee et al, 2009, Schmidt et al,
2009, Marq et al, 2010b). This finding is supported by the structural data, which shows
RIG-I’s helicase domain wrapping around the ten terminal base pairs with the CTD capping the 5’-strand of the RNA (Jiang et al, 2011, Luo et al, 2011, and Kowalinski et al, 2011). In chapter 5, I demonstrated that despite efficient binding to an RNA duplex
118 with both ends blocked with ssRNA overhangs, RIG-I ATPase stimulation was significantly diminished. Since RIG-I ATPase activity is critical for the activation of signaling in vivo, I would use the substrates described in chapter 5 to investigate directly how a lack of ATPase stimulation effects the activation of RIG-I signaling (Saito et al,
2007).
Previous groups demonstrated that even a single nucleotide RNA overhang had deleterious effects on the activation of antiviral signaling, and have suggested the use of short RNA overhangs as a mechanism of RIG-I avoidance by viruses (Schlee et al, 2009,
Schmidt et al, 2009, and Marq et al, 2010b). I would expect that the substrates with both ends blocked by ssRNA overhangs to be incapable of activating RIG-I signaling in vivo.
This set of experiments would directly demonstrate that the ATP hydrolysis dependent on
RIG-I binding to an available blunt duplex end is necessary for signaling activation in vivo.
6.3 Effect of ubiquitination of RIG-I on RIG-I RNA binding and ATP hydrolysis
Ubiquitination of the second RIG-I CARD domain by several E3 ubiquitin ligases has been shown to be critical for the activation of antiviral signaling in vivo (Gack et al,
2007, Oshiumi et al, 2009, Gao et al, 2009, and Oshiumi et al, 2010). Additionally, Zeng et al (2010) demonstrated that RIG-I signaling depended upon the interaction of the
CARD domains with free polyubiquitin chains. The structures of RNA bound RIG-I generated by Luo et al (2011), Kowalinski et al (2011), and Jiang et al (2011) demonstrated that in order to relieve the steric hindrance on the ubiquitination site in the
119 second CARD domain, RIG-I must be RNA bound. Collectively, these findings from the
literature suggest a model where one function of the conformational change that results from RNA binding is to exposes the CARD domains and allow access for the ubiquitination machinery and polyubiquitin chains. The ubiquitination of or interaction with free polyubiquitin by RIG-I stabilizes the active conformation, making RNA binding at that point unnecessary to continue signaling.
To further investigate the validity of the above model, I propose to measure the
RNA binding and ATPase activity of ubiquitinated RIG-I and RIG-I in the presence of polyubiquitin chains. Ubiquitinated RIG-I would be produced by purifying RIG-I from
E. coli as described in section 7.1.2, in addition to purifying the E3 ubiquitin ligases identified to ubiquitinate RIG-I, such as TRIM25 or Riplet. The remainder of the proteins required for in vitro ubiquitination would be purchased. Large scale ubiquitination reactions would be performed from which ubiquitinated RIG-I (RIG-I-Ub) would be purified.
Next, the RNA binding and RNA-dependent ATPase activity of RIG-I-Ub could be determined and compared with that of unmodified RIG-I. To examine the effect of polyubiquitin chains, the RNA binding and ATPase assays would simply be performed in the presence of polyubiquitin. Since a function of RIG-I RNA binding is to relieve the steric hindrance on the CARD domains and facilitate an open and active protein conformation, I anticipate that RIG-I RNA binding may be negatively affected by either the ubiquitination of RIG-I or its interaction with free ubiquitin chains. If RNA binding is diminished, then an accompanying decrease or abolishment of RNA-dependent
ATPase activity would also likely occur.
120 The functions of several different ubiquitination sites in RIG-I could also be ascertained with this approach. TRIM25 has been shown to ubiquitinate K172 in the second RIG-I CARD domain, while Riplet has been shown to ubiquitinate K172 and several lysine residues in the CTD (K849, K851, K888, K901, and K907). According to the model proposed above, ubiquitination of the CARD domain has a likely role in stabilizing the active conformation of RIG-I and a point mutant deficient in K172 ubiquitination would more than likely retain wild-type levels of RNA binding and
ATPase stimulation.
Meanwhile the role of the ubiquitination sites in the CTD remain unknown. One of the putative ubiquitination sites in the CTD, K888, is present in the positively charged groove that serves as an RNA binding site in the CTD and mutation of this residue by Cui et al (2008) diminished both RNA binding and interferon-β/firefly luciferase reporter gene activity (figure 6.1). Several other residues present in the positively charged groove also exhibited reduced RNA binding and reporter gene stimulation (Cui et al, 2008). The ubiquitination sites in the RIG-I CTD are positioned around the edges of (K849, K851,
K901, and K907) and directly in (K888) the positively charged groove, suggesting that when covalently modified with K63-linked ubiquitin the RNA may not have access to the binding site due to the bulky posttranslational modifications.
121
This hypothesis could be demonstrated by preparing ubiquitinated RIG-I, both wild-type and point mutants of these residues, particularly K888, and testing their ability to bind RNA in vitro. Data presented in chapter 3 of this thesis and that of many groups in the literature (Cui et al, 2008, Luo et al, 2011, Jiang et al, 2011, Vela et al, 2012, and
Kohlway et al, 2013) demonstrate that ubiquitination of RIG-I is not necessary for RNA binding. If wild-type RIG-I-Ub exhibits diminished or abolished RNA binding activity while point mutants with diminished ubiquitination retain RNA binding activity, this would suggest that the function of ubiquitination of the RIG-I CTD is to prevent RNA binding following stabilization of the protein’s active conformation. This could also
122 function to amplify the signaling response by preventing activated RIG-I that is locked into a stable signaling competent conformation by ubiquitin from binding viral RNA.
123 Chapter 7
Materials and Methods
7.1Materials
7.1.1 Plasmids
Plasmids expressing WT RIG-I, K270A RIG-I, RIG-IΔCARD, RIG-IΔRD, and
RIG-IΔCARDΔRD in the pET-28-His10-Smt3 vector were a gift from Dr. Dominique
Garcin. The cloning of these constructs is described in (Hausmann et al, 2008). To generate the plasmids R36C-pUC18, R41-3-pUC-18, R36-25-pUC18, R62-L21-pUC18, and R68-TRAP-L21-pUC18, synthetic DNA oligonucleotides encoding the sense strand and the antisense strand of each oligo were purchased from Sigma Aldrich (Table 7.1).
DNA oligonucleotides were annealed by incubation for four minutes at 95°C, followed by slow cooling to room temperature. DNA templates were ligated into pUC18 following digestion of the vector with BamHI and EcoRI. Transformants containing the desired inserts were identified with DNA sequence analysis. The plasmid encoding the L21-ScaI ribozyme (L21-ScaI-pUC19) was a gift from Dr. Mark Caprara (Caprara and Waring,
1994).
124
DNA Oligo Sequence Restriction RNA Name Enzyme to Transcript Linearize Produced R36C 5’GGATCCTAATACGACTCACTATAGGCACCG NaeI R36C Coding TAAAGACC CAATCATGCAGGGTCTGTGCCGGCGAATTC 3’ R36C 5’GAATCGCCGGCACAGACCCTGCATGATTG NaeI R36C Noncoding GGTCTTTACGGTG CCTTTAGTGAGTCGTATTAGGATCC 3’ R41-3 5’GGATCCTAATACGACTCACTATAGGGTCTT SnaBI R41-3 Coding TACGGTGCCCAGCATCAATGACATCAGCATC TACGTA GAATTC 3’
R41-3 5’GAATTCTACGTAGATGCTGATGTCATTGAT SnaBI R41-3 Noncoding GCTGGGCACCGTAAAGACCCTATAGTGAGTC GTATTAGGATCC 3’ R36-25 5’GGATCCTAATACGACTCACTATAGGCACAG SnaBI R36-25 Coding ACCCTGCATGATTGGGTCTTTACGGTGCCCA GCATCAATGACATCAGCATCTACGTAGAATT C 3’ R36-25 5’GAATTCTACGTAGATGCTGATGTCAATTGA SnaBI R36-25 Noncoding TGCTGGGCACGTAAAGACCCAATCATGCAGG CGTTGTGCCTATAGTGAGTCGTATTAGGATC C 3’
R62-L21 5’GGATCCTAATACGACTCACTATAGGGAGA SnaBI R62-L21 Coding CAAAGTATAGCCCTCTAACACAAACGCGTCA TGATTGACACTTTACGGTGCCCAGCATCAAT GACATCAGCATCTACGTAGAATTC 3’
R62-L21 5’GAATTCTACGTAGATGCTGATGTCATTGAT SnaBI R62-L21 Noncoding GCTGGGCACCGTAAAGTGTCAATCATGACGC GTTTGTGTTAGAGGGCTATACTTTGTCTCCCT ATAGTGAGTCGTATTAGGATCC 3’ R68- 5’GGATCCTAATACGACTCACTATAGGGACGC SnaBI R68-TRAP TRAP- CCTCTAGTAAATTTATATAAGATAAGATAAG L21Coding ATAAGATAAGATAAGATAAGATAAGATAAG ATACCCGAAGATCCTACGTAGAATTC 3’ R68- 5’GAATTCTACGTAGGATCTTCGGGTATCTTA SnaBI R68-TRAP TRAP-L21 TCTTATCTTATCTTATCTTATCTTATCTTATCT Noncoding TATCTTATATAAATTTACTAGAGGGCGTATA TACTTTGTCTCCCTATAGTGAGTCGTATTAGG ATCC 3’ Table 7.1. List of DNA Oligos for Transcription Templates
125 7.1.2 Proteins
Recombinant wild-type and mutant RIG-I proteins were purified from Escherichia
coli strains expressing wild-type and mutant RIG-I. BL21(DE3) E. coli strains expressing pET28-SMT-His10-RIG-I were grown to an OD600 of 0.6 to 0.8. Protein
expression was induced by the addition of 0.2M IPTG, and 2% ethanol was also added to
the culture at this time to induce heat shock protein expression and increase the amount
of RIG-I present in the soluble fraction (Hausmann et al, 2008, Strandberg and Enfors,
1991, and Thomas and Baneyx, 1996). Cultures were then incubated for sixteen hours at
16°C. Cells were pelleted, and lysed in ice cold lysis buffer containing 50 mM NaH2PO4, pH 7.4, 300 mM NaCl; 25% glycerol, 5 mM imidazole, and Roche Complete Mini Tablet protease inhibitors (buffer A). Clarified lysates were incubated with nickel-sepharose
(Qiagen Ni-NTA resin) for thirty minutes at 4°C. The sample was applied to a gravity flow column and washed sequentially with buffer A containing 5 mM imidazole, 30 mM imidazole, and 60 mM imidazole. RIG-I was eluted off the column with buffer A containing 500 mM imidazole. Fractions containing RIG-I protein were identified via analysis of flowthrough, all washes, and all elution fractions with SDS-PAGE.
RIG-I containing fractions were pooled, and the total salt concentration was reduced to 25 mM by dilution with buffer containing 50 mM Tris, pH 8.0, 1 mM EDTA, 2 mM
DTT, 25% glycerol, and 0.1% Triton X-100 (buffer B). Diluted fractions were applied to a phosphocellulose column equilibrated in buffer B. The column was washed with
Buffer B containing 50 mM and 100 mM NaCl, and RIG-I was eluted with buffer B containing 500 mM NaCl. RIG-I containing fractions were identified via SDS-PAGE
126 analysis. Total protein concentration was determined with the Bradford assay. Aliquots were flash frozen and stored at -80°C.
7.1.3 Oligonucleotides
RNA oligonucleotides were purchased from Dharmacon and Sigma Aldrich, and
DNA oligonucleotides were purchased from Sigma Aldrich. RNAs purchased from
Sigma Aldrich were further purified with denaturing PAGE. RNAs were visualized in the gel with UV shadowing, and excised for elution. The sequence and substrate design of all RNA oligonucleotides are shown in Table 7.2
RNA/DNA RNA/DNA Sequence Base Pairing Special Chapters Purchased Name Features Used From R10B 5’ UUACGGUGCU 3’ With R10C, R16C, and 3, 4 Dharmacon R19C R10C 5’ AGCACCGUAA 3’ With R10B, R16B, and 3, 4 Dharmacon R19B R10L21P 5’ CCCUCUAGCA L21-ScaI 2 Sigma- CCGUA 3’ cleavage Aldrich site R10L21Bot 5’ UACGGUGCUC 3’ With R10L21C and 2, 3 Sigma- pppR10L21 Aldrich R10L21C 5’ GAGCACCGUA 3’ With R10L21Bot 3 Sigma- Aldrich R13L21P 5’ CCCUCUAGCACC L21-ScaI 2 Sigma- GUAAAG 3’ cleavage Aldrich site R13L21B 5’ CUUUACGGU With R13L21C and 2, 3 Sigma- GCUC 3’ pppR13L21 Aldrich R13L21C 5’ GAGCACCGU With R13L21B 2, 3, 4 Sigma- AAAG 3’ Aldrich R13L21PBot2 5’ CCCUCUACU L21-ScaI 2 Sigma- UUACGGUGC 3’ cleavage Aldrich site R13L21D 5’ GACUUUACG With R13L21DBot 2, 3, 4 Sigma- GUGC 3’ Aldrich R13L21DBot 5’ GCACCGUAA With R13L21D and 2, 3, 4 Sigma- AGUC 3’ pppR13L21Bot Aldrich
127 RNA/DNA RNA/DNA Sequence Base Pairing Special Chapters Purchased Name Features Used From R16B 5’ GCGUCUUUA Same as R10B 3, 4 Dharmacon CGGUGCU 3’ R16C 5’ AGCACCGUAA Same as R10C 3, 4 Dharmacon AGACGC 3’ R19B 5’ GCUGCGUCU Same as R10B 3, 4, 5 Dharmacon UUACGGUGCU 3’ R19C 5’ AGCACCGUAA Same as R10C 3, 4, 5 Dharmacon AGACGCAGC 3’ R19B-UTR 5’ GUCUCCUCG With R19C-UTR, 3, 4, 5 Dharmacon UCUCCCUCGU 3’ R44C-5-UTR, R39C R19C-UTR 5’ ACGAGGGAG With R19B-UTR, R44- 3, 4, 5 Dharmacon ACGAGGAGAC 3’ 3-UTR, R39B R44-3-UTR 5’ GUCUCCUCU Same as R19B-UTR 5 Dharmacon CCCUCGUCAGCA UCAAUGACAUCA GCAUCAAA 3’ R44C-5-UTR 5’ AAACUACGA Same as R19C-UTR 5 Dharmacon CUACAGUAACUA CGACACGAGGGA GACGAGGAGAC 3’ R39B 5’ UAACUACGA Same as R19C-UTR 5 Sigma- CACGAGGGAGAC Aldrich GAGGAGACCAGC AUCAAU 3’ R39C 5’ UAACUACGA Same as R19B-UTR 5 Sigma- CGUCUCCUCGUC Aldrich UCCCUCGUCAGC AUCAAU 3’ R36D 5’ GGCACCGUA With R10B, R16B, 3, 4 Dharmacon AAGACGCAAUCA R19B, R36DBot, and UGCAGGGUCUGU D36DBot CAG 3’
R36DBot 5’ GGCACAGAC With R10C, R16C, 3, 4 Dharmacon CCUGCAUGAUUG R19C, R36D, and GGUCUUUACGGU D36D GCC 3’ D36D 5’ AGCACCGTA Same as R36D 3 Sigma- AAGACGCAATCA Aldrich TGCAGGGTCTGT CAG 3’ D36DBot 5’ GGCACAGAC Same as R36DBot 3 Sigma- CCTGCATGATTG Aldrich GGTCTTTACGGT GCC 3’ Table 7.2. List of RNA and DNA Oligos
128 5’-triphosphate containing RNA oligonucleotides and the L21-ScaI ribozyme
were prepared by in vitro transcription (Table 7.3). The DNA templates were prepared from maxiprep DNA (Qiagen Plasmid Maxi Kit). Plasmid DNA was linearized by restriction enzyme digestion with an enzyme producing blunt ends (Table 7.1). The linearized templates were purified by phenol chloroform extraction and ethanol precipitation. Linearization of the plasmids was confirmed via agarose gel analysis.
Linearized templates were quantified by measuring the absorbance at OD260, and the in vitro transcription reaction conditions were as described by the manufacturer
(MegaShortScript Kit, Ambion). RIG-I RNA substrate transcripts were gel purified with denaturing PAGE. L21-ScaI ribozyme transcription reactions were incubated with
DNase to remove the template DNA, and then purified via phenol chloroform extraction and ethanol precipitation (Ambion). The concentration of all RNAs was determined by measuring absorbance at OD260 (Eq. 7.1).
C = A/(ε * l) (Equation 7.1) C = concentration (µg/mL)
A = absorbance at OD260 ε = extinction coefficient of RNA (1/50 µg/mL for dsRNA) l = path length (cm)
129
RNA Transcript Sequence Base Pairing Special Features Name R36C 5’ GGC ACC GUA AAG ACC CAA R10B, R16B, Contains 5’- UCA UGC AGG GUC UGU GCC 3’ R19B, R36DBot, triphosphate and D36DBot R62-L21 5’ GGG AGA CAA AGU AUA GCC None L21-ScaI cleavage CUC UAA CAC AAA CGC GUC site AUG AUU GAC ACU UUA CGG UGC CCA GCA UCA AUG ACA UCA GCA UCU AC 3’ R68-TRAP 5’ GGG AC GCC CUC UAG UAA None L21-ScaI cleavage AUU UAU AUA AGA UAA GAU site AAG AUA AGA UAA GAU AAG AUA AGA UAA GAU AAG AUA CCC GAA GAU CCU 3’
Table 7.3. List of RNA Transcripts
7.1.4 Miscellaneous Reagents
ATP and GTP were purchased from Roche. ADP, ADPNP, GDP, and GMP-PMP were purchased from Sigma-Aldrich. BeF3, AlF4, and NaF were purchased from Acros
Organics. ADP-BeFx was prepared by combining ADP, BeF3, and NaF in a ratio of
1:5:25. ADP-AlFx was prepared by combining ADP, AlF4, and NaF in a ratio of 1:5:25.
ADP-NaF was prepared by combining ADP and NaF in a ratio of 1:25 (Fisher et al, 1995,
Kagawa et al, 2004, Chen et al, 2007, and Nielsen et al, 2009).
130 7.2 Methods
7.2.1 Radiolabeling and Gel Purification of RNA and DNA Oligonucleotides
RNA and DNA oligonucleotides were radiolabeled on their 5’-ends with T4
polynucleotide kinase (NEB) and γ-32P-ATP (Perkin Elmer). RNA or DNA (5 nmol) was
incubated with 1.6 µM γ-32P-ATP, 3.3 µM ATP, and 0.67 U/µL T4 polynucleotide kinase
at 37°C for one hour. Radiolabeled oligonucleotides were resolved via denaturing
PAGE. Following exposure of X-ray film to the gel, the labeled RNA or DNA was cut
from the gel and eluted with 400 µL of elution buffer (300 mM sodium acetate, pH 6.0, 1
mM EDTA, and 0.1% SDS). Radiolabeled RNA was isolated by ethanol precipitation,
lyophilized, and dissolved in double distilled water. Duplex annealing reactions
contained the radiolabeled oligonucleotide and 10 nmol of a complementary strand in
buffer containing 100 mM MOPS, pH 6.5, 500 mM KCl, and 10 mM EDTA, pH 8.0.
Duplexes were annealed by incubation at 95°C for two minutes, followed by slow
cooling to room temperature. Oligonucleotide duplexes were resolved with native
PAGE, and eluted and purified as described above for denaturing PAGE.
RNAs were 3’-end labeled by ligation of [5’-32P]pCp (England et al, 1978,
England and Uhlenbeck, 1978, and Kikuchi et al, 1978). [5’-32P]pCp was prepared by
incubating 13 µM cytidine 3’-monophosphate, 2.2 µM γ-32P-ATP, and 0.8 U/µL T4 polynucleotide kinase (NEB) at 37°C for one hour. In vitro transcribed L21-ScaI ribozyme substrates (10 µM) were incubated with 50 pmol of [5’-32P]pCp and 0.5 U/µL
T4 RNA ligase (Ambion) for sixteen hours at 4°C. 3’-end labeled RNAs were resolved on denaturing PAGE, and purified as described above.
131
7.2.2 Characterization of L21-ScaI Ribozyme Activity on RIG-I Substrates
Ribozyme cleavage reaction conditions were 50 mM Tris, pH 7.5, 20 mM NaCl,
and 10 mM MgCl2. The large excess of MgCl2 is required for activation of the ribozyme in vitro (7). L21-ScaI ribozyme (1 µM) was activated by incubation at 50°C for thirty minutes. The cleavage reaction was initiated by the addition of 70 nM precursor substrate RNA (1 nM 5’-end labeled RNA and 69 nM unlabeled RNA) and 35 µM GTP-
32 MgCl2, containing a trace amount of α- P-GTP. Incubation continued at 50°C, and
aliquots were removed at indicated timepoints. The reaction products were separated
with 20% denaturing PAGE. The gel was dried, and radiolabeled RNA in the gels was
visualized using a PhosphorImager and ImageQuant 5.2 software (GE Healthcare).
Characterization of L21-ScaI ribozyme activity with GTP analogs was performed
under the same reaction conditions as above. Following activation of the ribozyme, the
cleavage reaction was initiated by the addition of 70 nM precursor substrate RNA (1 nM
5’- and 3’-end labeled RNA and 69 nM unlabeled RNA) and either 35 µM GDP-MgCl2
or GMP-PNP-MgCl2. Samples taken at the indicated timepoints were analyzed as above.
7.2.3 End-Labeling of ssRNA with L21-ScaI Ribozyme
The reaction conditions and ribozyme activation were as described in section
7.2.1. The cleavage reaction was initiated by the addition of 70 nM precursor substrate
RNA and 0.16 µM α-32P-GTP and 1.8 µM GTP, and continued for one hour at 50°C.
132 The reaction products were separated with 20% denaturing PAGE. Following exposure of X-ray film to the gel, the labeled 5’-triphosphate-containing RNA was cut from the gel and eluted with 400 µL of elution buffer (300 mM sodium acetate, pH 6.0, 1 mM EDTA, and 0.1% SDS). Radiolabeled RNA was isolated by ethanol precipitation, lyophilized, and dissolved in double distilled water. 5’-triphosphate containing duplexes were created by annealing a complementary RNA sequence (Table 7.2), and purification of the duplex proceeded as described in section 2.1.
7.2.4 RIG-I Binding under Equilibrium Conditions
RIG-I-RNA binding reactions contained 20 mM Tris, pH 7.5, 100 mM NaCl, 0.5 mM MgCl2, 1 mM DTT, 0.01% (v/v) Igepal, 1 unit/µL RNasin (Roche), and 4% (v/v) glycerol. ATP and analogs (including equimolar MgCl2) were added to reactions at a final concentration of 1 mM where indicated. Varying concentrations of recombinant
RIG-I were incubated with 1 nM radiolabeled RNA duplex for four minutes at 25°C in a temperature-controlled aluminum block. Reactions were immediately applied to a running (0.3W/cm) native 4% polyacrylamide gel at 4°C. Following drying, radiolabeled
RNA in the gels was visualized using a Storm PhosphorImager (GE Healthcare) and
ImageQuant 5.2 software (GE Healthcare). Complex formation was plotted versus RIG-I concentration, and fitted to the Hill equation (Eq. 7.2) to measure dissociation constants of various RNA duplex substrates (Kaleidagraph 3.0, Synergy).
∗ (Equation 7.2)
133 Y = Fraction Bound Bmax = Maximum Fraction Bound L = Ligand Concentration h = Hill Coefficient
Kd = Dissociation Constant
7.2.5 Derivation of Equations for Coupled Equilibrium
⇌ ⇌
0
0
for S
0
for ES and replace in equation for S
134
Replace ES and S equations with Keq
Solve for E2S using
Solving the fraction of E2S and the total bound (E2S+ES)
1 1 1 1
When fbound = 0.5 then the following can be assumed
1. ~ 0.5 1 0.5 1 1
2. ~ 0.5
135 0.5
Solving for K1 and K2
Comparing with Fei Liu’s equation for K1
4
4
4 4 4
7.2.6 Dissociation Kinetics of RIG-I-RNA Complexes
Reaction conditions were the same as in section 2.3. Following incubation of 60 nM RIG-I with 1 nM radiolabeled RNA duplex, competition of RIG-I-RNA complexes
was initiated by the addition of the same unlabeled RNA duplex at a concentration of 1
µM. ATP or analogs were added to a final concentration of 1 mM (plus 1 mM MgCl2).
Aliquots were taken at the indicated timepoints and analyzed via 4% native PAGE at
136 4°C. Gels were dried and radiolabeled RNA was visualized using a Storm
PhosphorImager and ImageQuant 5.2 software (GE Healthcare). Dissociation rate
constants were determined by fitting the dissociation of RIG-I-RNA complexes over time
to a first order rate law (Eq. 7.3) (Kaleidograph 3.0, Synergy).
∗ (Equation 7.3)
B = Fraction of Bound RNA
B0 = Fraction of Bound RNA at time zero k = off rate t = time
7.2.7 Glutaraldehyde Crosslinking of RIG-I-RNA Complexes
Reaction conditions were the same as in section 2.3. RIG-I (100 nM) was incubated with unlabeled RNA duplexes (60 nM) and in the presence or absence of 1 mM
ATP-MgCl2 for four minutes in a temperature-controlled aluminum block at 25°C. The
RNA-protein or RNA-protein-nucleotide complexes were cross-linked chemically by the
addition of 0.02% glutaraldehyde and incubation at 25°C for an additional ten minutes.
Cross-linked RIG-I-RNA complexes were analyzed via SDS-PAGE and Coomassie blue
staining.
137 7.2.8 Measurement of RIG-I ATPase Activity with Thin Layer Chromatography
RIG-I ATPase activity was measured at 25°C in a buffer containing 400mM Tris pH 8.0, 0.1% IGEPAL, and 20 mM DTT. ATPase activity was measured under pre- steady state (1 nM RNA, 1-50 nM RIG-I) and multiple turnover (25 nM RIG-I, 1 µM
RNA) conditions. RIG-I was pre-incubated with RNA for ten minutes at 25°C in a temperature-controlled aluminum block. The reaction was initiated by the addition of
32 equimolar ATP and MgCl2, containing a trace amount of γ- P-ATP. Aliquots were removed at indicated times and spotted on a 5 cm by 20 cm thin layer chromatography plate. ATP and inorganic phosphate were resolved by thin layer chromatography in a buffer containing 1.5M formic acid and 0.3M lithium chloride. Plates were dried, and then ATP and inorganic phosphate were visualized with a Storm PhosphorImager (GE
Healthcare). The fraction of ATP hydrolyzed was quantified with ImageQuant 2.0 (GE
Healthcare) and plotted versus time. Kinetic parameters were determined by fitting to the
Michaelis-Menten equation (KaleidaGraph 3.0, Synergy) (Eq. 5).
∗ (Equation 5)
v = Initial Velocity
vmax = Maximum Velocity [S] = Concentration of Substrate (ATP)
KM = Michaelis Constant
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