REGULATION of MAMMALIAN Mrna DECAPPING

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REGULATION OF MAMMALIAN mRNA DECAPPING

You Li

A Dissertation submitted to the Graduate School-New Brunswick

Rutgers, The State University of New Jersey

and

The Graduate School of Biomedical Sciences

University of Medicine and Dentistry of New Jersey

in partial fulfillment of the requirements

for the degree of

Doctor of Philosophy

Graduate Program in Cell and Development Biology

written under the direction of

Dr. Megerditch Kiledjian

and approved by

______

New Brunswick, New Jersey

MAY, 2011

ABSTRACT OF THE DISSERTATION

Regulation of Mammalian mRNA Decapping

YOU LI

Dissertation Director

Dr. Megerditch Kiledjian

The modulation of mRNA degradation plays a critical role for regulation of gene

expression. A major mRNA decay pathway in mammals proceeding from the 5′ to 3′ end

is initiated with shortening of 3′ poly (A) tail, followed by the cleavage of the 5′ cap

structure (decapping) and degradation of the mRNA body by the Xrn1 exoribonuclease.

Dcp2 is a well characterized mRNA decapping enzyme. It is an RNA binding protein that

must bind the RNA in order to recognize the cap for hydrolysis. We identified mRNAs

bound by human Dcp2 by coimmunoprecipitation and microarray analysis. Further

biochemical assays identified a sequence element of 60 nucleotides at the 5´ terminus of the mRNA encoding Rrp41 as a specific Dcp2 substrate which could specifically bind

Dcp2 protein and enhance Dcp2 decapping. Mutational analysis of this element revealed

a stable stem-loop structure (termed DBDE) contained in the first 33 nucleotides is

ii critical for Dcp2 decapping stimulation. A bioinformatic search was carried out to identify a subset of mRNAs that have a comparable stem-loop structure at the 5′ end as potential Dcp2 substrates.

We identified a second cytoplasmic mRNA decapping enzyme Nudt16 in mammalian cells and established stable MEF cells that contained reduced level of Dcp2 and/or Nudt16 protein. Using these MEF cells, we demonstrated the distinct roles for

Dcp2 and Nudt16 in nonsense mediated mRNA decay (NMD), ARE-mediated decay and miRNA mediated gene silencing. Our results indicated that NMD preferentially utilizes

Dcp2 rather than Nudt16; Dcp2 and Nudt16 are redundant in miRNA mediated silencing; and Dcp2 and Nudt16 are differentially utilized for ARE-mRNA decay.

At last, we demonstrated that anti-viral immune response was regulated by Dcp2 in mouse embryonic fibroblast. In MEF cells lacking Dcp2 protein, interferon-mediated anti-viral response was significantly elevated. We identified IRF7, a crucial transcription factor in anti-viral immunity, as the direct target of Dcp2. Knockdown of Dcp2 led to stabilization of IRF7 mRNA and increased IRF7 protein level. Therefore, Dcp2 functions as a negative regulator of interferon-mediated anti-viral immunity. Interestingly, Dcp2 expression is also induced in virus infection, suggesting a negative feedback loop in the anti-viral immune response.

iii DEDICATION

This thesis is dedicated to my parents,

Xunqi Li and Guanrong Diao, who have been giving me love and support all through my life.

iv ACKNOWLEDGMENT

First of all, I would like to thank my advisor, Dr. Mike Kiledjian for all of his guidance, support and advice during the past five years. Mike is a great supervisor. He is knowledgeable and insightful and has always been patient and accessible to provide help all through my PhD study. He is open-minded and willing to discuss any ideas I have during my research. His way of critical thinking and enthusiasm to science set up a great model for me to follow. I am also grateful for the writing and presentation training under his guidance. I have learned a lot from him, not only about science but also personal and communicational skills. The research experiences in Kilejian lab will be a great asset of my life.

I would like to thank the members of my thesis committee, Dr. Lori Covey, Dr.

Sam Gunderson, Dr. Paul Copeland and Dr. Ping Xie for their advice and suggestions during my PhD study. In particular, I would like to thank Dr. Lori Covey and Dr. Ping

Xie for their advice and technical assistance in the study of regulation of Dcp2 on anti- viral immunity. Immunology is a new field to me and to the lab, and Dr. Lori Covey and

Dr. Ping Xie have taught me important concepts of immunology and have made great suggestions during the course of my research.

I’d like to express my gratitude to the current and past members of the lab, Xinfu

Jiao, Mangen Song, Madel Durens, Sophie Bail, Hudan Liu, Shin-wu Liu, and Vincent

Shen for their support, help and friendship in the past five years. I would especially like to thank Xinfu for his technical support and generous contribution of time in sharing his

v research experiences throughout my PhD study. I am grateful to Mangen Song who helped me a lot to get started when I initially joined the lab.

I would like to acknowledge the members in the neighboring labs, including

Covey lab, Xie lab, Denhardt lab, Gumderson lab for sharing facilities and reagents and providing indispensable help to my research work. I would like to especially express my gratitude to Carissa Moore in Xie lab, who spent numerous hours explaining concepts and experimental techniques in immunology to me, as well as sharing her research experience with me.

Lastly, and most importantly, I would like to express my deepest gratitude to my parents, Xunqi Li and Guanrong Diao, for their love and support throughout my life. I love them both with all of my heart.

vi TABLE OF CONTENTS

ABSTRACT OF THE DISSERTATION ...... ii

DEDICATION...... iv

ACKNOWLEDGMENT ...... v

TABLE OF CONTENTS ...... vii

LIST OF TABLES ...... xi

LIST OF FIGURES ...... xii

Introduction ...... 1

Eukaryotic decapping enzymes ...... 2

The Dcp2 mRNA decapping enzyme ...... 2

The DcpS scavenger decapping enzyme ...... 4

The Nudt16 (X29) decapping enzyme ...... 5

Regulators of decapping ...... 6

Activators of Dcp2 decapping ...... 6

Inhibitors of Dcp2 decapping...... 9

Regulators of DcpS decapping...... 11

Transcript specific decapping ...... 13

Protein-mediated recruitment of Dcp2...... 13

vii Direct recruitment of Dcp2 ...... 15

Localization of decapping enzymes ...... 17

Cytoplasmic recapping ...... 19

Implication of the decapping enzymes in human disorders ...... 20

Potential role of DcpS in Spinal Muscular Atrophy ...... 20

Potential role of Dcp2 in X-linked mental retardation...... 21

Anti-viral innate immunity ...... 22

Materials and Methods ...... 27

Plasmid constructs ...... 27

Plasmids used in characterization of DBDE ...... 27

Plasmids used in study of differential utilization of decapping enzymes ...... 28

Plasmids used in study of decapping in anti-viral immunity ...... 29

Generation of RNA in vitro ...... 30

Coimmunopurification of RNAs associated with Dcp2 ...... 31

Microarray Analysis ...... 32

Microarray of RNAs bound by Dcp2...... 32

Microarray of RNAs in MEF cells...... 33

RT-PCR ...... 33

GST fusion protein copurification ...... 34

viii In vitro RNA decapping assay ...... 34

Electrophoretic mobility shift assays ...... 35

RNase mapping ...... 35

Bioinformatics search for Dcp2 substrates ...... 36

siRNA transfection ...... 36

Cell based mRNA decay assays ...... 37

Electroporation of RNA ...... 37

Reporter assays ...... 38

Western Blot ...... 39

Immunofluorescence Assay ...... 39

Lentivirus production and infection ...... 40

Chapter I: Transcript-specific Decapping of Dcp2 and Characterization of a Dcp2- binding Element ...... 45

Summary ...... 45

Introduction ...... 46

Results ...... 49

Discussion ...... 86

Chapter II: Differential Utilization of Decapping Enzymes in Mammalian mRNA

Decay Pathways ...... 93

Summary ...... 93

ix Introduction ...... 94

Results ...... 98

Discussions ...... 116

Chapter III: Regulation of Decapping in Interferon-mediated Anti-viral Immunity

...... 121

Summary ...... 121

Introduction ...... 122

Results ...... 125

Discussion ...... 140

Concluding remarks ...... 146

References ...... 149

Curriculum Vita ...... 162

x LIST OF TABLES

Table I. Primers used in semi-quantitative RT-PCR. 41

Table II. Primers used for real-time PCR 42

Table III.Primers used for generating RNAs in vitro 43

Table IV. Dcp2 bound mRNAs 50

xi LIST OF FIGURES

Figure 1. Identification of Rrp 41 mRNA as a substrate directly

bound by Dcp2 54

Figure 2. Rrp41 mRNA is preferentially decapped by Dcp2. 57

Figure 3. Dcp2 target sequence on Rrp41 mRNA is located in

the 5′UTR. 60

Figure 4. The Rrp41 5´UTR is an autonomous element conferring

higher decapping activity. 62

Figure 5. Identification of hDcp2 binding element (2xDE). 64

Figure 6. Dcp2 regulates Rrp41 mRNA stability in cells. 67

Figure 7. The first 33 nucleotides of the 2xDE element are critical

for enhanced Dcp2 decapping and binding. 71

Figure 8. The first 33 nt of the 2xDE forms a stem-loop secondary

structure. 74

Figure 9. The intact stem-loop structure is critical for promoting

Dcp2 decapping. 76

Figure 10. Mutational analysis of the stem-loop structure. 78

Figure 11. Positioning of the DBDE stem-loop structure less than 10

nucleotides from the 5´ cap is important for Dcp2-mediated

enhanced decapping. 80

Figure 12. The Ndufb7 5´ UTR stem-loop structure is also capable

of enhancing decapping. 82

xii Figure 13. Enhancement of decapping by DBDE-like 5′ end stem-loop

structure in cellular mRNAs. 84

Figure 14. Nudt16 is a cytoplasm mRNA decapping enzyme. 99

Figure 15. Reduction of Nudt16 protein levels in wild type and

Dcp2β/β MEFs. 101

Figure 16. P-body numbers are unaltered in MEF cells with undetectable

levels of Dcp2 and/or reduced levels of Nudt16 protein. 103

Figure 17. Contribution of decapping enzymes to NMD. 106

Figure 18. Dcp2 and Nudt16 are redundant in miRNA mediated

gene silencing. 110

Figure 19. Differential utilization of Dcp2 and Nudt16 in ARE-mediated

decay. 114

Figure 20. A subset of genes involved in anti-viral immune response were

upregulated in Dcp2β/β MEF cells. 126

Figure 21. IFN mediated anti-viral immune response was elevated in

Dcp2β/β MEF cells following lentivirus infection. 129

Figure 22. Catalytic activity of Dcp2 is required for the regulation of

anti-viral immune response. 132

Figure 23. Dcp2 regulates mRNA stability and protein expression of IRF7. 135

Figure 24. Dcp2 expression is induced in virus infection. 138

xiii 1

Introduction

Regulation of mRNA stability has emerged as a key step in the control of gene

expression and provides a rapid response mechanism to cellular events. The steady-state

level of any given mRNA is determined by the rate of transcription and degradation. The

decay of mRNA can occur via different pathways. In eukaryotic cells, bulk mRNA decay

typically initiates with the shortening of the 3' poly(A) tail, followed by degradation of

the mRNA body in 5' to 3' direction or 3' to 5' direction (Coller & Parker, 2004). In the

5' end decay pathway, deadenylation leads to cleavage of the monomethyl guanosine

(m7G) mRNA cap by the Dcp2 decapping enzyme (Dunckley & Parker, 1999; Lykke-

Andersen, 2002; van Dijk et al, 2002; Wang et al, 2002b), exposing the 5' end monophosphorylated RNA to progressive 5' to 3' exoribonucleolytic decay by Xrn1 in the

cytoplasm (Decker & Parker, 1993; Hsu & Stevens, 1993). In the 3' decay pathway, degradation from the 3' end following deadenylation is carried out by the cytoplasmic

RNA exosome complex (Anderson & Parker, 1998; Wang & Kiledjian, 2001), which is a multisubunit 3'-to-5' exoribonuclease complex (Liu et al, 2006) and interestingly also contains endoribonuclease activity (Lebreton et al, 2008; Schaeffer et al, 2009; Schneider et al, 2009). The resulting cap structure from 3' end decay is hydrolyzed by the scavenger decapping enzyme DcpS (Liu et al, 2002). Therefore, there are at least two classes of decapping enzymes, each primarily functioning in one of the two exonucleolytic mRNA decay pathways. A third cellular decapping enzyme has also been identified in Xenopus, the X29 nuclear decapping enzyme, which can preferentially hydrolyzes the cap of U8 snoRNA in vitro (Ghosh et al, 2004). Interestingly, its mammalian homolog Nudt16 is a

cytoplasmic decapping enzyme which could regulate the stability of a subset of mRNAs

(Song et al, 2010). Therefore, there appear to be multiple decapping enzymes in

mammalian cells and each one could contribute to overall mRNA decay.

Eukaryotic decapping enzymes

The Dcp2 mRNA decapping enzyme

The Dcp2 decapping enzyme is a member of the NUDIX (nucleotide diphosphate

linked to moiety X) hydrolase family (Dunckley & Parker, 1999; Lykke-Andersen, 2002;

van Dijk et al, 2002; Wang et al, 2002b), which is characterized by the highly conserved

23-residue NUDIX motif, GX5EX7REUXEEXGU (where U represents a hydrophobic residue and X is any residue). It cleaves the 5' methylated cap of an mRNA to produce m7GDP and 5' monophosphate RNA in the presence of manganese or high concentration

of magnesium (Piccirillo et al, 2003; Steiger et al, 2003; van Dijk et al, 2002). Dcp2 can

cleave m7G-capped RNA and m2,2,7G-capped RNA while unmethylated capped RNA is a poor substrate (Cohen et al, 2005; Piccirillo et al, 2003; Steiger et al, 2003; van Dijk et al,

2002). Importantly, the RNA body contributes to the substrate specificity, since Dcp2 does not hydrolyze cap structure without a linked RNA moiety (Piccirillo et al, 2003) and

preferentially utilizes RNAs that are longer than 25 nucleotides (Steiger et al, 2003).

Consistent with the requirement of the RNA body for decapping, Dcp2 is an RNA

binding protein that can bind RNA in vitro and in cells (Li et al, 2008; Piccirillo et al,

2003) and its activity is competed by uncapped RNA but not m7GpppG cap structure

alone in vitro (Piccirillo et al, 2003; Steiger et al, 2003; van Dijk et al, 2002). These facts

indicate that Dcp2 recognizes the mRNA substrates by simultaneously interacting with

the cap and the RNA body.

The catalytic site within the NUDIX motif of Dcp2 contains three conserved

glutamate residues that coordinate a divalent cation (manganese or magnesium) essential

for cap hydrolysis (She et al, 2006). Mutations of the conserved glutamate residues abolish decapping activity (Dunckley & Parker, 1999; van Dijk et al, 2002; Wang et al,

2002b). In addition to the NUDIX motif, Dcp2 protein contains two other evolutionarily conserved regions, termed Box A and Box B. Box A is at the amino terminus of Dcp2

and appears to be important for the catalytic fidelity of decapping. Truncations in Box A

compromise Dcp2’s ability to exclusively generate m7GDP upon hydrolysis, and results

in release of m7GMP as a second product (Piccirillo et al, 2003). Box A is also important

for the interaction between yeast Dcp2p and the decapping activator Dcp1p (She et al,

2008). Box B is located at the C-terminus of the NUDIX domain and is shown to be

necessary for the RNA binding property of Dcp2 and its truncation decreases both RNA-

binding and decapping activity of Dcp2 in vitro (Piccirillo et al, 2003).

Structural studies by NMR spectroscopy revealed that yeast Dcp2 recognizes its

substrate by a bipartite surface formed by the NUDIX domain that interacts with the cap

and the RNA body (Deshmukh et al, 2008). The interaction with the cap is relatively

weak while the interaction with RNA body is relatively strong and mediated by a

conserved RNA binding channel formed by positive charged residues encompassing the

NUDIX domain containing Box B (Deshmukh et al, 2008). The channel confers

association with the RNA body by distributing weak ionic interactions over a large

surface. Mutations in several positive charged residues lining the channel decrease

decapping activity in vitro (Deshmukh et al, 2008). The cap structure is recognized

exclusively during the catalytic step, which is much slower compared to the substrate

binding and product release. Thus the chemical catalysis is the rate limiting step for Dcp2

decapping (Deshmukh et al, 2008).

The DcpS scavenger decapping enzyme

DcpS is a member of the Histidine Triad (HIT) family of nucleotide hydrolases

(Liu et al, 2002), which are characterized by the conserved HIT motif HUHUHU (where

U denotes a hydrophobic residue) (Seraphin, 1992) contained in a larger HIT fold region.

Unlike Dcp2, DcpS functions on cap structure or capped oligonucleotides shorter than 10

bases, but not on capped RNA. It cleaves the cap structure between the γ and β

phosphates to generate m7GMP and a nucleotide diphosphate (Liu et al, 2002). It can also

dephosphorylate m7GDP, generated by Dcp2, into m7GMP plus Pi (van Dijk et al, 2003).

Mutations of conserved histidines in the HIT motif abolish decapping activity confirming

the significance of this motif in catalysis (Liu et al, 2002). The N7-methyl moiety is

essential for substrate specificity of DcpS, as it does not function on unmethylated cap

(Liu et al, 2002).

Crystal structure of DcpS has revealed that it exists as a symmetric homodimer in

the ligand-free form and an asymmetric homodimer when bound to its ligand (Chen et al,

2005; Gu et al, 2004). The DcpS homodimer contains a distinct domain swapped N-

terminal segment and a C-terminal domain linked by a flexible hinge region generating

two distinct cap binding/hydrolysis pockets (Chen et al, 2005; Gu et al, 2004). In the

ligand-free form, both N-terminal domains of the homodimer are in the open

conformation; while in the ligand-bound form, the homodimer simultaneously forms a

closed productive conformation on one side and an open nonproductive conformation on

the other, with substrate bound at the C-terminal domain of each monomer (Chen et al,

2005). The structure suggests that the flexible hinge region enables the N-terminal domain to flip back and forth, alternating between open and closed conformations. This mechanism is supported by kinetic studies that demonstrate the significance of dynamic conformational changes of the N-terminal domain for cap hydrolysis and confirm the mutually exclusive hydrolysis function between the two catalytic active sites (Liu et al,

2008).

The Nudt16 (X29) decapping enzyme

X29 was initially identified in Xenopus as a U8 snoRNA binding protein. It is a member of the NUDIX family of proteins and was shown to possess decapping activity

(Ghosh et al, 2004). X29 is a nucleolar protein capable of specifically binding U8 snoRNA in vitro (Ghosh et al, 2004; Tomasevic & Peculis, 1999), and in the presence of

Mg2+, cap hydrolysis is highly specific for U8 snoRNA (Ghosh et al, 2004; Peculis et al,

2007); in contrast, in the presence of Mn2+ , X29 possessed a more pleiotropic decapping

activity where all RNAs tested were decapped at high efficiency (Peculis et al, 2007).

X29 can cleave both methyl-capped RNA and unmethyl-capped RNA (Ghosh et al, 2004).

Although it has been implicated in nucleolar decapping, a direct role for this protein in cellular U8 snoRNA stability has yet to be addressed.

The crystal structure of X29 confirmed the presence of a NUDIX fold and

demonstrated a homodimeric quaternary structure. Each X29 monomer consists of two

domains with one domain containing the Nudix motif and the other domain forming the monomer-monomer contact interface (Scarsdale et al, 2006). The structure of X29 displays a highly dipolar surface charge distribution that possibly creates a plausible

RNA binding channel on the positive face of the protein. Metal binding only occurred in the presence of nucleotides, suggesting substrates binding may be required for metal binding to form an active holo-enzyme (Scarsdale et al, 2006).

The Nudt16, mammalian ortholog of X29, also possesses decapping activity

(Taylor & Peculis, 2008). Although conserved in metazoans, an obvious ortholog of

Nudt16 is lacking in S. cerevisiae, C. elegans and Drosophila (Taylor & Peculis, 2008).

Interestingly, Nudt16 was reported to be a cytoplasmic decapping protein in mammalian cells which could regulate the stability of a subset of mRNAs (Song et al, 2010). For instance, Amotl2 mRNA was significantly stabilized when Nudt16 protein levels were reduced (Song et al, 2010). Therefore, unlike its Xenopus counterpart, the mammalian

Nudt16 protein is involved in cytoplasmic mRNA decay. Like Dcp2, Nudt16 decaps capped RNA but not cap structure alone (Song et al, 2010). Curiously, Nudt16 was also reported to be an inosine diphosphatase linked to prevention of single stranded DNA breaks (Iyama et.al 2010) which hydrolyzes purine nucleoside diphosphates to the corresponding nucleoside monophosphates. Whether the diphosphatase activity and mRNA decapping activity of Nudt16 constitute distinct activities and whether mRNA stability functions in single stranded DNA breaks remain to be determined.

Regulators of decapping

Activators of Dcp2 decapping

Several proteins have been identified to activate Dcp2 decapping. They can promote decapping by either directly accelerating the cap hydrolysis of Dcp2 or recruiting Dcp2 onto mRNA substrates to increase the local concentration of decapping enzyme. Dcp1 is a critical cofactor of Dcp2 that is required for efficient decapping in S. cerevisiae (Steiger et al, 2003). It is conserved in eukaryotic cells and two homologs,

Dcp1a and Dcp1b, are identified in humans. Structural analysis of Dcp1 protein reveals it contains an EVH1 (Ena/VASP homolog) domain that is a proline-rich protein interaction module (She et al, 2004). Two conserved regions are mapped on the molecular surface of

Dcp1. One is contained within the EVH1 domain and likely to be the binding site for regulatory proteins. The other site is essential for the stimulation of Dcp2 decapping (She et al, 2004). Dcp1 interacts with the N-terminal domain of Dcp2 protein and copurifies with Dcp2 in yeast and mammalian cells (Dunckley & Parker, 1999; Lykke-Andersen,

2002; She et al, 2006; She et al, 2008). Yeast Dcp2 decapping activity is stimulated by

Dcp1 in vitro (Steiger et al, 2003). Structural analysis of the yeast Dcp1/Dcp2 heterodimer complex demonstrated that Dcp1 enhances Dcp2 decapping by accelerating the chemical catalysis step. Binding of Dcp1 orients the N-terminus of Dcp2 towards the active site, transforming from a nonproductive open conformation to a productive closed conformation (She et al, 2008). However the interaction interface of yeast Dcp1 and

Dcp2 is not conserved in higher eukaryotes (She et al, 2008) suggesting these two proteins may not form a similar heterodimeric complex in higher eukaryotes. This point is further supported by the lack of detected stimulation of Dcp2 in vitro by recombinant

Dcp1 from C.elegans, drosophila and human (Cohen et al, 2005; Lin et al, 2008; Lykke-

Andersen, 2002), suggesting Dcp1-Dcp2 interaction in higher eukaryotes requires

additional factors.

Stimulators of decapping that appear species restricted have also been identified.

The enhancers of decapping, Edc1 and Edc2, are present exclusively in yeast and have been shown to stimulate Dcp2 activity in vitro (Steiger et al, 2003). In metazoans, the

Hedls protein (also referred to as Edc4 or Ge-1) enhances Dcp2 decapping in vitro

(Fenger-Gron et al, 2005b). The stimulation appears to be a consequence of its ability to promote a stable interaction of Dcp1 with Dcp2, which do not stably interact otherwise, as determined by immunoprecipitations (Fenger-Gron et al, 2005b). The capacity for yeast Dcp2 and Dcp1 to heteromerize in the absence of additional factors is consistent with the absence of a Hedls ortholog in this organism since a major function of Hedls appears to be to promote this heteromerization in metazoans. Unlike Edc1, Edc2 and

Hedls, Edc3 is a decapping stimulator that is conserved in all eukaryotes (Dunckley et al,

2001; Fenger-Gron et al, 2005b; Kshirsagar, 2004). It is an RNA-binding protein and can specifically bind to and promote decapping of the Rps28b mRNA in S.cerevisiae (Badis et al, 2004). A conserved hairpin structure in the 3' UTR of the Rps28b mRNA is recognized by the Rps28 protein, which in turn recruits the Edc3 protein (Badis et al,

2004). The decapping is stimulated by an interaction of Edc3 with multiple decapping factors including Dcp1 and Dcp2 (Decker et al, 2007; Harigaya et al; Tritschler et al,

2007). Its interaction with Dcp2 is mediated by a short peptide sequence located C- terminal to the catalytic domain of Dcp2 (Harigaya et al, 2010). Therefore Edc3 appears to serve as a platform to recruit the decapping machinery onto the Rps28b mRNA.

Another class of decapping activators, including Dhh1 (Rck/p54 in human) and

Pat1 (Pat1L in human), stimulate decapping by promoting translational repression (Coller

& Parker, 2005). Overexpression of Dhh1 and Pat1 results in general repression of yeast

mRNA translation and enhances decapping (Coller & Parker, 2005) by releasing the cap- binding complex from the mRNA 5´ end to enable access for the decapping complex.

Pat1 also forms a complex with the Lsm1-7 protein complex and has been shown to bind specifically to deadenylated mRNAs and activate decapping (Bouveret et al, 2000;

Chowdhury et al, 2007; Chowdhury & Tharun, 2009; Tharun et al, 2000; Tharun &

Parker, 2001). The Upf proteins, which are required for nonsense-mediated decay (NMD), also promote decapping. Decapping is induced by NMD (Couttet & Grange, 2004;

Lejeune et al, 2003; Muhlrad & Parker, 1994) and all three Upf proteins coimmunopurify with Dcp2 (Lejeune et al, 2003; Lykke-Andersen, 2002), indicating that Upf NMD factors recruit decapping protein to degrade mRNAs with a premature termination codon.

Inhibitors of Dcp2 decapping

It is a reasonable expectation that cap-binding proteins could provide a physical barrier to Dcp2 access and consequently prevent decapping. There are at least two classes of cap binding proteins. The first consists of canonical cap binding proteins that bind the m7G cap directly independent of other moieties. This class includes the

eukaryotic initiation factor 4E (eIF4E) cap-binding protein or the nuclear cap-binding protein, CBP20. The second is a more recently described non-canonical cap binding protein class that requires the cap to be linked to an RNA moiety and are unable to bind the m7G cap directly in the absence of a linked RNA (Jiao et al, 2006). An example of

this is the Dcp2 decapping protein itself (see above) and the decapping inhibitor, Variable

Charged X chromosome (VCX)-A protein.

The eIF4E cap-binding protein negatively regulates decapping due to its ability to bind the 5' cap structure and consequently limit access of the decapping complex to the mRNA 5' end. eIF4E can inhibit decapping in vitro in both yeast and mammalian cells

(Khanna & Kiledjian, 2004; Schwartz & Parker, 1999; Schwartz & Parker, 2000), and

RNAs with synthetic cap structures that bind eIF4E with higher affinity are more stable in vivo (Grudzien et al, 2006). The 3' end poly(A) tail can also negatively influence decapping. This is likely to be mediated by the poly(A) binding protein, PABP

(Caponigro & Parker, 1995; Wilusz et al, 2001a), as PABP tethered onto mRNAs impairs decapping, indicating its intrinsic inhibitory property of decapping is independent of the poly(A) tail (Coller et al, 1998). Although this inhibition is primarily mediated through an interaction with eIF4E, in vitro decapping reactions with mammalian PABP demonstrate a direct role of PABP in decapping inhibition as well. PABP is also a non- canonical cap binding protein that can directly associate with the 5' end of capped RNA to curtail decapping (Khanna & Kiledjian, 2004). Interestingly, the direct association with the cap is limited to the mammalian PABPs tested and is not observed with the yeast

Pab1p. Therefore, the poly(A) binding protein has the capacity to inhibit decapping either through a direct association with the cap in mammals, or more broadly, through a network of interactions involving eIF4E and eIF4G which juxtapose the two ends of the mRNA to stabilize the cap by preventing access to decapping enzymes.

Similar to Dcp2, the VCX-A protein is an RNA-binding protein that preferentially associates with the 5' cap and is a member of a novel family of non-canonical cap binding

proteins that require dual recognition of the cap and RNA since they recognize the cap structure when it is linked to an RNA (Jiao et al, 2006; Piccirillo et al, 2003). The cap binding property of VCX-A impedes Dcp2-directed decapping in vitro and stabilizes mRNAs in cells. Moreover, the affinity of VCX-A to the cap and its capacity to inhibit decapping is further accentuated by Dcp2 in vitro in a mechanism that further ensures stabilization of an mRNA in the presence of Dcp2 (Jiao et al, 2006). A similar non- canonical cap-binding property has also been reported for the YB-1 protein where cap recognition is dependent on the presence of an RNA (Evdokimova et al, 2001).

Expression of YB-1 is shown to stabilize mRNAs suggesting it too could be an inhibitor of decapping. Structural analysis of non-canonical cap binding proteins complexed with capped oligoribonucleotides will reveal the interesting dual recognition property of these proteins.

Regulators of DcpS decapping

One well characterized modulation of DcpS decapping involves the DcpS ortholog in S.cerevisiae, Dcs1p, by the catalytically inactive Dcs2p (Malys & McCarthy,

2006). Despite 90% similarity between the two Dcs proteins and the presence of an intact HIT hydrolase motif in both proteins, Dcs2p does not possess the ability to hydrolyze cap structure (Liu et al, 2002; Liu et al, 2004b). The Dcs1p-Dsc2p heterodimer shows remarkably reduced rates of cap hydrolysis compared to the Dcs1p homodimer in vitro. Interestingly, Dcs2p is induced upon nutrient stress in yeast and can function as a negative regulator of Dcs1p under stress conditions (Malys & McCarthy, 2006).

However, this regulatory system appears to be restricted to yeast as a Dcs2p homolog

does not appear to be present in other organisms.

Factors regulating mammalian DcpS have not been extensively studied and only two interactions have been reported. First, consistent with its role in degrading cap structure generated by exosome-mediated 3´ to 5´ decay of an mRNA, DcpS activity copurifies with the exosome (Wang & Kiledjian, 2001). Whether this is a direct or

indirect interaction and the stoichiometry involved are not known. Second, the DcpS

ortholog in C.elegans is encoded by a bicistronic transcript together with the cytosolic flavin reductase, fre-1 (NR-1 in humans) and induced by stress, suggesting a functional association between these two proteins in stress response (Kwasnicka et al, 2003).

Although both proteins are encoded by distinct genes in humans, they nevertheless

appear to be coordinately expressed in human tissues (Kwasnicka et al, 2003) and coimmunopurify (Kwasnicka-Crawford & Vincent, 2005). Intriguingly, the increased

cytotoxicity by menadione in human embryonic kidney cells overexpressing NR-1, is

reversed by the expression of DcpS (Kwasnicka-Crawford & Vincent, 2005). The

physiological significance of DcpS under various cellular stress environments is not

known. However, exposure of cells to environmental stresses can trigger a stress

response leading to intracellular changes that manifest in alterations of mRNA synthesis,

processing, export, translation and stability (Anderson & Kedersha, 2008; Bond, 2006;

Kedersha & Anderson, 2002). These alterations may result in a greater cellular cap

structure load as a result of the block in pre-mRNA processing and increased mRNA

decay. DcpS may be relevant as an adaptive response following a stress response to help

clear cap structure and assist cells upon return to normal homeostasis.

Transcript specific decapping

Protein-mediated recruitment of Dcp2

Initial studies of Dcp2 have focused on its role as a general mRNA decapping

enzyme that can target all mRNAs. Subsequent studies have revealed that Dcp2 can be

activated by specific sequence elements within mRNA transcripts. One such element is

the AU-rich elements (ARE) found in the 3′ untranslated region (UTRs) in a class of

mRNAs encoding transcription factors or cytokines and proto-oncogenes that regulate

either cell proliferation or response to external stimuli. AREs confer rapid degradation

onto the host mRNAs by a process referred as ARE-mediated decay (AMD) which involves the interactions between ARE elements and multiple ARE-binding proteins

(Barreau et al, 2005). AREs seem to differentially activate both 3′ to 5′ and 5′ to 3′ decay pathways by recruiting the exosome complex or decapping complex respectively (Chen et al, 2001; Gao et al, 2001; Lykke-Andersen & Wagner, 2005; Mukherjee et al, 2002;

Murray & Schoenberg, 2007; Stoecklin et al, 2006). Several ARE-binding factors that destabilize the mRNA, including TTP, BRF-1 and KSRP interact with the decapping complex and are thought to promote rapid decay of an mRNA by directly or indirectly recruiting Dcp2 (Chou et al, 2006; Fenger-Gron et al, 2005b; Lykke-Andersen & Wagner,

2005). In particular, TTP can specifically promote ARE-dependent Dcp2 decapping in vitro (Fenger-Gron et al, 2005b), suggesting these factors recruit decapping enzymes onto the target mRNA to activate decapping. Consistently, overexpression of Dcp2 results in accelerated decay of a reporter mRNA containing the GM-CSF ARE (Fenger-Gron et al,

2005b), and knockdown of Dcp2 is reported to impair 5' decay of c-fos ARE-containing mRNAs (Murray & Schoenberg, 2007). Depletion of the decapping activator Lsm1 by

RNAi impairs ARE-mediated decay in human cells (Stoecklin et al, 2006), suggesting recruitment of decapping enzymes onto ARE-mRNA could also be mediated by the

Lsm1-7 complex. Interestingly, depletion of 5' exoribonuclease Xrn1 significantly stabilizes the reporter mRNA containing the GM-CSF ARE in mammalian cells while depletion of Dcp2 has no effect on the decay of the same RNA (Stoecklin et al, 2006).

This discrepancy might be due to an inefficient knock down of Dcp2 in these cells or indicate commitment to the 5' decay pathway requires an initial decapping step followed by an inhibition of the 3' decay pathway. Alternatively, multiple decapping enzymes may contribute to cap hydrolysis in mammalian cells.

A more recently identified cis-element that can trigger decapping when positioned at the 3′ end is a stretch of U residues or U tract (Mullen & Marzluff, 2008; Rissland &

Norbury, 2009; Song & Kiledjian, 2007). The initial indication that a 3′ terminal U-tract can promote decapping was provided by Shen and Goodman (Shen & Goodman, 2004).

They demonstrated that following miRNA-directed cleavage of mRNAs in plants and mice, 3′ oligouridylated mRNA fragments are detected corresponding to the 5′ cleavage product (Shen & Goodman, 2004). Interestingly, a majority of the mRNA fragments containing a U-tract were missing sequences at the 5′ end indicating they have undergone decapping. A U-tract with an optimal length of 5 U’s at the 3' end of a generic RNA triggers Dcp2-mediated decapping in mammalian cell extract and the stimulation is directed by the Lsm1-7 complex (Song & Kiledjian, 2007) which has the capacity to recognize terminal oligo U stretches (Chowdhury et al, 2007; Song & Kiledjian, 2007).

Significantly, a similar stimulation of decapping by oligouridylation is demonstrated for endogenous mRNAs. The cell cycle dependent histone mRNAs, which contain a stem-

loop structure at their 3' end rather than the poly(A) tail, are shown to undergo 3´ end oligouridylation to promote degradation (Mullen & Marzluff, 2008). Sequencing of histone mRNA decay intermediates indicates that the U-tract could promote decay from both the 5' and/or 3' ends and is dependent on the Lsm1-7 complex. A role of decapping is confirmed by a decrease in decay of histone mRNAs following a knockdown of Dcp2

(Mullen & Marzluff, 2008). At present it is unclear whether the uridylation occurs only on unadenylated RNAs or whether it also occurs on adenylated RNAs in mammalian cells. Uridylation in S. pombe can occur at the 3' end of polyadenylated mRNAs, but only one or two uracil residues appear to be added (Rissland & Norbury, 2009).

Decapped polyadenylated mRNA decay intermediates with a terminal U could be detected in S. pombe, indicating the uridylation triggers deadenylation-independent decapping (Rissland & Norbury, 2009). Therefore, uridylation is a novel and perhaps widespread mechanism for triggering decapping and mRNA decay.

Direct recruitment of Dcp2

Dcp2 is an RNA-binding protein, and like other RNA-binding proteins, also contains the intrinsic property to distinguish between different mRNA substrates. Dcp2 can specifically associate with and selectively decap a subset of cellular mRNAs (Li et al,

2008), indicating activation of decapping can be regulated by the affinity of Dcp2 to target mRNAs. Hybridization of DNA oligonucleotides to the RNA 5´ end significantly reduces decapping activity in vitro demonstrating the significance of the 5' cap proximal region of an mRNA for Dcp2 decapping (Steiger et al, 2003). Microarray analysis of mRNAs selectively bound by human Dcp2 identified a subset of mRNAs including the

mRNA encoding a subunit of the exosome complex protein, Rrp41. The 5' UTR of human Rrp41 mRNA can specifically recruit Dcp2 and direct decapping of the mRNA in vitro and in cells (Li et al, 2008). Particularly, a stem-loop structure, termed the Dcp2 binding and decapping element (DBDE) at the 5' terminus of the Rrp41 mRNA is essential to promote decapping (Li et al, 2009). Importantly, the secondary structure, but not the primary sequence, is critical for Dcp2 decapping. The stem loop structure consists of an 11 nucleotide stem and 8 nucleotide loop. While a minimal length of an

8bp stem is required to stimulate decapping, the size of the loop does not significantly contribute to decapping stimulation. Positioning of the stem-loop is important with optimal Dcp2 binding and decapping stimulation observed when placed within 10 nucleotides of the cap. The broad significance of the DBDE as a general activator of decapping when positioned at the 5' end of mRNAs was demonstrated by the identification of related structures at the 5' end of specific mRNAs that are selectively decapped by Dcp2 in vitro (Li et al, 2009). 5' end structures can also suppress decapping as demonstrated by the C. elegans splice leader (SL) sequence, which reduces Dcp2 decapping (Cohen et al, 2005). RNAs containing the 22-nt trans-spliced SL sequence at their 5' ends are inefficiently decapped by Dcp2 in vitro (Cohen et al, 2005). In addition, all trypanosome mRNAs contain a 39 nt spliced leader sequence at their 5' end, and this sequence also inhibits Dcp2-like decapping activity in trypanosome extract (Denker et al,

1996). Therefore the sequence at the 5' end of an mRNA is critical for either the recruitment or accessibility to Dcp2 and ultimately stability of the mRNA.

Localization of decapping enzymes

The Dcp2 decapping enzyme is primarily a cytoplasmic protein and enriched in

distinct cytoplasmic foci termed processing bodies (P-bodies), which also contain the

decapping activators Dcp1, Edc3, Hedls, Dhh1 and the Lsm1-7 complex. Additional

components of the mRNA decay machinery, including the deadenylase Ccr4 and the 5′ to

3′ exoribonuclease Xrn1 also localize within P-bodies (Fenger-Gron et al, 2005b; Sheth

& Parker, 2003; Teixeira & Parker, 2007; van Dijk et al, 2002; Yu et al, 2005). Moreover,

proteins involved in NMD, AMD, and miRNA mediated silencing machineries, as well

as mRNA decay intermediates and translationally repressed mRNAs are all found in P-

bodies (Eystathioy et al, 2003; Franks & Lykke-Andersen, 2007; Liu et al, 2005a; Sen &

Blau, 2005; Sheth & Parker, 2003; Sheth & Parker, 2006; Teixeira et al, 2005),

suggestive of a role for P bodies as sites of cytoplasmic mRNA decapping and

degradation. However several lines of evidence do not support a direct role of

microscopically visible P bodies for general mRNA decay. First, loss of detectable P-

bodies by depletion of Edc3 or Lsm4 proteins in yeast does not lead to detectable

alterations in MFA2 or PGK1 mRNA decay (Decker et al, 2007). Second, disruption of

P-bodies does not affect mRNA decay mediated by NMD or RNAi (Chu & Rana, 2006;

Eulalio et al, 2007a; Stalder & Muhlemann, 2009). Third, the decay of ARE-containing

mRNAs is not impacted by a significant reduction in the number of P-bodies following

the knockdown of GW182 in mammalian cells (Stoecklin et al, 2006). Therefore the

integrity of P-bodies does not appear to be critical for the control of general or specified

mRNA degradation. In addition, at least a subset of mRNAs can be decapped co-

translationally on ribosomes in S. cerevisiae (Hu et al, 2009). In strains lacking Xrn1p

where decapped mRNAs accumulate, decapped mRNAs are detected on polyribosomes

and 5' end decay occurs when the transcripts are associated with actively translating

ribosomes. This suggests that a fraction of Dcp2 enzyme is associated with polyribosomes and active for decapping, and indeed human Dcp2 protein co-sediments in the cytoplasmic high speed S130 pellet fraction enriched with polyribosomes (Wang et al,

2002b). These data indicate that sequestration of mRNA into a ribosome-free state is not a prerequisite for decay and the significance of cytoplasmic foci containing the decapping machinery still remains to be elucidated.

Contrary to Dcp2, the DcpS scavenger enzyme is primarily localized in the nucleus in S. pombe and human cells as determined by immunocytochemistry (Liu et al,

2004b; Salehi et al, 2002), despite its original purification from cytoplasmic extract (Liu et al, 2002). DcpS is a nucleocytoplasmic shuttling protein that can transiently enter the cytoplasm in mammalian cells in the Crm1-mediated export pathway via a leucine-rich nuclear export signal (Shen et al, 2008). Notwithstanding its well-characterized cytoplasmic function to clear the residual cap structure after 3' to 5' degradation of mRNA, the predominant nuclear localization suggests a nucleus-associated function for

DcpS. One nuclear function appears to be modulation of cap-proximal pre-mRNA splicing. Reduction of DcpS levels in cells compromises splicing of the first intron of both generic and endogenous pre-mRNAs tested (Shen et al, 2008). The proposed mechanism involves a sequestration and reduction of the nuclear cap-binding complex

(CBC) pool by unhydrolyzed cap structure, which would otherwise bind the 5' cap of a pre-mRNA and facilitating cap proximal splicing. Therefore, a major function of DcpS

could be as a modulator of cap structure concentrations in cells and potentially a regulator

of multiple processes mediated by cap-binding proteins (Bail & Kiledjian, 2008).

Cytoplasmic recapping

Addition of the N7 methyl guanosine (m7G) cap onto the 5' end of a nascent

mRNA is a well-characterized process involving three enzymatic reactions: 5'- triphosphatase, guanylyltransferase and methyl transferase (Shuman, 1995). In

mammalian cells, the first two activities are carried out by a bifunctional capping enzyme.

The 5´-triphosphatase activity hydrolyzes the 5' terminal triphosphate of the pre-mRNA

to a diphosphate and the guanylyltransferase activity covalently links a GMP to the 5'

diphosphate. Methylation at the N7 position is carried out by methyltransferase to

generate the m7G cap at the 5' end of an mRNA. Until recently, decapping was believed

to be an irreversible process that committed an mRNA for decay, as the capping enzyme

was restricted to the nucleus and could not function on 5'-monophosphate RNA generated

by Dcp2 decapping. However, this premise must be reevaluated by the recent

demonstration of a cytoplasmic capping enzyme complex that contains a kinase capable

of phosphorylating a 5'-end monophosphorylated RNA (Otsuka et al, 2009). The

resulting 5'-end diphosphorylated RNA can function as a substrate for recapping.

Therefore decapping may not necessarily always equate with the kiss of death and may

be reversible. The extent to which recapping occurs and whether the process is selective

in substrate utilization or can occur on bulk mRNA remains to be determined. However,

the demonstration that β-globin mRNAs harboring a premature stop codon generated 5'

end decay intermediates that appeared to be recapped (Lim & Maquat, 1992; Otsuka et al,

2009) and a role for cytoplasmic recapping in recovery of cells from oxidative stress

(Otsuka et al, 2009) suggests this process could be broadly utilized in cells.

Implication of the decapping enzymes in human disorders

Potential role of DcpS in Spinal Muscular Atrophy

The significance of mRNA decapping is now beginning to emerge as an important regulator of gene expression in at least two neurological disorders. The first involves a link between DcpS and spinal muscular atrophy (SMA) and the second with a potential role of Dcp2 in X-linked mental retardation. DcpS is the target cellular protein for a candidate therapeutic compound for the treatment of spinal muscular atrophy (SMA)

(Singh et al, 2008). SMA manifests in 1 out of 6000 live births and is the leading cause of hereditary infant death (Pearn, 1978). It is caused by lack of the survival motor neuron protein (SMN) due to a deletion or mutation of the SMN1 gene (Lefebvre et al, 1995;

Lorson et al, 1999). The human genome contains a second copy of the SMN1 gene,

SMN2, which primarily differs by a mutation in exon 7 resulting in approximately 90% of the SMN2 mRNAs generated to lack exon 7 and encode a truncated protein (Lorson et al, 1999) and 10% of the SMN2 pre-mRNAs spliced into an mRNA encoding wild type full length SMN protein. Therapeutic strategies aimed at increasing overall SMN2 gene expression levels could also increase the absolute levels of fully spliced SMN mRNA.

High throughput screening of small molecules identified C5-substituted quinazolines that could increase the expression of the SMN2 gene (Jarecki et al, 2005; Singh et al, 2008;

Thurmond et al, 2008). A proteomic screen with the quinazoline compound identified

DcpS as a cellular target that binds the compound. Importantly, the compound is a potent

inhibitor of DcpS decapping and the efficiency of decapping inhibition is proportional to

the level of SMN2 gene activation suggesting that the compound functions through DcpS

(Singh et al, 2008). Co-crystal structure of the compound bound to DcpS revealed that the

quinazoline compound binds and traps the DcpS dimer in an inactive conformation

(Singh et al, 2008). The mechanism by which DcpS inhibition mediates an increase of

SMN2 expression is not yet clear. Possible mechanisms include an indirect consequence

of decapping inhibition leading to cap structure accumulation that in turn influences cap-

binding protein functions. However, a novel decapping independent function of DcpS

cannot be ruled out.

Potential role of Dcp2 in X-linked mental retardation

The VCX-A gene was identified as the defining interval in a subset of patients

with X-linked mental retardation (Fukami et al, 2000) and the VCX-A protein was

recently shown to be an inhibitor of decapping (Jiao et al, 2006) and mRNA translation

(Jiao et al, 2009). VCX-A is a member of a highly related protein family VCX/Y, consisting of four distinct genes on the X chromosome and two identical genes on the Y chromosome (Fukami et al, 2000; Lahn & Page, 2000). Expression of the VCY genes is

testes-restricted while the VCX genes are ubiquitously expressed (Jiao et al, 2009). The

VCX/Y gene family are simian primate-restricted and do not have an obvious ortholog in

lower mammals (Fukami et al, 2000; Lahn & Page, 2000). Although VCX-A was

specifically implicated in mental retardation, the role of the VCX proteins in cognitive

function is likely more complex and may involve a cumulative role of the multiple VCX

genes and additional modifier genes rather than only the VCX-A gene (Cuevas-

Covarrubias & Gonzalez-Huerta, 2008; Hosomi et al, 2007; Macarov et al, 2007; Van

Esch et al, 2005). VCX-A functions as an inhibitor of Dcp2 decapping, at least in part by

its cap binding capacity to prevent Dcp2 from accessing the cap (Jiao et al, 2006).

Importantly, the binding affinity of VCX-A to the cap increases upon association with

Dcp2, in a mechanism to restrict access of the cap to Dcp2 and further ensure the

integrity of the mRNA 5′ in the presence of the decapping enzyme. The mRNA 5' end

binding property of VCX-A also promotes inhibition of mRNA translation most likely by

competing with the eIF4E cap binding protein for cap access (Jiao et al, 2009).

The functional significance for the VCX-A mediated inhibition of decapping and

translation appears to manifest in the regulation of neuronal projections. Expression of

VCX-A in rat hippocampal neurons, which normally lack the VCX/Y genes, leads to

increased neurite numbers. Conversely, a knockdown of the VCX genes in human

neuroblastoma cells results in a decrease of neuritogenesis (Jiao et al, 2009). Moreover,

VCX-A can preferentially bind to a subset of mRNAs as well as aggregate into staufen-

containing neuronal granules within neurites of human neuroblastoma cells (Jiao et al,

2009). Therefore, VCX-A appears to function by sequestering and transporting specific

mRNAs in a Dcp2 resistant and translationally silenced state in neurons to modulate

neuritogenesis. These findings provide a potential link between VCX-A, and in turn

decapping and translational suppression, to cognitive function.

Anti-viral innate immunity

Virus infection activates the expression of a group of genes encoding antiviral proteins that trigger first the innate immune response and subsequently the adaptive

immune response. The interferon (IFN) system is the first line of defense against viral

infection in mammals. They belong to a class of important biological regulatory proteins

called cytokines. Thus, IFNs are the mediators of innate immune responses to virus

infection in mammalian cells: they are induced in response to viruses, secreted to circulation and act on yet uninfected cells to activate a global antiviral state in which virus replication is inhibited. Interferons are divided into two types, type I and type II, which function through different receptors and are structurally unrelated. There are many members of the type I or IFN-α/β superfamily but only one member of the type II family,

IFN-γ. Type I interferons, which are the major mediators of anti-viral response, consist of multiple numbers of IFN-α genes and only one IFN-β gene in mammals. They exhibit profound pleiotropic effects on many aspects of cellular functions, mediating antiproliferative, antitumor, and antibacterial activities besides their primary role of

antiviral cytokines. Investigation of the biochemical mechanisms responsible for IFNs’

effects has led to the discovery of hundreds of IFN-stimulated genes.

Type I IFNs can be induced in most cell types, and are the major initial response

to many viral infections of mammalian hosts. The transcriptional activation of IFN-α/β

genes are mediated by complex signaling networks involving many transcription factors.

A general principle has emerged that receptors recognizing viral nucleic acids such as

single-stranded (ss)RNA, doublestranded (ds)RNA or dsDNA accounts for most virus

detection (Fensterl & Sen, 2009). A well studied pathway is triggered by double-stranded

RNA, a common feature of both DNA and RNA viruses that’s recognized by the immune

system. Receptors of dsRNA are RIG-I (retinoic acid-induced gene-I) and MDA-5

(melanoma differentiation-associated gene-5) in the cytoplasm (Andrejeva et al, 2004;

Fensterl & Sen, 2009; Yoneyama et al, 2004), or TLR3 (Toll-like receptor 3) in the endosome (Alexopoulou et al, 2001). Upon binding to the dsRNA, these receptors can intiate signaling cascades, either via a TRAF3-containing branch or TRAF6-containing branch, to eventually phosphorylate critical transcription factors in the activation of type I

IFNs, including IRF-3, IRF-7, NF-κB, ATF-2, and c-JUN (Fitzgerald et al, 2003; Saha et al, 2006; Seth et al, 2005; Sharma et al, 2003; Xu et al, 2005).These factors then activate the transcription of type I IFNs as well as other cytokine/chemokine genes, such as

CXCL10 and CCL5 (Andersen et al, 2008). IRF3 and IRF7 are members of a family of transcription factors called IFN regulatory factors (IRF) that share homology in the amino-terminal 115 amino acids. Altogether, nine cellular IRFs have been identified, and they exert distinct roles in diverse biological processes, such as response to pathogens, cytokine signaling and cell growth regulation (Nguyen et al, 1997). IRF3 and IRF7 are crucial factors mediating induction of type I IFNs in viral infection. IRF-3 is constitutively expressed in cells while IRF7 is usually induced in response to the initial infection in many cell types. Most of IFN-α subtypes require the induced IRF-7, whereas

IFN-β can be induced without IRF-7 by the constitutively expressed IRF-3 (Honda et al,

2006). In mouse fibroblasts, IFN-β expression is required for IFN-α production by inducing IRF-7; however in leukocytes IFN-α can be induced without IFN-β (Erlandsson et.al, 1998).

Type I IFNs activate downstream genes by binding to the cell surface heterodimeric receptor IFNAR1/IFNAR2c, whose cytoplasmic domains recruit specific

protein tyrosine kinases that get activated upon IFNs binding. The kinases involved in

this process are members of the Janus kinase (Jaks) family including JAK1 and TYK2.

These kinases then phosphorylate transcription factors STAT1 and STAT2 (signal transducers and activators of transcription). Activated STATs form heterodimers, associate with IRF-9, and then translocate to the nucleus, bind to specific DNA sequences on the promoter of ISGs (interferon-stimulated genes) to activate transcription (Fensterl

& Sen, 2009). Numerous ISGs are induced by IFNs to help eliminate viruses or build an

“antiviral state”. These proteins include enzymes, transcription factors, chemokines, signaling proteins and apoptotic factors. One well-characterized IFN-induced protein is the dsRNA-activated protein kinase, PKR. It’s activated upon binding to dsRNA, which induces its dimerization and autophosphorylation (Clemens & Elia, 1997; Proud, 1995).

Activated PKR can phosphorylate selected cellular target proteins, one of which is the translation initiation factor eIF-2α. Phosphorylation of eIF-2α causes an inhibition of translation which blocks cellular and viral protein synthesis (Samuel, 1979; Samuel,

1993). Another group of IFN-induced proteins are the 2'-5' oligoadenylate synthetases

(OAS). These proteins, activated by dsRNA, generate 2'-5' linked oligoadenylate from

ATP, which serves as the activator of a latent ribonuclease, RNase L, by inducing its dimerization (Sadler & Williams, 2008; Silverman, 2007). The RNaseL cleaves cellular and viral single stranded RNA and thereby inhibits protein synthesis after viral infection

(Sadler & Williams, 2008; Silverman, 2007).

Because of its antiviral and immunomodulatory effects, recombinant interferons have been used as therapeutics to a number of viral and nonviral diseases. IFNα/β is clinically administered to treat chronic active hepatitis caused by infection of HCV or

HBV. IFN-β is commonly used in treatment of multiple sclerosis (MS) although the mechanism of its beneficial effects is largely unknown (Friedman, 2008). The inhibitory effects on cell growth and apoptotic response by IFNs have been observed on certain types of tumor cells, and this antigrowth property of IFNs have been exploited in their clinic management to treat several types of cancer such as certain leukemias, lymphoma and malignant melanoma (Friedman, 2008).

Materials and Methods

Plasmid constructs

Plasmids used in characterization of DBDE

The plasmid pGEM-Rrp41 containing human Rrp41 open reading frame was

constructed by inserting the Rrp41 cDNA containing a T7 promoter sequence at the 5´

end into the pGEM-T vector (Promega; Madison, WI). The cDNA was obtained by reverse transcription of 293T cell RNA and PCR amplified with primers 5'-

TAATACGACTCACTATAGGGACCTCCGGAAACCGTAGATTCC-3' and 5'-

GTGGGCAGAGGAGGGTTTTATTC-3'. pGEM-Stx7 was similarly constructed with primers 5'-TAATACGACTCACTATAGGGGTGACTGCTTAGAAAACTGC-3' and 5'-

GTAAACATCAGACTTAAATAGACC -3' to amplify the Stx7 cDNA.

The pGEM-R5´UTR-S which contains the chimeric sequence of Rrp41 5′UTR and the Stx7 coding region (Stx7∆5′UTR) was constructed in two steps. Initially, the Stx7

coding region was amplified with a primer to insert a 5′ BamHI site (primers 5'-

GCGGATCCATGTCTTACACTCCAGGAGTTG-3' and 5'-GTAAACATCAGACTTA

AATAGACC -3' ) and inserted into the pGEM-Teasy vector. The Rrp41 5′UTR was

generated with primers 5'-CAGTCCATGGACCTCCGGAAACCGTAGATTCC-3' and

5'- CAGTAGGATCCGCTGCCCGGCCGCCAGGTCC -3' and inserted into the NcoI

and BamHI sites of pGEM- S∆5′UTR plasmid. The plasmid containing the Rrp41 5′UTR

terminal 60 nucleotides 1-60 (pGEM-R5´1-60-S) was generated by digesting pGEM-

R5´UTR-S with SacII and BamHI and self ligating.

The plasmids for GST-Dcp2 (pGEX6p1-Dcp2; (Wang et al, 2002b)), GST-mDazl

(pGEX6p1-mDazl; (Jiao et al, 2002)) and pcDNA3-Flag-Dcp2 (Fenger-Gron et al, 2005b) were constructed as described previously. pcDNA3-Flag-mDazl was constructed by removing Dcp2 cDNA from pcDNA3-Flag-Dcp2 plasmid and replacing it with the mDazl coding region.

Plasmids used in study of differential utilization of decapping enzymes

hNudt16 cDNA was excised from pET28a-hNudt16 plasmid with BamH1 and

HindIII and inserted into the same sites in pCDNA3-FLAG to generate pcDNA3-FLAG-

hNudt16. pcDNA3-hNudt16-FLAG was constructed similarly using primers 5′-

GATCAAGCTTGCCGCCATGGCCGGAGCCCGCAGGCT-3′ and 5′-GATCG

CGGCCGC TCACTTGTCATCGTCCTTGTAGTCGTGATGAGCTGGAATCTTAA-

3′.to amplify hNudt16-FLAG fragment first.

The pmCMV-G1 Norm and pmCMV-G1 39Ter NMD reporter plasmids that

encode the normal or nonsense mutation-containing versions of the β-globin transcripts

respectively were kindly provided by Dr. L.E. Maquat (University of Rochester). The β-

495 and β-496 NMD reporter plasmids that encode the normal or nonsense mutation-

containing versions of the TCR-β transcripts respectively were provided by Dr. Miles

Wilkinson (UCSD)(Wang et al, 2002a). The microRNA reporter plasmids encoding the

Renilla luciferase (pCMV-RL), Renilla luciferase with three let-7 imperfectly basepaired

target sites in the 3´UTR (pCMV-RL-3Xbulge), the Renilla luciferase with a

complementary let-7 target site in the 3´UTR (pCMV-RL-perfect) and the pri-let7

expression plasmid (pcDNA3-priLet7) were kindly provided by W. Filipowicz (Friedrich

Miescher Institute for Biomedical Research)(Schmitter et al, 2006). The pRL-SV40

plasmid encoding the Renilla luciferase reporter protein was obtained from Promega

Corp. (Madison, MI) and the pSV2AL 5'-Luc plasmid expressing the Firefly luciferase

gene has previously been described (de Wet et al, 1987).

The plasmids pLKO.1-puro-mNudt16 expressing shRNA against Nudt16 and

pLKO.1-puro-mUpf1 expressing shRNA directed against Upf1 were obtained from

Sigma-Aldrich (St. Louis, MO). To construct the retroviral plasmid pBMN-Nudt16mut which overexpresses a shRNA-resistant Nudt16 mutant, pGEM-Nudt16 was used as template with the following PCR primers 5′-CTCGACCCAGGGATCATAG

CCAAACTAAAGATCCCAGATTCTAAGTAGAATCGGAC-3′ and 5′-GGCTATG

ATCCCTGGGTCGAGAAGTTTCAAGTCCTGAAGGGCTTCTAG-3′ to generate pGEM-Nudt16mut. pGEM-Nudt16mut was subsequently used as template with primers

5′-CAGCTCGAGACCATGGAGGGGCATCGGAAAGTG-3′ and 5′-ACAGCGGC

CGCGTCCATGCTGGTCCGATTCTAC-3′. The PCR product was purified and digested by Xho I and Not I and inserted into pBMN-I-GFP (Addgene, Cambridge, MA).

Plasmids used in study of decapping in anti-viral immunity

The pLKO.1-puro, pCMV-VSV-G and pSPAX2 plasmids used in the generation of lentiviral particles were purchased from Sigma-Aldrich (St. Louis, MO). To construct the retroviral plasmids pBMN-Dcp2 and pBMN-Dcp2 EE/Q which expresses wild type or cayalytic mutant Dcp2 proteins respectively, pET-Dcp2 and pET-Dcp2 EE/Q plasmids

(Wang et al, 2002b) were used as template with the following PCR primers 5′-

CAGTCTCGAGATGGAGACCAAACGGGTGG-3′ and 5′-CAGTGCGGCCGCTC

AAAGGTCCAAGATTTT-3′. The PCR products were purified and digested by Xho I and Not I and inserted into pBMN-I-GFP (Addgene, Cambridge, MA).

Generation of RNA in vitro

RNAs were in vitro transcribed from PCR-generated templates that contained T7 or Sp6 promoter sequences at the 5′ end. Cap labeling was carried out as previously

described (Wang & Kiledjian, 2001). The 5′ 900 nucleotides of Stx7 mRNA used as negative control for decapping assays was transcribed with T7 polymerase from a template generated with primers 1 and 2 (all primer sequences are listed in Table III).

Rrp41 mRNA truncations in Figure 3A were generated with T7 polymerase from templates generated by the following primer sets: fragment A was generated with

primers 3 and 4; fragment B with primers 5 and 6; C with primers 8 and 7; D with

primers 3 and 9; and E with primers 10 and 4. Template for RNAs in Figure 5A were

transcribed with T7 polymerase using the same 3′ primer (primer 2) and the following

different 5′ primers: 1-110 and 50-110 were made from pGEM-R5′UTR-S with 5′ primers

3 and 11 respectively; 1-60 was made from pGEM-R5´1-60-S with 5′ primers 3; RNA

probe used in the electrophoretic mobility shift assays in Figure 5D was in vitro

transcribed in the presence of [α32P]GTP from template generated by primers 16 and 17.

2xDE RNA mutations used in decapping assays were transcribed as chimeric

RNAs with the coding region of the Stx7 RNA attached to the 3′ end (in order to

maintain a uniform size for all the RNAs) by T7 RNA polymerase from templates

generated by 3′ primer 34 and the following 5′ primers: for Figure 7A, Mut1-10 was

generated with primer 18; Mut11-22 with primer 19; Mut23-33 with primer 20; Mut34-

44 with primer 21; and Mut45-55 with primer 22. For Figures 9 and 10, mutant A was

generated with primer 23; B with primer 24; C with primer 25; D with primer 26; E with

primer 27; F with primer 28; G with primer 29; H with primer 30; and I with primer 31.

The RNAs in Figure 11A, 10-2xDE and 20-2xDE were generated with 5´ primers 32 and

33 respectively and the same 3´ primer, 34. Templates for Ndufb7 mutant RNAs in

Figure 12 were transcribed with T7 RNA polymerase using the same 3′ primer (primer 35) and the following different 5′ primers: wild type with primer 36; Mut35-44 with primer

37; and Mut2-11/35-44 with primer 38. For Figure 13A, Hip1 RNA was generated with primer 39 and 40; Kcnj2 with primers 41 and 42; Zcrb1 with primers 43 and 44. Mutants of Hip1 RNA in Figure 13B were generated with 3′ primer 40 and the following 5′ primers: Mut23-31 with primer 45 and Mut4-13/23-31 with primer 46. RNA probes used in the electrophoretic mobility shift assays were in vitro transcribed in the presence of [α32P]GTP from template generated by the following primer sets: for Figure 7B, wild

type 2xDE was generated with primers 47 and 48; Mut1-10 with primers 18 and 48;

Mut11-22 with primers 19 and 48; Mut23-33 with primers 20 and 49; Mut34-44 with

primers 21 and 50; Mut45-55 with primers 22 and 51; Stx7 RNA with primers 1 and 52.

For Figure 11B, 10-2xDE was generated with primers 32 and 52; 20-2xDE with primers

33 and 52. All primer sequences are listed in Table III.

Coimmunopurification of RNAs associated with Dcp2

pcDNA3-Flag-Dcp2 (16µg) or pcDNA3-Flag-mDazl (7µg) plasmids were

transfected into 293T cells in 10-cm dishes by using Lipofectamine 2000 (Invitrogen;

Carlsbad, CA) following recommendations of manufacturer. These concentrations were

chosen to yield equivalent levels of expressed Flag-tagged protein in the cells. Twenty

four hours after transfection, cells were harvested in 1 ml lysis buffer (PBS+0.5%

TritonX-100) and disrupted by sonication. After a brief spin at 10,000g for 10 min, the amount of supernatant used in the subsequent binding steps was adjusted such that an equivalent amount of Flag-tagged protein was used in the binding studies. The supernatant was precleared with 100µl agarose-IgG (mouse) beads for 30 min to remove non-specific binding to the beads. The precleared supernatant was incubated with 150µl

o anti-Flag M2 beads at 4 C for 2h, with addition of MgOAc2 and MnCl2 to a final

concentration of 2mM and 0.5mM respectively. Beads were then washed 4 times with

washing buffer (PBS+150mM NaCl+300mM urea +0.5% TritonX-100). RNAs were isolated from the beads by boiling for 3 min in 300µl TE/1% SDS and purified by

RNeasy Mini Kit (Qiagen; Valencia, CA)

Microarray Analysis

Microarray of RNAs bound by Dcp2

RNA quality was assessed on an RNA chip using an Agilent 2100 Bioanalyzer

(Agilent Technologies; Palo Alto, CA), and microarray experiments were carried out at

the Transcriptional Profiling Facility, Cancer Institute of New Jersey. RNA samples

were labeled using BioArray™ HighYield™ RNA Transcript Labeling Kit (T7) (Enzo

Life Sciences, Inc.; Farmingdale, NY) and hybridized to Human Genome U133A 2.0

GeneChip® (Affymetrix Inc.; Santa Clara, CA) following manufacturers’

recommendations. Arrays were then washed and stained with Streptavidin–phycoerythrin conjugate (Invitrogen Corp.; Carlsbad, CA) using an Affymetrix Fluidics Station 450 and scanned on a GeneChip scanner (Affymetrix Inc.; Santa Clara, CA). The data were processed using the Microarray Suite 5.0 software (MAS 5.0, Affymetrix, Inc.) and gene annotations were referenced to the Gene Ontology database.

Microarray of RNAs in MEF cells

Wild type and Dcp2β/β MEF were infected with lentiviruses expressing control

vector or shRNA against Nudt16. Total RNAs were harvested 2 days post-infection by

TRIzol reagent (Invitrogen; Carlsbad, CA) following the manufacturer’s instructions.

RNA quality was assessed on an RNA chip using an Agilent 2100 Bioanalyzer (Agilent

Technologies; Palo Alto, CA). The mRNA was amplified with TotalPrep RNA amplification kit with T7-oligo(dT) primer according to the manufacturer’s instructions

(Ambion) and microarray analysis was carried out with the Illumina Sentrix MouseRef-8

24K Array at The Burnham Institute (La Jolla, CA).

RT-PCR

RNAs were reverse transcribed by M-MLV reverse transcriptase (Invitrogen;

Carlsbad, CA) with random primer according to manufacturer’s instruction. PCR

amplifications were carried out with gene-specific primers. Primers used in Figure 1A for

Spen, Rrp41, Ndufb7, Ndufs8, Psmc3, Ruvb2, Tceb2, Ada, Ap2s1, Gltscr, Edf1, Narf,

Hmox2, Ppib, and Stx7 are listed in Table I.

GST fusion protein copurification

The GST-Dcp2 and GST-mDazl fusion proteins were expressed in Escherichia

coli BL21 as previously described (Jiao et al, 2002; Wang et al, 2002b). 4µg of GST-

fusion protein was bound to 50µl GST-beads in a total volume of 400µl in PBS with

protease inhibitor cocktail (Roche; Mannheim, Germany) at 4oC for 1h, followed by

extensive washes in 300mM NaCl/500mM urea/0.5% TritonX-100. The washed beads

were resuspended in 400µl of RBB (10 mM Tris-HCl, pH 7.5, 1.5 mM MgCl2, 150 mM

KCl, 0.5mM DTT) with RNase inhibitor (Promega; Madison, WI ), and incubated with in vitro transcribed [α32P]UTP labeled RNA at 4oC for 1 h. After washes in RBB+150mM

NaCl +300mM urea+0.25% TritonX-100, RNAs were isolated from the beads as

previously described (Trifillis et al, 1999). Copurified RNA was resolved on a 5%

polyacrylamide gel with 7M urea.

In vitro RNA decapping assay

In vitro RNA decapping assays were carried out essentially as described

previously (Wang et al, 1999). Reactions were carried out in 20µl reaction volume with

25-100 ng of bacterial expressed recombinant Dcp2 or indicated proteins incubated with

cap-labeled RNA (2,000 cpm) at 37°C for 30 min in IVDA-2 buffer (10 mM Tris:HCl at

pH 7.5, 100 mM potassium acetate, 2 mM magnesium acetate, 0.5 mM MnCl2 and 2 mM

dithiothreitol). An aliquot of each sample was resolved by phosphoethyleneimine-thin

layer chromatography (PEI-TLC) developed in 0.45 M (NH4)2SO4 and exposed to

PhosphorImager. Quantifications were carried out using a Molecular Dynamics

PhosphorImager using ImageQuante-5 software. Percent decapping was determined as

the level of m7GDP relative to total RNA used in the reaction

Electrophoretic mobility shift assays

Electrophoretic mobility shift assays were carried out with in vitro transcribed

[α32P]GTP uniformly labeled RNA substrate ( 4,000 cpm per reaction). Binding

reactions were carried out in RBB (10 mM Tris [pH 7.5], 50 mM KCl 1.5 mM MgCl2, 0.5

mM DTT ) with 1 or 2µg of His-Dcp2 protein in a 20-µl total volume containing 1.5 µg

yeast RNA and 0.25 µg heparin. Following a 25-min binding reaction at room temperature, the complexes were resolved on a 5% polyacrylamide gel (60:1

acrylamide:bis) in 0.5x Tris-borate-EDTA buffer at 8 V/cm and exposed to

PhosphorImager.

RNase mapping

2xDE RNA was in vitro transcribed and incubated with Calf Intestinal

Phosphatase (CIP) at 370C for 1 hour to dephosphorylate the 5′ end. A phenol/chloroform

(1:1) extraction was carried out following the reaction to remove CIP. The

dephosphorylated RNA was incubated with [γ-32P] ATP and T4 Polynucleotide Kinase

(Ambion; Austin, TX) to label the 5′ end following the manufacturer’s protocol. The 5′

end labeled RNA (1500 cpm) was incubated with the indicated amounts of RNase A,

RNase T1, RNase V1 (Ambion; Austin, TX) or alkaline hydrolysis buffer (Ambion;

Austin, TX) following the manufacturer’s instructions and the digested products were resolved on a 15% polyacrylamide gel and exposed to PhosphorImager.

Bioinformatics search for Dcp2 substrates

The PatSearch program and UTR database provided by this link

(http://www.ba.itb.cnr.it/BIG/PatSearch/) was used to search available human 5’ UTR

sequences (Grillo et al, 2003). The stem-loop secondary structure constraint criteria consisted of an 8-15 nucleotide stem portion with either perfectly paired or 1 mismatch with no bulge or a one-base bulge with no mismatch, and a 6-15 nucleotide loop. G-U

base pairing was also permitted in addition to Watson-Crick base pairing. The following

pattern was submitted to the PatSearch website: r1={at,ta,gc,cg,gt,tg} (p1=8...15 6...15

~p1[1,0,0] | p2=8...15 6...15 ~p2[0,1,1]). Human 5´ UTRs that adhered to the search

criteria were returned with a proprietary reference ID. A Python script was developed to

match and identify each reference ID to its gene description in the UTRdb from

ftp://bighost.ba.itb.cnr.it/pub/Embnet/Database/UTR/.

siRNA transfection

293T cells (30% confluent) were cultured in 6-well plates and transfections were

carried out with 80nM Dcp2 siRNA (Dharmacon SMARTpool M00842500; Chicago, IL)

or control siRNA (Qiagen 1027281; Valencia, CA) by using Lipofectamine 2000 reagent

(Invitrogen Corp.; Carlsbad, CA) following recommendations of the manufacturer. A

second transfection was carried out 24 hours following the first.

Cell based mRNA decay assays

293T or MEF cells were treated with actinomycin D (5µg/ml) to stop

transcription. Cells were harvested at the indicated times post-actinomycin D addition

and total RNA was isolated by TRIzol reagent (Invitrogen; Carlsbad, CA) following the

manufacturer’s instructions. RNAs were reverse transcribed by M-MLV reverse transcriptase (Invitrogen; Carlsbad, CA) with random primers according to the manufacturer’s instruction. mRNA levels were quantified by real-time PCR using SYBR green PCR core reagent (Applied Biosystems, Foster City, CA), and the abundance of

specific mRNAs were quantified using the standard curve method according to the

recommendations of the manufacturer. Values were normalized to the stable U6 or β-

actin RNA. Each mRNA was amplified using the appropriate specific primers. Primers used for real-time PCR are listed in Table II. Real-time PCR was carried out with an ABI

Prism 7900HT sequence detection system, and the specificity of the amplified PCR

products was assessed by a melting curve analysis after the last cycle by the

manufacturer's suggested program

Electroporation of RNA

Cap labeled RNA (2 × 105 cpm) was electroporated into 293T cells (2× 106) in a

total volume of 200 μl OPTI-MEM media in a 4 mm gap cuvette using a BioRad

Genepulser charged at 300mV, 250μF. All reagents were kept on ice prior to

electroporation. Following discharge, the cells were resuspended in culture media

containing 200 U/ml micrococcal nuclease (USB Corporation, Cleveland, Ohio) and 1

o mM CaCl2 for 10 min at 37 C to degrade residual RNA remaining outside the cell.

Following the incubation, cells were transferred to culture media and an aliquot was removed for time zero; subsequent aliquots were isolated at the indicated time points.

RNA was isolated using Trizol Reagent (Invitrogen, Carlsbad, California) and resolved by denaturing polyacrylamide gel electrophoresis.

Reporter assays

Transfections were performed in 6-well plates using Lipofectamine 2000 reagent

(Invitrogen Corp.; Carlsbad, CA) according to the manufacturers recommendations. For

the NMD reporter assays, transfections were carried out with 1µg of pmCMV-G1-Norm

or pmCMV-G1-39Ter plasmid or 0.5µg of β-495 or β-496 plasmid, along with 1µg pRL-

SV40 plasmid. Total RNA was isolated 24h post-transfection, β-globin or TCR-β mRNA

levels analyzed by real-time reverse transcription-PCR as described above, and

normalized to Renilla luciferase mRNA levels. Analysis of microRNA silencing was

carried out with 0.3µg of pCMV-RL or pCMV-RL-3Xbulge or pCMV-RL-perfect

plasmids transfected into MEF cells in 6 well plates together with 1.5µg pcDNA3-priLet7

and 1µg pSV2AL 5'-Luc plasmids. Nudt16 complementation studies were carried out

with the transduction pBMN-Nudt16mut-derived retrovirus according to manufacturer’s

instruction to express an shRNA resistant Nudt16 24h prior to transfection of the dual

luciferase reporter construct. Renilla and Firefly luciferase activities were detected using

a Dual Luciferase Assay System (Promega, Madison, WI) 24h post-transfection according to the manufacturer’s guidelines using GloMax®-Multi Luminescence Module

(Promega, Madison, WI).

Western Blot

Cells were sonicated in PBS and protein extract was resolved by 12.5% SDS-

PAGE. Nudt16 polyclonal antibody (1:100 dilution; (Song et al, 2010), monoclonal anti- eIF4E antibody (1:1500 dilution; Transduction Laboratories, Lexington, KY), affinity purified Dcp2 polyclonal antibody (1:500 dilution) (Wang et al, 2002b), IRF7 polyclonal antibody (1:500 dilution; Sigma-Alderich) and monoclonal anti-GAPDH antibody

(1:2000 dilution; Abcam Inc., Cambridge, MA), were used for Western blot analysis and visualized using secondary antibodies coupled to horse radish peroxidase (Jackson

ImmunoResearch, West Grove, PA) and chemiluminescence (ECL; GE Healthcare Life

Science, NJ).

Immunofluorescence Assay

To study the localization of Nudt16 protein, U2OS cells were transfected with

pCDNA3-Flag-hNudt16 or pCDNA3-hNudt16-Flag plasmids using lipofectamine 2000

(Invitrogen) according to the manufacturer's protocol. Flag tagged proteins were detected

by indirect immunofluorescence with 1:200 dilution of anti-FLAG antibody (Sigma) and

1:200 dilution of FITC-conjugated anti-mouse secondary antibody. Nuclei were stained

for 3 min with DAPI (1 µg/ml). Images were captured from a LSM510 META confocol

microscopy (Carl Ziess, Thornwood, NY).

To visualize P-bodies in Dcp2 or Nudt16 knockdown cells, MEF cells were

grown on glass coverslips in six-well plates to 50% confluency. The coverslips containing the cells were treated as previously described (Liu et al, 2004b). Affinity-

purified Dcp2 rabbit polyclonal antibody was used as the primary antibody to detect

endogenous Dcp2 and monoclonal Dcp1a antibody (1:200 dilution; Abnova, Walnut, CA)

was used to detect Dcp1a containing P-bodies. Goat anti-rabbit or goat anti-mouse secondary antibodies conjugated with Texas Red or FITC (Jackson ImmunoResearch,

West Grove, PA) were used at a 1:200 dilution. The nuclei were stained with 1µg/ml

DAPI for 10 minutes. The images were obtained with a Zeiss Axiovert 100 M microscope. P-body numbers per cell were counted and the average number of 30 cells was graphed.

Lentivirus production and infection

Lentiviral particle were produced in 293T cells with cotransfection of pLKO.1-

puro empty vector or pLKO.1-puro-mNudt16, pCMV-VSV-G and pSPAX2 plasmids at a

ratio of 4:1:3. Cells were washed and supplemented with fresh media 24 hours post- transfection and the supernatant containing viral particles harvested following an additional 24 hours. MEF cells were infected with the viral particles in the presence of

8µg/ml hexadimethrime bromide, washed and fed with fresh medium 24 hours later.

Cells were harvested for Western Blot analyses or real-time PCR analysis at indicated times post-infection.

Table I: Primers used in semi-quantitative RT-PCR

primer name Primer sequence Rrp41F TACATTGAGCAGGGCAACAC Rrp41R ATGGCTGCTTCGAAAGTCTG SpenF TCC TCA GTA TGC GTT TCT GC SpenR ATC TAG CCA CAC GCA GTT TG Ndufb7F CACACAGCAGGAGATGATGG Ndufb7R AACTCTGCCGCCTTCTTCTC Ndufs8F ATCCCGAGATGGACATGAAG Ndufs8R CCGTTGTTGAGCAACTTCTC Psmc3F ACAGACGTACTTCCTTCC Psmc3R CCAATGAACATCTGCACCAG Ruvb2F GCGAGGAAGAAGATGTGGAG Ruvb2R CAACTCAGGAGGTGTCCATG Tceb2F TGGCTTCACCAGTCAAACAG Tceb2R AACAATGGCTTGGGTCTCAG AdaF GTGTAAGAAGTACCAGCAGC AdaR TTGAGCCGAATGACTGCATG Ap2s1F AACTGGACCTGGTGTTCAAC Ap2s1R AGACGCTTATGGCATCACAC GltscrF TCATCCTCGAGAACACATCC GltscrR TGGTGGTCTTCAAAGGATGG Edf1F TGCTGGCCAGAACAAACAAC Edf1R TTGTGTTCATTTCGCCCTAG NarfF AGCTGTTCAACGAGGATGTG NarfR TTAAGCATCCTCCAGCACAG Hmox2F ATGAGAATGGCTGACCTCTC Hmox2R TACTCCATGTCCTTGGTCAG PpibF CAAGGACTTCATGATCCAGG PpibR CAGTCTGCGATGATCACATC Stx7F GCTCGAGTAAGAGCCAGTTC Stx7R AAGACGGAGGTCATCCTCTG

Table II. Primers used for real-time PCR

Gene/primer name Primer sequence

Rellina luciferase 5′-CAGTGGTGGGCCAGATGTAAACAA-3′ 5′-TAATACACCGCGCTACTGGCTCAA-3′ β-globin 5′-ACCACCGTAGAACGCAGATCG-3′ 5′-GGGTTTAGTGGTACTTGTGAGC-3′ TCRβ 5′-TATGTCGCTGACAGCACGGAGAAA-3′ 5′-AGCTGTCTGAGAAAGGGAAGCCAA-3′ β-actin 5′-TCTCCTTCTGCATCCTGTCAGCAA-3′ 5′-TCTTGGGTATGGAATCCTGTGGCA-3′ c-Fos 5′-AGAGAAACGGAGAATCCGAAGGGA-3′ 5′- ATTGGCAATCTCAGTCTGCAACGC-3′ c-Jun 5′-TGAAGTTGCTGAGGTTGGCGTAGA-3′ 5′- GAACTGCATAGCCAGAACACGCTT-3′ c-Myc 5′-AATCCTGTACCTCGTCCGATTCCA-3′ 5′-TTGCTCTTCTTCAGAGTCGCTGCT-3′ IFNα2 5′- TGAAGGACAGGCAGGACTTTGGAT-3′ 5′- AGGAGGGTTGTATTCCAAGCAGCA-3′ IL4 5′-GGCTTCCAAGGTGCTTCGCATATT-3′ 5′-GCAGCTTATCGATGAATCCAGGCA-3′ P53 5′- ACAAGAAGTCACAGCACATGACGG-3′ 5′- TTCCTTCCACCCGGATAAGATGCT-3′ Dcp2 5′-ACTGCGAATCAATGACCAGCTTGC-3′ 5′-AGCTTGGACTTGGGAGTCATGTCA-3′ GAPDH 5′-ACCATGGAGAAGGCTGGGGC-3′ 5′-TGGACTGTGGTCATGAGTCC-3′ Firefly luciferase 5′-CTCACTGAGACTACATCAGC-3′ 5′-TCCAGATCCACAACCTTCGC-3′ Rrp41 5′-AGACTTTCGAAGCAGCCATCCTCA-3′ 5′-ATCTGCCTGTAGCACCTGCACATA-3′ Stx7 5′-AAGCAAATCAGCTGTCAAGGG-3′ 5′-ATGTAGTGCGACAGTGTGCTCCTT-3′ U6 5′-TCGCTTCGGCAGCACATATA-3′ 5′-CACGAATTTGCGTGTCATCCTTGC-3′

Table III. Primers used for generating RNAs in vitro

primer Sequence 1 TAATACGACTCACTATAGGGGTGACTGCTTAGAAA ACTGC 2 CATGATTACAGGAATCTTCCTAC 3 TAATACGACTCACTATAGGGACCTCCGGAAACCGTAGATTCC 4 CAAGCTGCATAGGTCCCACCATC 5 TAATA CGACT CACTA TAGGG TGTGAATGCAGCCACGC 6 GTGGGCAGAGGAGGGTTTTATTC 7 GCGCA ATCTG TCCTG AGGCT G 8 TAATA CGACT CACTA TAGGG GCCTGCAGCTCCGCCAG 9 GCTGCCCGGCCGCCAGGTCC 10 TAATA CGACT CACTA TAGGG ATGGCGGGGCTGGAGCTC 11 TAATA CGACT CACTA TAGGGTCCCGCGGCTCAGAGAAG 12 TAATA CGACT CACTA TAGGG CGGTCGGAGCCGCCGGGA 13 ACCTCCGGAAACCGTAGATTCCGGGATGTCTTACACTCCAGG 14 ATTTAGGTGACACTATAGAATACCGCACAGATGCGTAAG 15 CAACTGTTGCCTCAATTCAGGTG 16 TAATA CGACT CACTA TAGGGACCTCCGGAAACCGTAGATTCCGGGCGGTCGG 17 TTCTCTGAGCCGCGGGAGAACTACGCTCCCGGCGGCTCCGACCGCCCGGAAT C 18 TAATACGACTCACTATAGGGCAAGAATTCCACCGTAGATTCCGGGCGGTCG 19 TAATACGACTCACTATAGGGACCTCCGGAACAATGCTCGGAAGGGCGGTCGG AGCCGCCG 20 ACCTCCGGAAACCGTAGATTCCTTTATTGATTCGCCGCCGGGAGCTGTAGTTC 21 TCCGGAAACCGTAGATTCCGGGCGGTCGGATAATCCTTTCTCTGTAGTTCTCC CGATCC 22 GTAGATTCCGGGCGGTCGGAGCCGCCGGGAGAGTGCTGGAGCCCGTCCATGT CTTACAC 23 TAATACGACTCACTATAGGGAAATAATTAAACCGTAGATTCCGTAGATTCCGGG CGGTCG 24 TAATACGACTCACTATAGGGACCTCCGGAAACCGTAGATTAATTGCTTTCGGA GCCGCCGGGAGCTGTAG 25 TAATACGACTCACTATAGGGAAATAATTAAACCGTAGATTAATTGCTTTCGGAG CCGCCGGGAGCTGTAG 26 TAATACGACTCACTATAGGGAGGTGGCCAAACCGTAGATTGGCCGCCCTCGGA GCCGCCGGGAGCTGTAG 27 TAATACGACTCACTATAGGGACCTCCGGAAAAAAAAAATTCCGGGCGGTCGG AGCCG 28 TAATACGACTCACTATAGGGACCTCCGGAAGCAATTCCGGGCGGTCGGAGCC G 29 TAATACGACTCACTATAGGGACCTCCGGAAGCAAACCGTCGATTCCGGGCGGT CGGAGC 30 TAATACGACTCACTATAGGGACCTCCGGAAACCGTAGATTCCGGGGGTCGGAG CCGCCGGGAGCTGTAG 31 TAATACGACTCACTATAGGGTCCGGAAACCGTCGATTCCGGGGGAGCCGCCG GGAGCTGTA 32 TAATACGACTCACTATAGGGGTGACTGCTTACCTCCGGAAACCGTAGAT 33 TAATACGACTCACTATAGGGGTGACTGCTTAGAAAACTGAACCTCCGGAAACC

GTAGAT 34 GTAAACATCAGACTTAAATAGACC 35 GGCCTGAAGGCTTTTATTTGACTG 36 TAATACGACTCACTATAGGGACTGAGGGGTCAGTGGTTCC 37 TAATACGACTCACTATAGGGACTGAGGGGTCAGTGGTTCCGGGTAGGAGCTAG CTCAAAGATTCTGCTGCAGGGATCTGCA 38 TAATACGACTCACTATAGGGAAGTCTTTGGACGTGGTTCCGGGTAGGAGCTAG CTCAAAGATTCTGCTGCAGGGATCTGCA 39 TAATACGACTCACTATAGGGCGGGGCAGCCGAGGGCCCCTGA 40 TATTGATGCTGACAGTCTGAGTCCGC 41 TAATACGACTCACTATAGGGGCGCACTGGAGCCCTGGCCAGC 42 TGACTCAGCTGACATCCAGAGAAC 43 TAATACGACTCACTATAGGGCTCGCCTCTGGGCCCGCCTTCGG 44 TGGGAGACCAGAAGAGTGCGA 45 TAATACGACTCACTATAGGGCGGGGCAGCCGAGGGCCCCTGAAGATTAGAATC GCGGCGACATGGATCGG 46 TAATACGACTCACTATAGGGCGGTTCCTAATCTGGCCCCTGAAGATTAGAATCG CGGCGACATGGATCGG 47 TAATACGACTCACTATAGGGACCTCCGGAAACCGTAGATTCCGGGCGGTCGGA GCCGCCG 48 GCGGGAGAACTACAGCTCCCGGCGGCTCCGACCGCCCGG 49 GCGGGAGAACTACAGCTAACGGCGGCGAATCAATAAAGGAATCTACGGTTTC CGGAGGT 50 GCGGGAGAACTACAGAGAAATTATTATCCGACCGCCCGGAATCTACGGTTTCC GGAGGT 51 GCGGGCTCCAGCACTCTCCCGGCGGCTCCGACCGCCCGGAATCTACGGTTTC CGGAGGT 52 GGTTGATGTTCTTATTCGCTAATTTCATGAGATGCTGTG

Chapter I: Transcript-specific Decapping of Dcp2 and

Characterization of a Dcp2-binding Element

Summary

mRNA decapping is a critical step in the control of mRNA stability and gene expression and is carried out by the Dcp2 decapping enzyme. Dcp2 is an RNA binding protein that must bind the RNA in order to recognize the cap for hydrolysis. We now demonstrate that human Dcp2 preferentially binds to a subset of mRNAs and identify sequences at the 5´ terminus of the mRNA encoding Rrp41, a core subunit component of the RNA exosome, as a specific Dcp2 substrate. A minimal element of 60 nucleotides, which is termed 2xDE, at the 5´ end of Rrp41 mRNA was identified and shown to confer more efficient decapping onto a heterologous RNA. Moreover, reduction of Dcp2 protein levels in cells resulted in a selective stabilization of the Rrp41 mRNA, confirming it as a downstream target of Dcp2 regulation. These findings demonstrate that Dcp2 can specifically bind to and regulate the stability of a subset of mRNAs. Further examination of the 2xDE element by mutagenesis assays demonstrated that the stem-loop secondary structure, but not the primary sequence, contained within the first 33 nucleotides is critical for Dcp2 binding and decapping. In addition, we showed other mRNAs containing similar secondary structures could also promote Dcp2 decapping, suggesting a general role of 5′ stem-loop in enhancing decapping activity and the utilization of this structure as a predictive tool for Dcp2 target RNAs.

Introduction

mRNA decay plays an important role in the control of gene expression and

response to regulatory events. In both yeast and mammalian cells, bulk mRNA decay

typically initiates with the removal of′ poly(A)3 tail followed by degradation of the mRNA in a 5′ to 3′ direction or a 3′ to 5′ direction (Wilusz et al, 2001b). Degradation

from 3′ end is carried out by the cytoplasmic RNA exosome, which is a multisubunit 3′ to

5′ exoribonuclease complex (Liu et al, 2006), and the resulting cap structure is

hydrolyzed by the scavenger decapping enzyme DcpS (Liu & Kiledjian, 2006). In the 5′

to 3′ decay pathway, the monomethyl guanosine (m7G) mRNA cap is cleaved first by the

Dcp2 decapping enzyme (Dunckley & Parker, 1999; Lykke-Andersen, 2002; van Dijk et al, 2002; Wang et al, 2002b) and the monophosphate RNA is degraded progressively by the 5′ to 3′ exoribonuclease Xrn1 (Decker & Parker, 1993; Hsu & Stevens, 1993).

Decapping is a highly regulated process influenced by both positive and negative regulators. In yeast, Dcp1p forms a complex with Dcp2p and is required for optimum decapping activity (She et al, 2004; Steiger et al, 2003). The Edc1p, Edc2p, and Edc3p proteins, as well as Dhh1p and Lsm1-7 protein complex, are all reported to stimulate

Dcp2p decapping (Coller & Parker, 2004). In mammals, an additional protein Edc4 (also known as Hedls and Ge-1) is shown as a positive effector of Dcp2 decapping, by either directly facilitating Dcp2 activity or promoting the association of Dcp1a with Dcp2 to possibly further enhance decapping activity in cells (Fenger-Gron et al, 2005b). In addition, AU rich elements (ARE) which confer rapid mRNA decay onto an mRNA, have also been shown to stimulate decapping in yeast (Vasudevan & Peltz, 2001) and mammals (Fenger-Gron et al, 2005b; Gao et al, 2001; Lykke-Andersen & Wagner, 2005).

In addition to these decapping activators, Dcp2 decapping can also be negatively

regulated. In yeast, both the eIF4E cap-binding protein and the poly(A) tail negatively impact decapping (Caponigro & Parker, 1995; Ramirez et al, 2002; Schwartz & Parker,

1999; Schwartz & Parker, 2000; Wilusz et al, 2001a). In mammals, eIF4E can inhibit

Dcp2 decapping in vitro (Khanna & Kiledjian, 2004) and RNAs with synthetic cap structures that bind eIF4E with higher affinity are more stable in vivo (Grudzien et al,

2006). The poly(A) tail can also negatively influence decapping and the poly(A)-binding protein (PABP) can directly inhibit decapping in vitro (Khanna & Kiledjian, 2004). The testis-specific VCX-A protein was also identified as a cap-binding protein and inhibitor of Dcp2 decapping (Jiao et al, 2006).

Dcp2 is an RNA binding protein, and can only cleave cap structure that is linked to an RNA moiety (Piccirillo et al, 2003). Uncapped RNA, but not cap analog, can inhibit Dcp2 decapping efficiently (Piccirillo et al, 2003; van Dijk et al, 2002; Wang et al,

2002b). These facts suggest that Dcp2 detects its cap substrate by RNA binding. A typical RNA binding protein usually has a basal level of nonspecific binding to all RNAs but much higher affinity for a subset of RNAs. If this is the case with Dcp2, then Dcp2 could bind RNAs differentially, and preferentially regulate a subset of mRNA. In fact, the

X29 protein, which is the nuclear decapping enzyme and a NUDIX fold protein as Dcp2, contains substrate specificity. X29 specifically binds U8 snoRNA (Ghosh et al, 2004;

Tomasevic & Peculis, 1999) and in the presence of Mg2+, cap hydrolysis is highly

specific for U8 snoRNA (Ghosh et al, 2004; Peculis et al, 2007); in contrast, in the presence of Mn2+ , all RNAs tested are decapped at high efficiency (Peculis et al, 2007).

These data suggest that decapping proteins can have preference for their RNA substrates

and raise the intriguing possibility that Dcp2 can differentially associate with and decap

specific mRNAs.

In this chapeter, we demonstrate that similar to other RNA binding proteins, Dcp2 can preferentially bind specific mRNAs and identify the 5´ termini of the mRNA encoding a core subunit of the exosome, Rrp41, as a specific Dcp2 substrate. Moreover, differential binding of Dcp2 to the Rrp41 mRNA can impact the stability of this mRNA.

Further characterization the Dcp2 binding element (2xDE) by mutational analysis revealed that the first 33 nucleotides are critical for Dcp2 decapping stimulation and this region forms a stable stem-loop structure. Mutations that disrupt the stem-loop significantly reduced decapping activity by Dcp2, which can be restored by compensatory mutations that restore the stem-loop. This suggestes the secondary structure, but not the primary sequence, is critical for Dcp2 recognition. A bioinformatic search identified a subset of mRNAs that have a comparable stem-loop structure at the 5′ end as potential Dcp2 substrates in cells. These data indicate that Dcp2 has the capability to directly and specifically regulate decapping of a subset of mRNAs in the cell.

Results

Identify mRNAs specifically bound by Dcp2

The Dcp2 human decapping enzyme is an RNA binding protein responsible for mRNA decapping. Although it has the capacity to decap any mRNA tested, we reasoned that similar to other RNA-binding proteins, it should have both a basal level of nonspecific binding to RNA as well as high affinity binding to a subset of mRNA. To begin identification of RNAs preferentially bound by Dcp2 we isolated and identified mRNAs specifically associated with Dcp2. Flag epitope-tagged human Dcp2 (Flag-Dcp2) or an epitope-tagged control RNA binding protein, the murine autosomal Deleted in

Azoospermia-like protein, mDazl (Flag-mDazl) were overexpressed in 293T cells, and

immunoprecipitated with anti-Flag antibody. Copurifying RNAs were extracted and

subjected to microarray analysis using an Affymetrix Human Genome U133A 2.0

GeneChip®. Since we were interested in RNAs specifically bound by Dcp2, RNAs that

were bound by both proteins were designated as nonspecific and not further considered.

A stringent cutoff of six fold greater binding to Dcp2 above that bound to the mDazl

control was used to designate potential mRNAs specifically bound to Dcp2.

Of the 22277 probes on the chip, 10133 RNAs with a discernable signal were

detected to be bound by either Dcp2, mDazl or both. Of those, 1461 were detected to be

preferentially bound by Dcp2 relative to mDazl, 710 were two fold, 351 were three fold,

199 were four fold, 103 were five fold and 98 were six fold or greater. Genes that were

immunopurified with Dcp2 at a level six fold or greater than that detected with mDazl are

listed in Table IV. Fourteen of the potential Dcp2 specifically bound mRNAs identified

Table IV. Dcp2 bound mRNAs

Dcp2 IP / Gene mDazl IP Descriptions SPEN 16.56 Homo sapiens spen homolog(Drosophila), transcriptional regulator EXOSC4 14.42 exosome component Rrp41 NADH dehydrogenase (ubiquinone) 1 beta subcomplex, 7 (18kD, NDUFB7 12.55 B18) unknown 12.13 gb:BF215644 NDUFS8 11.71 NADH dehydrogenase (ubiquinone) Fe-S protein 8 (23kD) unknown 11.71 gb:BE138647 NDUFS7 10.93 NADH dehydrogenase (ubiquinone) Fe-S protein 7, 20kDa PSMC3 10.93 proteasome (prosome, macropain) 26S subunit, ATPase 3 RUVBL2 10.93 Homo sapiens RuvB (E coli homolog)-like 2 NMOR2 10.56 NAD(P)H menadione oxidoreductase 2, dioxin-inducible eukaryotic translation elongation factor 1 delta (guanine nucleotide EEF1D 10.20 exchange protein) transcription elongation factor B (SIII), polypeptide 2 (18kD, TCEB2 10.20 elongin B) ADA 9.51 Homo sapiens adenosine deaminase adaptor-related protein complex 2, sigma 1 subunit, transcript AP2S1 9.51 variant AP17delta GLTSCR2 9.51 Homo sapiens glioma tumor suppressor candidate region gene 2 WDR74 9.51 WD repeat domain 74 DCXR 9.19 Homo sapiens dicarbonyl/L-xylulose reductase EDF1 9.19 endothelial differentiation-related factor 1 NARF 9.19 nuclear prelamin A recognition factor Homo sapiens structural maintenance of chromosomes SMC3 9.19 3 ,chondroitin sulfate proteoglycan 6 UQCRC1 9.19 ubiquinol-cytochrome c reductase core protein I RFC2 8.88 Human replication factor C, 40-kDa subunit (A1) WDR45 8.88 Homo sapiens WD repeat domain 45, transcript variant 1 HMOX2 8.57 heme oxygenase (decycling) 2 PSMC5 8.57 proteasome (prosome, macropain) 26S subunit, ATPase, 5 bromodomain adjacent to zinc finger domain, 1A;ATP-dependent BAZ1A 8.28 chromatin remodeling protein unknown 8.28 gb:AA669799 MRPL12 8.00 mitochondrial ribosomal protein L12 PPIB 8.00 peptidylprolyl isomerase B (cyclophilin B) RPSA 8.00 laminin receptor 1 (67kD, ribosomal protein SA) DNA segment on chromosome X (unique) 9928 expressed XAP5 8.00 sequence APRT 7.73 adenine phosphoribosyltransferase EIF3S4 7.73 eukaryotic translation initiation factor 3, subunit 4 (delta, 44kD) MAP2K2 7.73 mitogen-activated protein kinase kinase 2 NARG1 7.73 Homo sapiens NMDA receptor regulated 1 nucleoside diphosphate kinase type 6 (inhibitor of p53-induced NME6 7.73 apoptosis-alpha) Homo sapiens nucleolar protein 7, 27kDa ;retinoic acid repressible RARG-1 7.73 protein ,

serine/arginine repetitive matrix 2, RNA binding protein; AT-rich SRRM2 7.46 element binding factor unknown 7.46 gb:AW238632 FLII 7.46 Homo sapiens flightless I (Drosophila) homolog HADH2 7.46 hydroxyacyl-Coenzyme A dehydrogenase, type II JMJD6 7.46 jumonji domain containing 6, phosphatidylserine receptor MTX1 7.46 Homo sapiens metaxin 1 gb:AK000822, highly similar to Homo sapiens mRNA for PBK1 unknown 7.46 protein. CYC1 7.21 cytochrome c-1 Homo sapiens transcriptional regulator protein, SAP30 binding HCNGP 7.21 protein HSP90B1 7.21 heat shock protein 90kDa beta (Grp94), member 1 MRPS11 7.21 mitochondrial ribosomal protein S11 POLR2L 7.21 RNA polymerase II (DNA directed) polypeptide L (7.6kD) RPS5 7.21 ribosomal protein S5 DGCR6 7.21 Homo sapiens DiGeorge syndrome critical region gene 6 ETFB 7.21 Homo sapiens electron-transfer-flavoprotein, beta polypeptide IDH3G 7.21 isocitrate dehydrogenase 3 (NAD+) gamma MRPL23 7.21 mitochondrial ribosomal protein L23 PITRM1 7.21 Homo sapiens pitrilysin metalloproteinase 1 Homo sapiens translocated promoter region (to activated MET TPR 7.21 oncogene) UBXD1 7.21 Homo sapiens UBX domain containing 1 CHMP2A 6.96 Homo sapiens chromatin modifying protein 2A, transcript variant 1 Homo sapiens deformed epidermal autoregulatory factor DEAF1 6.96 1(Drosophila) DLEC1 6.96 deleted in lung and esophageal cancer 1 GATA zinc finger domain containing 1,ocular development GATAD1 6.96 associated PDXK 6.96 pyridoxal (pyridoxine, vitamin B6) kinase PPP1R7 6.96 protein phosphatase 1, regulatory subunit 7 TST 6.96 thiosulfate sulfurtransferase (rhodanese) Homo sapiens N-acetyltransferase, homolog of S. cerevisiae ARD1 6.73 ARD1 BID 6.73 Homo sapiens BH3 interacting domain death agonist CHAF1A 6.73 chromatin assembly factor 1, subunit A (p150) GCP60 6.73 golgi resident protein GCP60 LSM3 6.73 Homo sapiens LSM3 homolog, U6 small nuclear RNA associated MRPS12 6.73 mitochondrial ribosomal protein S12 human mRNA turnover 4 homolog (S. cerevisiae), 60S acidic MRTO4 6.73 ribosomal protein PO MYG1 6.73 Homo sapiens MYG1 protein NOP56 6.73 nucleolar protein (KKED repeat) NT5DC2 6.73 5'-nucleotidase domain containing 2 PP3111 6.73 human dihydrouridine synthase 1-like (S. cerevisiae) Human D9 splice variant A ,stimulated by retinoic acid 13 homolog STRA13 6.73 (mouse) TRAPPC2L 6.73 trafficking protein particle complex 2-like TRIB3 6.73 Human tribbles homolog 3 (Drosophila),neuronal cell death

inducible putative kinase TTC3 6.73 tetratricopeptide repeat domain 3 unknown 6.73 gb:AW237172 unknown 6.73 gb:AI588986 ZNF593 6.73 Homo sapiens zinc finger protein 593 LOC728233 6.50 phosphatidylinositol 4-kinase, catalytic, alpha pseudogene PHGDH 6.50 phosphoglycerate dehydrogenase unknown 6.50 gb:BF001806,antigen identified by monoclonal antibody Ki-67 APEH 6.28 N-acylaminoacyl-peptide hydrolase chromosome 11 open reading frame 48,hypothetical protein C11orf48 6.28 LOC79081 CCDC44 6.28 coiled-coil domain containing 44 CUEDC2 6.28 CUE domain containing 2 Homo sapiens terminal deoxynucleotidyltransferase interacting DNTTIP2 6.28 protein 2 FAHD2A 6.28 fumarylacetoacetate hydrolase domain containing 2A NDUFA13 6.28 NADH dehydrogenase (ubiquinone) 1 alpha subcomplex, 13 SAS 6.28 N-acetylneuraminic acid phosphate synthase; sialic acid synthase DGS-I 6.06 DiGeorge syndrome critical region gene 14 ENO1 6.06 Human enolase 1 PRPF40A 6.06 PRP40 pre-mRNA processing factor 40 homolog A (S. cerevisiae) Homo sapiens SCO cytochrome oxidase deficient homolog 2 SCO2 6.06 (yeast) UFC1 6.06 ubiquitin-fold modifier conjugating enzyme 1

by the microarray were further tested to confirm their binding by RT-PCR.The Stx7

mRNA which was specifically bound by mDazl was used as a negative control.

Approximately 65% of the candidate mRNAs (nine out of 14) were confirmed to specifically associate with Dcp2 relative to mDazl by their selective coimmunopurification with Flag-Dcp2 (Figure 1A). Surprisingly, the top candidate from the microarray data with a 16.6 fold greater association to Dcp2, the Spen mRNA, was not reproducibly detected to specifically bind Dcp2. However, the second highest candidate in the microarray analysis, the Rrp41 mRNA, was reproducibly bound by Dcp2 in the RT-PCR. In addition to Rrp41, mRNAs encoding Ndufb7, Ndufs8, Psmc3, Ruvb2,

Gltscr, Edf1, Hmox2 and Ppib were selectively immunopurified by Flag-Dcp2 expressed in 293T cells while the Tceb2, Ada, Ap2s1 and Narf mRNAs were not reproducibly coimmunnopurified (Figure 1A). As expected, the Stx7 mRNA was only detected in the mDazl-bound fraction (Figure 1A). These data show that despite the ubiquitous requirement of Dcp2 for mRNA decapping, Dcp2 can specifically associate with a set of mRNAs and may exhibit selective regulation of mRNA decay.

The above analysis was carried out with mRNA coimmunopurified with Flag-

Dcp2 expressed in cells and therefore, the isolated mRNA could either be a consequence

of direct Dcp2 binding to the mRNA or indirect association through a protein-protein

interaction network. To test whether the binding to Dcp2 was direct or indirect, bacterial

expressed GST-Dcp2 was used to determine which mRNA could copurify with Dcp2 in

the absence of additional proteins. Three of the Flag-Dcp2 copurifying RNAs were randomly chosen and tested for their ability to be bound by recombinant Dcp2 protein: a

subunit of the exonuclease exosome complex, Rrp41 mRNA (Liu et al, 2006); the endo-

Figure 1. Identification of Rrp 41 mRNA as a substrate directly bound by Dcp2.

(A) Identification of mRNAs specifically associated with Dcp2. 293T cell mRNAs bound by Flag-Dcp2 (lane 3) or Flag-mDazl (lane 2) were isolated, reverse transcribed with random primer, and amplified by gene-specific primers for the indicated mRNAs.

An aliquot of RT-PCR products using 293T total RNA to designate the size of the correct band are shown in lane 1, and RT-PCR products from a mock IP are indicated in lane 4 as negative control. RT-PCR results that were reproducibly detected to be preferentially bound by Flag-Dcp2 in three independent sets are indicated as “+”. Those that were not reproducible in all three experiments are indicated as “-”. The Stx7 negative control mRNA was associated with mDazl.

(B) Rrp41 mRNA is bound by Dcp2 directly. GST pull-down assays were carried out with 4µg of the indicated fusion protein and 32P-labeled Rrp41 or Stx7 RNA. RNA bound to GST-Dcp2 or GST-mDazl were isolated and resolved on a denaturing PAGE. Five percent of each RNA used was included in the input lanes.

thelial differentiation-related factor 1 (Edf1) mRNA that functions as a transcriptional

coactivator in endothelial cell differentiation and the mRNA encodes a subunit of the

mitochondria NADH:ubiquinone oxidoreductase (complex I), Ndufb7. mRNA encoding

the syntaxin7 (Stx7) protein which is involved in post-Golgi vesicle-mediated transport

was included as a negative control mRNA. Stx7 was selectively bound by mDazl relative

to Dcp2 in the microarray screen. RNAs corresponding to the full length Rrp41, Edf1,

Ndufb7 and Stx7 mRNAs were transcribed in vitro in presence of 32P-UTP. The labeled

RNAs were incubated with GST-Dcp2 or a control GST-mDazl protein. After extensive washes, copurified RNAs were isolated and resolved on a polyacrylamide gel. We were unable to detect consistent binding of Edf1 and Ndufb7 to GST-Dcp2 indicating that the above detected binding is most likely due to indirect association with Dcp2 (data not shown). However, the Rrp41 RNA was reproducibly and specifically bound to GST-

Dcp2, while the Stx7 RNA only bound to GST-mDazl (Figure 1B). These data demonstrate that the binding of Dcp2 to the Rrp41 RNA is direct and does not require additional proteins.

Collectively, the above data demonstrate that Dcp2 has the capacity to specifically bind a subset of mRNA either directly or indirectly. Furthermore, the specificity of Dcp2 binding to the Rrp41 mRNA was intriguing considering that Rrp41 is an essential component of the exosome complex. As Dcp2 is the critical component of the 5´ to 3´ mRNA decay pathway and the exosome is the critical 3´ to 5´ decay complex, the binding of Dcp2 to Rrp41 mRNA was of particular interest and the binding of Dcp2 to this substrate RNA was further characterized.

Rrp41 mRNA is preferentially decapped by Dcp2 in vitro

We next addressed whether the preferential binding of Dcp2 to the Rrp41 mRNA

correlated with increased decapping. The decapping efficiency of the Dcp2-bound Rrp41

RNA substrate was compared to that of the Stx7 RNA which appears to be a poor Dcp2

binding substrate. The RNAs were in vitro transcribed and cap labeled with [α32P]GTP

and used in decapping assays with bacterial expressed His-Dcp2. The Rrp41 RNA consisted of the 900nt complete Rrp41 mRNA containing a 110 nucleotide 5´ UTR, 740 nucleotide coding region and 50 nucleotide 3ÚTR. To maintain consistency of the RNA substrate size, the 5′ 900 nucleotides of the Stx7 RNA was used that included 60 nucleotide 5ÚTR and 780 nucleotide coding region and 60 nucleotide 3ÚTR. The cap-

labeled RNAs were incubated with a titration of His-Dcp2 and the decapping products

were resolved by polyethyleneimine-cellulose (PEI) thin layer chromatography (TLC) developed in 0.45 M (NH4)2SO4. As expected, the decapping efficiency for both RNAs increased with increasing His-Dcp2 protein used (Figure 2A). The increase is also an indication that the reactions were carried out within a linear range for His-Dcp2 protein.

Interestingly, decapping of the Rrp41 RNA was considerably more efficient than the

control Stx7 RNA, with an average value of 6.5 fold greater decapping compared to the

decapping observed with Stx7 RNA. This suggests that specific binding of Dcp2 to an

RNA leads to enhanced decapping of the RNA.

The preferential decapping of a Dcp2-bound RNA was not restricted to bacterial

expressed Dcp2 and was also observed using both Flag-Dcp2 expressed in cells as well as

endogenous Dcp2. Flag-Dcp2 expressed in 293T cells was immunoprecipitated and the

bead associated protein was used in a decapping assay with Rrp41 or Sxt7 RNAs. With

Figure 2. Rrp41 mRNA is preferentially decapped by Dcp2.

(A) Rrp41 mRNA is preferentially decapped by Dcp2. Decapping assays were carried out using the indicated amount of bacterial expressed His-Dcp2 with cap labeled Rrp41 or

Stx7 RNA, and the reaction products were resolved by PEI TLC. Position of the capped

RNA substrate and m7GDP decapping product are indicated on the left. Decapping efficiency of Stx7 RNA was arbitrarily set to 1. The fold increase in decapping observed of the Rrp41 RNA relative to the Stx7 RNA obtained from three independent experiments are presented in the graph to the right with Standard Deviation (SD) denoted by the error bars. Quantitations obtained with the decapping observed with 80ng of His-

Dcp2 are shown, but are similar to that observed with 40ng protein.

(B) Rrp41 mRNA is preferentially decapped by immunoprecipitated Flag-Dcp2.

Decapping assays were carried out as described above except the decapping reactions

were carried out with the indicated volume of Flag-bead slurry containing

immunopurified Flag-Dcp2. Labeling and quantitations are as described in A above.

were carried out with the indicated concentration of K562 cell cytoplasmic P50 extract.

His-Dcp2 was used in lane 1 as a positive control. Labeling and quantitations are as

described in A above.

this assay system, a consistent average increase of two fold in the efficiency of decapping

Rrp41 RNA was observed (Figure 2B). Preferential decapping of the Rrp41 RNA was

also observed with endogenous Dcp2. Although Dcp2 activity is not readily detected in

cell extract due to decapping inhibitors, low levels of decapping can be detected in the

P50 cytoplasmic fraction (Jiao et al, 2006). With the level of K562 cell cytoplasmic P50

fraction used, Dcp2-mediated decapping that generates m7GDP can be detected with the

Rrp41 RNA and significantly lower levels with Stx7 RNA (Figure 2C). Decapping of the

Rrp41 RNA was approximately 3.5 fold greater than that observed with the Stx7 RNA.

Although the fold increase of decapping observed with the Dcp2 expressed in eukaryotic

cells (Figure 2B and 2C) is less than that observed with the bacterial expressed Dcp2

(Figure 2A), this is consistent with the presence of Dcp2 decapping inhibitors in cells

(Jiao et al, 2006). Taken together, these data reveal a novel role of Dcp2. Dcp2 itself

contains an intrinsic property to distinguish different RNA substrates and can

preferentially associate with and decap specific mRNAs and does not indiscriminately

hydrolyze all mRNAs equally.

Identification of the Dcp2 target sequence in the Rrp41 mRNA

In order to identify the sequence element within the Rrp41 mRNA that was

responsible for the recruitment of Dcp2, different truncations spanning the Rrp41 mRNA

were generated (Figure 3A) and tested for their ability to be decapped by Dcp2 in an in vitro decapping assay. Three fragments corresponding to the first half, second half and middle third of the Rrp41 mRNA spanning the junction of the two halves were tested. As

Figure 3. Dcp2 target sequence on Rrp41 mRNA is located in the 5′UTR.

(A) A schematic representation of the various Rrp41 mRNA truncations is shown with

the 5′UTR, coding region and 3′UTR indicated.

(B) 5′UTR of Rrp41 mRNA is the stimulatory region for Dcp2 decapping. RNAs shown

in panel A were used as substrates in decapping assays with the indicated amount of His-

Dcp2 protein. Stx7 RNA substrate was used as a negative control to designate a basal level of Dcp2 decapping.

(C) Quantitation of the percent decapping observed in panel B is presented. Fold increase in decapping observed of the Rrp41 RNA fragments relative to the Stx7 RNA is shown.

Quantitations from three independent experiments are presented in the graph with

Standard Deviation (SD) denoted by the error bars.

shown in Figure 3B, the 5´ half of the RNA was decapped three fold more efficiently

than the other two RNAs that were comparable to the level detected with the Stx7

negative control. Therefore the decapping stimulatory region, is contained within the 5´

half of the Rrp41 mRNA.

The Dcp2 recruitment region within the 5´ half of Rrp41 mRNA was further narrowed by separately testing decapping of the 110 nucleotide 5´ UTR and the remaining 400 nucleotide coding region. As shown in Figure 3B, decapping activity comparable to the full length 5´ half of the Rrp41 RNA is exclusively associated with the

5´UTR (lanes 13-15 and 16-18), but not the 5´portion of the coding region (lanes 19-21).

From these data (summarized in Figure 3C), we conclude that the 5´UTR of Rrp41

mRNA is essential for stimulating Dcp2 decapping and most likely contains the Dcp2

binding region. This suggests a model in which Dcp2 directly binds to the 5´end of a

RNA and preferentially decaps it.

The Rrp41 5´UTR can function on a heterologous RNA to promote decapping

The above data indicate that the Rrp41 5′UTR is involved in recruitment of Dcp2.

We next asked whether it can also function when placed onto a heterologous RNA. To

test this, the 110 nucleotide Rrp41 5´UTR was used to replace the 5′UTR of the Stx7

RNA since this RNA is normally inefficiently decapped (Figures 2A). The Rrp41-Stx7

chimeric RNA was used as substrate for in vitro decapping assays with bacterial

expressed recombinant His-Dcp2. Figure 4 shows that while the Stx7 RNA with its own

5´UTR was poorly decapped by Dcp2 (lanes 7-9), decapping efficiency of the chimeric

Figure 4. The Rrp41 5´UTR is an autonomous element conferring higher decapping

activity. Chimeric RNA that harbors the Rrp41 5´UTR at the 5´ end followed by Stx7 sequences was used in decapping assays with bacterial expressed His-Dcp2 (lane 4-6).

Stx7 RNA with its own 5´UTR was used as negative control (lane 7-9) and the Rrp41

RNA was used as a positive control (lane 1-3). Quantitations of the fold increase in percent decapping relative the Stx7 RNA are shown in the graph to the right. Decapping efficiency of the chimeric RNA is comparable to that of Rrp41 RNA. Labeling and quantitation of three independent experiments is as described in the legend to Figure 1.

RNA that bears the Rrp41 5´UTR was enhanced by approximately 10 fold (lanes 4-6).

Therefore, the Rrp41 5´UTR can recruit Dcp2 onto a chimeric RNA and appears to be an autonomous element that can confer higher decapping activity onto a heterologous RNA.

Identification of a minimal Dcp2 binding element

The fact that the Rrp41-Stx7 chimeric RNA was efficiently decapped enabled us to use this RNA to further narrow down the minimal sequence capable of stimulating decapping. Initially two chimeric RNAs were generated containing the first or second half of the Rrp41 5′UTR fused to the Stx7 RNA (Figure 5A). The capacity of Dcp2 to decap each RNA was tested and compared to the chimeric RNA containing the full length

Rrp41 5′UTR. As seen in Figure 5B, the RNA bearing nucleotides 1-60 of Rrp41 was decapped as efficiently as the RNA bearing the full length Rrp41 5´UTR (compare lanes

8-9 to 2-3). In contrast, the decapping efficiency of the RNA bearing nucleotides 50-110 was significantly reduced relative to the Rrp41 5′UTR-containing RNA.

Examination of the 60 nucleotide sequence revealed it to be rich in guanosine and cytosines with an interesting duplication of a 25 nucleotide motif (Figure 5C) and we term the 60 nucleotide element 2xDE (Dcp2 decapping element). The above analysis demonstrates that 2xDE at the 5´ terminal sequence of the Rrp41 mRNA can be specifically and efficiently decapped by Dcp2. However, whether this was due to the structure of the RNA being more accessible for decapping or an active selective binding process remained to be determined. To address this question, we tested whether Dcp2 can specifically bind the RNA by electrophoretic mobility shift assays. Uniformly 32P-

Figure 5. Identification of Dcp2 binding element (2xDE).

(A) A schematic representation of the various chimeric RNAs that contain different fragments of Rrp41 5´UTR upstream of the Stx7 coding region are shown. Numbers represent the corresponding nucleotides of Rrp41 5´UTR included in the RNA.

(B) The first 60 nucleotides of the Rrp41 5´UTR are responsible for the stimulation of

decapping. Decapping assays were carried out with the indicated chimeric RNAs and

Stx7 as a negative control. Quantification of decapping efficiency of each RNA relative to Stx7 is shown.

(C) An alignment of the two halves of first 60 nucleotides of Rrp41 5´UTR is shown with the corresponding nucleotide numbers on the left.

(D) Specific binding of Dcp2 to the Rrp41 5´UTR. An electrophoretic mobility shift

assay was carried out with 1µg of His-Dcp2 and uniformly 32P-labeled uncapped RNA

corresponding to the first 60 nucleotides of the Rrp41 5´UTR, and competed with an increasing amount of the indicated unlabeled RNAs. The control RNA was the 110 nucleotide α-globin mRNA 3´UTR.

labeled uncapped RNA consisting of the cap proximal 60 nucleotides of the Rrp41 mRNA was incubated with His-Dcp2 and the bound complex resolved by native gel electrophoresis. As shown in Figure 5D, a Dcp2-RNA complex can be detected upon addition of Dcp2 (lane 2). The binding was specific as it was competed by unlabeled self competitor but not an unrelated control RNA (compare lanes 3-6 to 7-10). Therefore the

2xDE appears to facilitate Dcp2 decapping by an active selective binding process.

Dcp2 can specifically regulate Rrp41 mRNA stability in cells

To address the functional significance of Dcp2 binding to the Rrp41 mRNA, we determined whether the stability of the Rrp41 mRNA is specifically influenced by Dcp2.

Our rational was that since Dcp2 is a component of the mRNA decay machinery, the

Rrp41 mRNA should be selectively stabilized in cells containing a reduction of Dcp2 protein relative to the Stx7 mRNA. siRNA directed against the Dcp2 mRNA was used to reduce Dcp2 expression in 293T cells. Dcp2 protein levels were efficiently reduced by

90% following two successive transfections of siRNA at 72 hours post-transfection

(Figure 6A). Therefore, this time interval was used in the subsequent experiments to assess the influence of Dcp2 protein. 293T cells containing a reduction in Dcp2 levels were subjected to actinomycin D to block transcription at the 72 hour siRNA post- transfection time point. RNA was isolated from cells transfected with either Dcp2- specific siRNA or control siRNA at increasing time intervals following actinomycin D up to eight hours. Rrp41 mRNA levels were determined by quantitative real time RT-PCR and the values normalized to the U6 snRNA. The U6 snRNA is uncapped and should not

Figure 6. Dcp2 regulates Rrp41 mRNA stability in cells.

(A) siRNA-directed knockdown of Dcp2 expression in 293T cells. 293T cells were transfected twice during a period of 72h with a control siRNA or siRNA specific for

Dcp2. Endogenous Dcp2 was detected by western blot analysis using affinity purified

Dcp2 antibody. The level of an unrelated decapping enzyme, DcpS, was monitored as an internal control. Bacterial expressed His-Dcp2 was used in the last lane as a positive control.

(B) Stabilization of the Rrp41 mRNA upon Dcp2 knockdown. 293T cells were treated with actinomycin D to block transcription 72 hours following the second of two series of siRNA transfections. The levels of Rrp41 or Stx7 mRNA remaining following 0, 2, 4, 8

hour actinomycin D treatment were determined by quantitative real-time RT-PCR

(qPCR). Values of quadruplicate qPCRs from two sets of independent RNA preparation normalized to the U6 snRNA are shown with +/- Standard Deviation represented by the error bars.

(C) The Rrp41 5´UTR 60nt element (2xDE) can enhance decapping in cell. Cap labeled and G-tract tailed Stx7 RNAs (510nt) contianing either its own 5’UTR (60 nt) or the

Rrp411-60 element, were electroporated into 293T cells. Cells were harvested at the

indicated time points and RNAs were isolated and resolved on a 5% polyacrylmide gel.

Quantitative of three independent experiments normalized relative to the internal control,

which was included in the harvest buffer, are presented in the graph with Standard

Deviation represented by the error bars.

be affected by Dcp2. As expected, the half-life of the Rrp41 mRNA was increased two fold from 3.5 hours in the control siRNA knockdown cells to 7 hours in Dcp2 knock down cells (Figure 6B). In contrast, the decay rate for the Stx7 mRNA, which is a poor substrate for Dcp2 in vitro, did not change when Dcp2 was knocked down and was similar to that observed in the control knockdown cells. These data indicate that the

Rrp41 mRNA is a true Dcp2 target in cells and validates the in vitro analysis above demonstrating selective binding and decapping of mRNA by Dcp2.

To further clarify the role of the Dcp2 binding-decapping element in the regulation of Rrp41 mRNA stability in cells, we tested whether this element could enhance decapping in cells. Cap-labeled Stx7 RNA (510nt) were made with either its own 60 nt 5′UTR or the 1-60 nt element of Rrp41 5′UTR, and tailed with poly(G) tract consisting of 16 guanosines to protect the 3′ termini from 3´ to 5´ exonucleolytic degradation (Wang & Kiledjian, 2001). The two cap-labeled RNAs were electroporated into 293T cells and RNAs remaining at distinct time intervals up to 4 hours were resolved by denaturing polyacrylamide gel. Since the RNAs are labeled exclusively at the cap and is protected from 3´ exonucleolytic decay by the G-tract, loss of detectable RNA directly correlates with decapping (Wang & Kiledjian, 2001). As shown in Figure 6C, the Stx7

RNA containing the Rrp41 60nt 2xDE at its 5´ termini degrades significantly faster, with a half-life of 1 hour compared to approximately 2.2 hours for the Stx7 RNA lacking the

Rrp41 sequence. These data demonstrate that the 60nt element can enhance mRNA decay by promoting mRNA decapping and it can function on a heterologous RNA in cells.

The first 33nt of 2xDE is critical for Dcp2 binding and decapping

To further uncover the mechanism of how the 2xDE element facilitates Dcp2

decapping, we made a series of mutations within this element. Every ten or eleven

successive nucleotides were mutated within the 2xDE element as indicated in Figure 7.

Chimeric RNAs were in vitro transcribed harboring the various mutated 2xDE sequences

placed upstream of the Stx7 mRNA sequence, which is normally a poor substrate for

Dcp2 decapping. These RNAs were cap-labeled at the first phosphate following the

methylguanosin and tested in an in vitro decapping assay with recombinant histidine-

tagged Dcp2 (His-Dcp2) protein. Consistent with the data above, the chimeric Stx7 RNA

containing the wild type (WT) 2xDE sequence at the 5´ end (lane 1-3) displayed an

enhancement of decapping of approximately 4 fold relative to the level of decapping

detected with the Stx7 RNA (Figure 7A, compare lanes 1-3 to 19-21). RNAs containing

mutations within the first 33 nucleotides of the 2x-DE (nucleotides 1-10, 11-22, and 23-

33) were less efficiently decapped by Dcp2 to a level comparable to that observed with

the Stx7 RNA (lanes 4-12); while mutations of nucleotides 34-44 or 45-55 were

decapped at approximately wild type 2xDE decapping levels. These results suggest that

the first 33 nucleotides of the 2xDE are critical for enhancement of Dcp2-mediated decapping.

To address whether the reduced decapping of the mutated RNAs correlated with reduced Dcp2 binding, we tested the binding of Dcp2 to each 2xDE mutant by electrophoretic mobility shift assays. The same series of mutated 2xDE elements were in

vitro transcribed and uniformly 32P labeled, and incubated with His-Dcp2. The bound complex was resolved by native gel electrophoresis and as shown in Figure 7B, a specific

Figure 7. The first 33 nucleotides of the 2xDE element are critical for enhanced

Dcp2 decapping and binding.

(A) The first 33 nucleotides of the 2xDE are important for Dcp2 decapping. Decapping assays were carried out using increasing concentrations of bacterial expressed His-Dcp2 with cap labeled 2xDE wild type or mutant RNAs as indicated or the Stx7 RNA negative control. Transversion mutations were generated where adenines were substituted by cytosines, thymines by guanosines, cytosines by adenines, and guanosines by thymines.

All RNAs were transcribed as chimeric RNAs with the coding region of Stx7 RNA linked to the 3′ end to maintain the same size as the negative control Stx7 RNA. The

reaction products were resolved by PEI-TLC and the position of the capped RNA substrate and m7GDP decapping products indicated on the left. Decapping efficiency of wild type 2xDE RNA with 60ng of His-Dcp2 was designated as 100 and the corresponding decapping efficiencies of the 2xDE mutant RNAs obtained from three independent experiments are graphically presented on the right with Standard Deviation

(SD) denoted by the error bars. Similar results were observed with the 30ng protein assays.

(B) The first 33 nucleotides of the 2xDE are important for binding of Dcp2. An electrophoretic mobility shift assay was carried out with increasing concentrations of His-

Dcp2 and uniformly 32P-labeled uncapped RNA described in A. The slower migrating

complex detected with the wild type RNA was designated as the Dcp2-complex while the

faster migrating complex detected with the Stx7 negative control was denoted as an

alternative complex. Binding efficiency of wild type 2xDE RNA with 2µg of His-Dcp2

was designated as 100 and the corresponding binding efficiencies of the 2xDE mutant

RNAs from three experiments are presented relative to the wild type value on the right with Standard Deviation (SD) denoted by the error bars. A direct correlation between formation of the slower migrating complex and efficiency of decapping was observed.

Dcp2-RNA complex can be detected on the wild type 2xDE RNA (lane 2-3). A similarly

migrating complex was inefficiently formed on the Stx7 control RNA (lanes 19 - 21).

However, an alternate faster migrating band was observed (Figure 7B, lanes 19 - 21), which is likely to be non-specific and is primarily detected in the absence of a high affinity binding site. Furthermore, formation of the slower migrating Dcp2 complex observed on the wild type 2xDE RNA was also more prominent on mutants that retained the ability to stimulate decapping (Figure 7B, lanes 13 - 18). Interestingly, this complex was considerably diminished with the 2xDE RNAs containing substitutions within the

first 33 nucleotides that did not support stimulated decapping (lanes 4 - 12). These mutants were more likely to form the faster migrating alternate complex analogous to the one formed on the Stx7 RNA. Collectively, these data demonstrate that the first 33nt of the 2x-DE are important for specific binding of Dcp2 that corresponded to enhanced decapping and appear to constitute the functional unit of the 2xDE for Dcp2 activity.

The presence of a 5′ end stem-loop structure within the 2xDE

The function of an RNA element can be attributed to either its primary nucleotide

sequence or its higher order structure. Examination of the 2xDE element by MFold prediction revealed a stable stem-loop structure within the critical first 33nt region. To

determine whether this region can form a stable stem-loop structure, we carried out an

RNase-mapping assay. The 2xDE RNA was transcribed in vitro and 32P-labed at the 5′

terminus. The full length RNA was gel purified and incubated with RNase A, RNase T1 or RNase V1. RNase A, which cleaves single strand pyrimidines, showed strong cleavage

Figure 8. The first 33 nt of the 2xDE forms a stem-loop secondary structure.

(A) RNase mapping of the 2xDE element. In vitro transcribed and 5′ end 32P-labeled

2xDE RNA was incubated with the indicated RNases for 15 minutes and resolved by a

15% polyacrylamide gel. AH designates the alkaline hydrolysis lane to generate a single nucleotide ladder. A RNA size marker is shown on the rightmost lane.

(B) Deduced secondary structure of the 2xDE RNA from A. Secondary structure of the

2xDE deduced by MFold program is shown along with the indicated RNase cleavage sites denoted by arrows. The predicted secondary structure is consistent with the RNase mapping results.

of the 2xDE RNA at U18 and additional cleavages at C15 and C16 (Figure 8A, lane 3

and 4). RNase T1 cleaves at single stranded guanosine residues and efficiently cleaved

G17 and G20. RNase V1 which cleaves double stranded regions of RNA showed

predominant cleavage from nucleotide 5 to nucleotide 12. The combined cleavage pattern

of the three RNases is shown in the right panel, and is consistent with that predicted by

the Mfold program. This confirms that the 5′ end region of the 2xDE can form a stable

stem-loop structure, with the first strand of the stem containing nucleotide 3-13, second

strand containing nucleotide 22-33 and an intervening loop of eight bases in between.

The 5′ stem-loop structure facilitates Dcp2 decapping

To test whether the 5′ stem-loop structure within the 2xDE contributes to the enhanced decapping activity or whether the enhanced decapping can be attributed to the primary sequence, we generated a series of mutations that disrupted or regenerated the stem-loop structure as indicated in Figure 9A and tested the in vitro decapping activity of

each mutant. As shown in Figure 9B, disruption of the stem-loop structure by mutating

either the first strand (mutant A) or the second strand (mutant B) of the stem,

significantly decreased decapping of the RNA (lane 4-9) to approximately 20-40 percent

of the activity of that detected with the wild type 2xDE. To distinguish whether the

decrease was due to changes in the primary or secondary sequence, a compensatory set of

mutants were generated such that the primary sequence was changed but the stem-loop

structure was restored (mutant C). Reconstituting the stem-loop recovered decapping to

80 percent of the wild type level (lane 10-12) indicating that the stem-loop structure and

Figure 9. The intact stem-loop structure is critical for promoting Dcp2 decapping.

(A) A schematic representation of the structures of the wild type and various mutant

2xDE RNAs are shown. WT: wild type; A: mutations in the first strand of the stem; B: mutations in the second strand of the stem; C: complementary mutations in both strands of the stem; D: the two strands of the stem are flipped.

(B) Decapping assays were carried out with the indicated mutant 2xDE RNAs and the

Stx7 negative control. Quantification of decapping efficiencies of each RNA obtained

from three independent experiments are shown on the right with the decapping efficiency

of wild type 2xDE RNA arbitrarily set to 100 and the Standard Deviation (SD) denoted

by the error bars.

not the primary sequence within the 5′ end of 2xDE is important for Dcp2 recognition.

An additional confirmation of this conclusion was provided with a mutation that flipped

the two strands of the stem (mutant D), such that the mutant has the same stem-loop

structure as the wildtype 2xDE but with a different primary sequence. Consistently, this mutant maintained the same level of decapping activity (lane 13-15) as the wild type element. Collectively, these data support the conclusion that the stem-loop structure at the

5′ end of 2xDE element enhances Dcp2 decapping and constitutes a Dcp2 binding and preferential decapping element we will refer to as the Dcp2 Binding and Decapping

Element (DBDE).

Having demonstrated that the stem-loop structure was critical for Dcp2 recognition and decapping, we next set out to further determine parameters of the loop region and the length of the stem that can be tolerated as a substrate for Dcp2. An additional set of mutations were generated that changed the sequence within the loop region of the DBDE (Figure 10A, mutant E); shortened the loop to 4nt (mutant F) or increased it to 12nt (mutant G); removed the single nucleotide bulge in the stem (mutant

H); or shortened the stem to 7 base pairs (mutant I) by deleting the lower 4 base pairs of

the stem. The decapping activity of each mutant was tested in vitro and compared to the

wild type 2xDE. As shown in Figure 10B, the mutations in the loop region (E,F,G) had a

moderate effect (20-30 percent) on the decapping activity of Dcp2, which suggest

nucleotides in the loop are not critical for Dcp2 recognition. The bulge in the stem

appears dispensable for decapping since its removal (H) did not significantly compromise

Dcp2 decapping of the RNA. Importantly, when the stem was shortened to 7 base pairs

(I), the decapping activity significantly decreased to approximately 30 percent of the wild

Figure 10. Mutational analysis of the stem-loop structure.

(A) A schematic representation of the predicted structures for the various mutant 2xDE

RNAs are shown. WT: wild type; E: substitution of the loop region; F: deletion of the

loop to 4 nucleotides; G: increase the loop to 12 nucleotides; H: deletion of the bulge at nucleotide 29; I: deletion of the lower 4 base pairs of the stem.

(B) Decapping assays were carried out with the indicated mutant 2xDE RNAs and the

Stx7 negative control. Decapping efficiencies from three independent experiments were quantitated for each RNA and graphically presented. The decapping efficiency of the wild type 2xDE RNA was arbitrarily set to 100.

type level, suggesting a stem longer than 7 base pairs was required for Dcp2 recognition.

Requirement for cap proximity of the DBDE

Since the DBDE exerts a function on the 5´ cap, we reasoned proximity to the cap

might be an important parameter. The stem region of the Rrp41 DBDE begins three

nucleotides downstream of the cap. Decapping was significantly compromised when the

stem-loop was positioned either 10 or 20 nucleotides from the 5´ terminus of the RNA by

insertion of Stx7 sequences at the 5´ end (Figure 11A, compare lanes 1-3 to lanes 4-6 and

7-9). To determine whether positioning of the DBDE relative to the 5´ cap influenced

Dcp2 binding to the RNA, binding of Dcp2 to the variant RNAs was tested by an

electrophoretic mobility shift assay. Consistent with the decapping data, movement of

the DBDE distal to the cap an addition seven nucleotides resulted in reduced formation of

the slower migrating Dcp2-RNA complex as determined by the gel shift assay (Figure

11B, compare lanes 1-3 to lanes 4-6 and 7-9). These data indicate that the position of the

stem-loop structure is important for facilitating enhanced Dcp2-mediated decapping and

should be less than 10 nucleotides from the 5′ end of the RNA.

Generality of decapping enhancement by a stem-loop structure at the 5′ end of an

mRNA

The dependence on structure rather than sequence of the DBDE indicates that the presence of this element at the 5´ end of other mRNAs could also promote decapping by

Dcp2. As an initial approach we first examined mRNAs we demonstrated above that are

Figure 11. Positioning of the DBDE stem-loop structure less than 10 nucleotides from the 5´ cap is important for Dcp2-mediated enhanced decapping.

(A) Decapping assays were carried out with wild type 2xDE RNA and 2xDE RNA containing 10 or 20 nucleotides of the Stx7 5′ end sequences. All RNAs were transcribed as chimeric RNAs with the coding region of Stx7 RNA attached to the 3′ end to maintain the same size as the negative control Stx7 RNA.

(B) Electrophoretic mobility shift assays were carried out with 32P-uniformly labeled

uncapped RNAs described in A above. Binding efficiencies relative to the wild type

RNA from two experiments are presented on the right with Standard Deviation (SD) denoted by the error bars.

specifically copurified with Dcp2, including the mRNA encoding Rrp41 and the Ndufb7

mRNA (Figure 1A). Similar to the predicted structure of the Rrp41 mRNA, MFold

predictions of the Ndufb7 mRNA also revealed a stem-loop structure at the 5′ terminus

(Figure 12A). We tested the decapping activity of Dcp2 on this mRNA by an in vitro decapping assay. As shown in Figure 12B, compared to the Stx7 mRNA negative control

(lane 10-12), the Nudfb7 mRNA demonstrated a 2.5 fold enhancement of decapping activity (lane 1-3). To test whether the predicted stem-loop structure influenced the decapping, nucleotide 35-44 in the second strand of the predicted double stranded stem region were mutated. The RNA containing this mutation was decapped with an approximate 40 percent reduced efficiency (lane 4-6). However, a compensatory mutation of nucleotide 2-11 in the first strand fully restored decapping activity (lane 7-9).

This result suggests the predicted stem-loop structure contributes to decapping of the

Ndufb7 mRNA and could be a general modulator of mRNA decapping. To further test the predictive potential of the DBDE, the human 5´UTR database was searched for mRNAs that could potentially form a 5´ stem-loop structure analogous to the DBDE.

The bioinformatics search parameters included a stem region ranging from 8-15 base pairs and the loop ranging from 6-15 nucleotide positioned in the proximity of the 5´ cap.

Two hundred thirty nine mRNAs with the potential of forming a stem-loop structure starting within 10nt of the 5′ termini were identified. Each mRNA was subsequently subjected to MFold prediction to determine which mRNAs are likely to form the stem- loop structure within the context of the entire 5′UTR. Three representative examples were chosen for further analysis. The Hip1 and Kcnj2 mRNA 5´ ends have the potential

of forming a DBDE-like stem-loop structure and expected to be preferentially decapped

Figure 12. The Ndufb7 5´ UTR stem-loop structure is also capable of enhancing decapping.

(A) A schematic of the Ndufb7 5′UTR predicted secondary structure is shown with the numbers denoting the nucleotides from the 5′ end.

(B) Decapping assays were carried out with the indicated Ndufb7 RNAs and Stx7 negative control. WT: wild type; Mut35-44: mutations in nucleotide 35-44 to disrupt the second strand of the stem; Mut2-11/35-44: complementary mutations in both strands of the stem that maintain the same structure. Decapping efficiency quantitations for each

RNA obtained from three independent experiments are shown with the decapping efficiency of wild type Ndufb7 RNA arbitrarily set to 100.

by Dcp2. The third RNA, Zcrb1, also has the propensity to form a stem-loop structure,

but the stem region is expected to be positioned 10 nucleotides from the 5´ end. Based on

our analysis in Figure 11, this RNA should not be recognized as a Dcp2 substrate and is not expected to be efficiently decapped. The DNA corresponding to the first 150

nucleotides of each mRNA was transcribed in vitro with T7 polymerase, cap labeled with

32P and tested in an in vitro decapping assay with Dcp2. As expected, the Hip1 RNA was

decapped at an efficiency three fold greater than the Stx7 control RNA (Figure 13A,

compare lane 1-3 to 10-12). Similarly, the Kcnj2 RNA was two fold more efficiently

decapped (lanes 4-6). Interestingly, the Zcrb1 RNA which is predicted not to be

preferentially decapped, was decapped at a level comparable to that of the Stx7 negative

control (lane 7-9).

To further substantiate that the increase in decapping observed with the Hip1

RNA was a consequence of the stem-loop structure, the putative stem structure was

mutated. To minimize alterations of primary sequence adjacent to the 5´ cap, mutations

were introduced in the cap distal strand of the stem at nucleotides 23-31 (Figure 13B).

As shown in Figure 13C, this mutation significantly reduced the decapping activity by

about 50 percent (lane 4-6). Compensatory changes in nucleotides 2-11 designed to

reform base pairing and formation of the stem also restored decapping activity to 80

percent of the wild type level, indicating the intact stem-loop structure is important for the enhancement of decapping in this RNA as well. Collectively, our data indicates Dcp2 can recognize and preferentially decap mRNAs containing a stem loop structure at their

5´ end.

Figure 13. Enhancement of decapping by DBDE-like 5′ end stem-loop structure in

cellular mRNAs.

(A) Decapping assays were carried out with Hip1, Kcnj2 and Zcrb1 RNAs which are all

predicted to contain potential stem-loop structures at their 5′ end, and the Stx7 RNA.

Decapping efficiency of Stx7 RNA was arbitrarily set to 1 and the fold increase relative to Stx7 decapping is plotted. Quantitations were obtained from three independent experiments and the Standard Deviations (SD) are denoted by the error bars.

(B) Schematic of the Hip1 5´ UTR wild type and substitution mutations are shown. WT: wild type; Mut23-31: mutations in nucleotide 23-31 to disrupt the second strand of the stem; Mut4-13/23-31: complementary mutations in both strands of the stem that maintain the same structure.

PEI-TLC. Quantification of the decapping efficiency for each RNA obtained from three independent experiments are shown on the right with the decapping efficiency of wild type Hip1 RNA arbitrarily set to 100.

Discussion

In this chapter, we demonstrate that the Dcp2 decapping enzyme can selectively bind to a subset of mRNAs and identified the Rrp41 mRNA as a substrate that’s specifically bound by Dcp2. This binding is direct as bacterial expressed recombinant

Dcp2 can copurify this RNA without additional proteins. Furthermore, this RNA is preferentially decapped by Dcp2, most likely because of its specific binding by Dcp2.

The 5′UTR of Rrp41 was identified as an autonomous element for stimulating Dcp2 decapping. The maximal stimulatory sequence is contained within the first 60 nucleotides of the 5′UTR (2xDE). Moreover, we demonstrate that the Rrp41 mRNA is a true target of Dcp2 in cells, and its stability is regulated by Dcp2. Mutational assay of the 2xDE element identified the Dcp2 binding region as the first 33 nucleotides termed the Dcp2

Binding and Decapping Element, DBDE, which forms a stable stem-loop structure.

Importantly, the intact stem-loop secondary structure but not the primary sequence was responsible for enhanced Dcp2-directed decapping. Furthermore, a bioinformatics search for mRNAs that harbor a DBDE-like stem-loop in the 5′UTR identified a subset of mRNA with the propensity to form such a structure. Interestingly, the tested RNAs with a DBDE-like element positioned within 10 nucleotides of the 5´ cap were preferentially decapped, indicating that the DBDE 5′ end stem-loop structure could be a general predictor of mRNAs that could be specifically regulated by Dcp2 decapping.

Microarray analysis of Dcp2-bound mRNA revealed RNA profiles that were associated with Dcp2. A small subset of 98 mRNAs were identified to be bound by Dcp2 at a level six fold or higher than that bound to a control unrelated testis-specific RNA- binding protein, mDazl (Table IV). These RNAs derived from a wide range of functional

classes, including cell metabolism, electron transport, transcription factors, and mRNA

encoding translation factors. In Dcp2 knockout plants, the levels of a set of mRNAs are significantly elevated, among those the best represented are the heat-shock proteins

(Goeres et al, 2007). And in a recent report, degradation of HSP70 mRNA in Drosophila requires Dcp2 decapping (Bonisch et al, 2007). In our microarray, mRNAs of several heat shock proteins, including HSP70, HSP90B1, DNAJB12 (Hsp40 homolog), were found to be associated with Dcp2, approximately 5 fold more than the control mDazl protein (data not shown). Collectively, these may indicate a conserved function of Dcp2 in regulating mRNA stability.

RNAs can be associated with Dcp2 either directly or indirectly. Given that Dcp2 can form different complexes with various protein factors, it is conceivable that many of the mRNAs that specifically associated with Dcp2 were through indirect binding via other protein(s) that can recruit Dcp2. For example, TTP can bind an ARE-containing mRNA and recruit a decapping complex including Dcp2 (Fenger-Gron et al, 2005b;

Lykke-Andersen & Wagner, 2005). Similarly, mRNAs harboring premature termination codons can be recognized by Upf1 and recruit an Dcp2 decapping complex (Lykke-

Andersen, 2002). In yeast, the Rps28 protein can bind its own mRNA and engage the

Dcp1p/Dcp2p decapping complex via Edc3p (Badis et al, 2004). Alternatively, the RNA

binding property of Dcp2 suggests that it can bind its substrates directly without

additional factors. In this study, we demonstrate Dcp2 can directly bind the Rrp41 mRNA and decap it specifically, indicating Dcp2 alone has preference for RNA substrates. This

is similar to the observation with the X29 nuclear decapping enzyme, which alone can

bind U8 snoRNA and preferentially decap it (Ghosh et al, 2004). It is interesting that

specificity of X29 is dependent on cationic ions. In the presence of Mg2+, it preferentially

decapped the U8 snoRNA, while in the presence of Mn2+, it decapped all RNAs tested

(Peculis et al, 2007). However, this does not appear to be true for Dcp2. Similar specific

decapping of Rrp41 mRNA by Dcp2 was observed regardless of whether Mg2+ or Mn2+

was used as the divalent cation (data not shown).

The 2xDE element, consisting of the first 60nt of the Rrp41 mRNA 5′UTR is an element that is specifically bound by Dcp2 and can enhance the decapping of a heterologous RNA when placed at the 5´ end. The element contains two stretches of similar sequences from nucleotides 1 to 25 and 26 to 60. We have refined these findings

to show that the first 33 nucleotides of the 2xDE termed DBDE are sufficient for the

Dcp2 recognition and enhanced decapping (Figure 7A). The functional significance of

the second element is currently unclear since its positioning approximately 30 nucleotides

from the 5´ cap would be expected to not promote decapping. Whether it can serve to

augment the recruitment of Dcp2 in cells at its endogenous position 30 nucleotides distal

to the cap remains to be determined.

RNA binding proteins can recognize their RNA substrates by either the primary

sequence or the higher order structure of the RNA. Binding of the Iron Response

Element (IRE)-binding protein (IRP) to the IRE is one example of the importance of

structural integrity to RNA binding. IRP binding to the ferritin mRNA 5´UTR is

dependent on the IRE stem-loop structure (Hentze & Kuhn, 1996). Similarly, the SECIS-

binding proteins recognize a SECIS element which is a structural motif that directs the

cell to translate UGA codons as selenocysteines (Walczak et al, 1996; Copeland et al.

2001). In this study, we found that the intact stem-loop structure within the 5′ end of the

Rrp41 mRNA 2xDE element is critical for promoting Dcp2 decapping. The Dcp2 protein appears to recognize the secondary structure of its target RNAs, but not the primary sequence, since alterations in the primary sequence of the stem-loop structure did not have a significant effect on Dcp2 decapping as long as the structural integrity of the stem loop was maintained (Figure 9, mutant C and D).

The stem-loop structure within the first 33nt of 2xDE consists of a double- stranded stem region of 11 base pairs with a single nucleotide bulge at position 29, and a single stranded loop region of 8 nucleotides. Mutational assays in the stem-loop region in the 2xDE indicated that the exact nucleic acid sequences in the stem or in the loop region are not important for Dcp2 decapping. The single nucleotide bulge is also dispensable for

Dcp2 decapping (Figure 10, mutant H). Increasing the size of the loop to 12 nucleotides

(Figure 10, mutant G) resulted in a slight increase in decapping activity, while decreasing the size of the loop to 4 nucleotides (Figure 10, mutant F) modestly decreased the decapping activity, suggesting the size of the loop does not significantly contribute to the promotion of decapping. In contrast to the loop, the length of the stem was important to decapping. Decreasing the size of the stem region to less than 8 basepairs resulted in a significant reduction of decapping activity, indicating either the Dcp2 protein requires more than 7 base pairs to recognize the RNA or shortening the stem compromises the structural integrity of the overall stem-loop structure.

We previously demonstrated that Dcp2 is an RNA-binding protein (Piccirillo et al,

2003) and proposed a model in which the RNA-binding property anchors Dcp2 onto a capped RNA substrate adjacent to the cap structure. The presence of a DBDE stimulatory element at the 5′ termini of Rrp41 mRNA is consistent with this model. Furthermore, the

fact that placing the DBDE element 10 nucleotides downstream of the 5′ cap diminished decapping stimulation and reduced its capacity to form the Dcp2 complex observed on the wild type 2xDE RNA (Figure 11) indicates the distance between these two elements is important. The significance of the cap proximal 5´ region is also important for the yeast

Dcp2p protein, which can be inhibited by annealing antisense oligonucleotides to the 5′ end of its RNA substrate (Steiger et al, 2003). Further significance for the 5´ terminal sequence is apparent in C. elegans, in which the presence of a 22 nucleotides spliced leader sequence at the 5′ end of an RNA dramatically inhibits Dcp2 decapping activity

(Cohen et al, 2005). Interestingly, the splice leader (SL) RNA, which functions as the donor of the 22 nucleotides spliced leader sequence, is efficiently decapped by Dcp2

(Cohen et al, 2005). Consistent with our study, the initial 30 nucleotides of the SL RNA can fold into a stem-loop structure within the context of the SL RNA (Denker et al, 1996) which is likely disrupted in the trans spliced mRNAs that only contain the first 22 nucleotides of the SL RNA. In addition, all trypanosome mRNAs contain a 39 nucleotide spliced leader sequence at their 5΄ end and this sequence has been shown to inhibit Dcp2- like decapping activity in trypanosome extract (Milone et al, 2002). Although this region has been proposed to adopt two forms of secondary structures containing an internal stem-loop (Harris et al, 1995), the proposed stem-loop structures are more than 10 nucleotides downstream of the 5´ cap which might explain their poor decapping propensity. Collectively these data support direct binding of Dcp2 to the 5′ end of its

RNA substrate to hydrolyze the cap structure and an RNA containing a stem-loop structure preferentially bound by Dcp2 at its 5´ end, is decapped more efficiently.

Consistent with the data that Dcp2 preferentially decapped Rrp41 mRNA in vitro, the stability of Rrp41 mRNA is regulated by Dcp2 in cells. The fact that Rrp41 is an

essential component of the exosome complex (Liu et al, 2006) is intriguing. It suggests

Dcp2, which is a key component in the 5′ to 3′ RNA decay machinery, can also impact the 3′ to 5′ RNA decay machinery. Even though the exosome is a multisubunit complex,

Rrp41 is an important structural subunit and its level would affect the whole complex, since the knockdown of hRrp41p led to reduced levels of other core exosome subunits

(van Dijk et al, 2007). Such a regulatory mechanism could provide an interesting

regulation feedback between 5′ to 3′ and 3′ to 5′ RNA decay machineries.

A search of all available human RNA 5´ UTRs in the 5´ UTR database as of 2008

(see Materials and Methods) revealed 239 mRNAs containing the potential to form a

DBDE-like structure starting within the first 10 nucleotides from the 5′ end of the mRNA.

Although the search parameters identify local regions of the RNA capable of forming the

stem-loop structure, further analysis of each mRNA within the context of the entire 5´

UTR by MFold program, predicted approximately twenty percent of these mRNAs could

form a stable stem-loop structure at the 5´ end. Therefore, only a small subset of mRNAs

appear to have the capacity to form into an appropriate structure that would render them

as direct substrates of Dcp2 decapping in cells. One important observation was that a

small subset of mRNAs that copurified with Dcp2 in the microarray (Table IV) were

identified by the bioinformatic search, indicating only a small portion of Dcp2-bound

mRNAs contain a DBDE-like 5' stem-loop structure. One explanation is that as Dcp2

could form complexes with many interacting proteins, many of the Dcp2-associated

mRNAs indirectly copurified with Dcp2 by binding to these interacting proteins.

Therefore these mRNAs are likely not direct targets of Dcp2 and do not necessarily have the 5' stem-loop structure. Secondly, the bioinformatic search was carried out using the

stem-loop structure of Rrp41 mRNA as a prototype. Some Dcp2 target mRNAs may have

different stem-loop structures that are not included in the bioinformatic search. Of the

three mRNAs we further characterized, Hip1 RNA was the most efficiently decapped and

contains a 13-nucleotide stem with a single nucleotide bulge and a nine-nucleotide loop

region. The Kcnj2 RNA contains a nine-nucleotide stem with one mismatch and an 8-

nucleotide loop. It had a modest 2 fold increase in decapping activity compared to the

Stx7 RNA negative control. The Zcrb1 RNA contains a nine nucleotide stem and a seven

nucleotide loop but was located 10 nucleotides downstream of the cap structure and as

expected was not decapped at a level greater than that observed with the negative control.

These data indicate that a general stem-loop structure formed at the 5′ terminus of an

RNA, of which the primary sequence and the composition may vary, is capable of

promoting Dcp2 decapping. Furthermore, this feature can be used to predict potential

Dcp2 target RNAs in cells. However, considering multiple positive and negative

regulators of Dcp2 decapping have been reported (Coller & Parker, 2004), it is likely that

decapping is regulated by the combined total contributions of both the direct intrinsic

properties of Dcp2 to associate with the 5´ cap as well as the decapping auxiliary proteins

that will dictate the consequence of Dcp2 on an mRNA. Our studies now provide a

framework for future studies addressing the direct and indirect decapping activities of

Dcp2 on mRNAs in cells.

Chapter II: Differential Utilization of Decapping Enzymes in

Mammalian mRNA Decay Pathways

Summary

mRNA decapping is a crucial step in the regulation of mRNA stability and gene

expression. Dcp2 is an mRNA decapping enzyme that has been widely studied. Here we

identify the presence of a second mammalian cytoplasmic decapping enzyme, Nudt16. In this chapter we address the differential utilization of the two decapping enzymes in specified mRNA decay processes. Using mouse embryonic fibroblast (MEF) cell lines derived from a hypomorphic knockout of the Dcp2 gene with undetectable levels of Dcp2 or MEF cell lines harboring a Nudt16-directed shRNA to generate reduced levels of

Nudt16, we demonstrate the distinct roles for Dcp2 and Nudt16 in nonsense mediated mRNA decay (NMD), ARE-mediated decay and miRNA mediated silencing. Our results indicated that NMD preferentially utilizes Dcp2 rather than Nudt16; Dcp2 and Nudt16 are redundant in miRNA mediated silencing; and Dcp2 and Nudt16 are differentially utilized for ARE-mRNA decay. These data demonstrate that the two distinct decapping enzymes can uniquely function in specific mRNA decay processes in mammalian cells.

Introduction

Regulation of RNA degradation plays an important role in the control of gene

expression. In eukaryotic cells, most mRNAs have a 5′ monomethyl guanosine cap

structure (Shatkin, 1976) and a 3′ poly(A) tail (Sachs, 1993) which are important for

mRNA translation and stability. Bulk mRNA decay usually initiates with the removal of

the 3′ poly(A) tail (Decker & Parker, 1993). The resulting deadenylated mRNAs can be

degraded by two exonucleolytic pathways involving either 5′-end or 3′-end decay (Badis

et al, 2004; Cougot et al, 2004). The 3′-end decay is carried out by a cytoplasmic

multisubunit exosome complex (Anderson & Parker, 1998; Mitchell et al, 1997) and the

resulting cap dinucleotide subsequently hydrolyzed by the scavenger decapping enzyme,

DcpS (Liu et al, 2002; Wang & Kiledjian, 2001). Alternatively, following deadenylation, the mRNA can be decapped to remove the m7GDP of the cap to expose a

monophosphorylated 5´ end, which is subsequently degraded by the 5´ monophospate-

dependent 5′ to 3´ exoribonuclease, Xrn1 (Decker & Parker, 1993; Hsu & Stevens, 1993).

Removal of the 5´cap structure (decapping) is therefore an important prerequisite for

decay of the mRNA body from the 5´ end. The Dcp2 protein has been identified as the

major mRNA decapping enzyme in cells (Dunckley & Parker, 1999; Lykke-Andersen,

2002; van Dijk et al, 2002; Wang et al, 2002b).

The Dcp2 decapping enzyme catalysis has been implicated in various mRNA decay processes, including nonsense-mediated mRNA decay (NMD), ARE-mediated decay and microRNA directed gene silencing (Franks & Lykke-Andersen, 2008). The

NMD pathway, which targets mRNAs with premature translational termination for rapid

decay, was reported to trigger decapping in both yeast and human cells (Couttet &

Grange, 2004; Lejeune et al, 2003; Muhlrad & Parker, 1994), and a reduction of Dcp2

levels in Hela cells impaired both nuclear-associated and cytoplasmic NMD (Lejeune et

al, 2003). All three Upf proteins, key factors in NMD, coimmunopurified with Dcp2 (He

& Jacobson, 1995; Lejeune et al, 2003; Lykke-Andersen, 2002), indicating that these

factors recruit decapping protein to degrade nonsense-containing mRNAs.

AU-rich element (ARE)-mediated decay rapidly degrades mRNAs that contain

AREs in their ′3 untranslated region (UTRs). Different A REs seem to differentially

activate both 5′ to 3′ and 3′ to 5′ decay pathways (Chen et al, 2001; Gao et al, 2001;

Lykke-Andersen & Wagner, 2005; Mukherjee et al, 2002; Murray & Schoenberg, 2007;

Stoecklin et al, 2006). TTP, a key ARE binding factor, interacts with the Dcp2 protein and enhances decapping of a target ARE-containing RNA in vitro (Fenger-Gron et al,

2005a). Depletion of the decapping activator Lsm1 by siRNA, impairs ARE-mediated decay in human cells (Stoecklin et al, 2006). Finally, decapping has also been implicated in miRNA-mediated gene silencing. Knockdown of Dcp1:Dcp2 decapping complex effectively relieved miRNA-mediated silencing of firefly luciferase expression in

Drosophila cells (Rehwinkel et al, 2005). Depletion of several decapping activators including Hedls (also referred to as Ge-1 and Edc4), Dcp1a, Edc3 and Lsm1 in

Drosophila S2 cells suppressed gene silencing mediated by several miRNAs (Eulalio et al,

2007b). Moreover, the decapping activator RCK/p54 has also been shown to be important in miRNA-mediated silencing in human cells (Chu & Rana, 2006). Furthermore, Dcp2 can preferentially bind to a subset of mRNA substrates (Li et al, 2008). Highly transcript- specific decapping of Dcp2 has been reported in vitro and in mammalian cells (Li et al,

2009; Li et al, 2008). Collectively, it appears that in addition to a potential general

mRNA decay function, Dcp2 can also target various specific mRNA decay processes.

One striking feature of the decapping machinery is that the decapping enzyme

Dcp2 and other decapping activators including Dcp1a, Edc3, Hedls, Lsm1-7 complex, and the 5′ to 3′ exoribonuclease, Xrn1 colocalize to distinct cytoplasmic foci called P - bodies (Fenger-Gron et al, 2005a; Franks & Lykke-Andersen, 2008; Sheth & Parker,

2003; van Dijk et al, 2002; Yu et al, 2005). In addition, proteins involved in NMD, ARE-

mediated decay, and miRNA mediated silencing machineries localize to P-bodies (Ding

et al, 2005; Eystathioy et al, 2003; Franks & Lykke-Andersen, 2007; Sen & Blau, 2005;

Sheth & Parker, 2006). Moreover, mRNA decay intermediates and translational repressed

mRNAs can also be found in P-bodies (Franks & Lykke-Andersen, 2007; Liu et al, 2005b;

Sheth & Parker, 2003; Sheth & Parker, 2006) suggestive of a role for P bodies in mRNA decay. However, loss of visible P-bodies do not lead to detectable alterations in mRNA decay (Decker et al, 2007; Eulalio et al, 2007a; Stalder & Muhlemann, 2009) and decapping can occur within polysomes that are devoid of visible P-bodies (Hu et al,

2009); therefore the assembly of P-bodies does not seem to be necessary for mRNA degradation. The importance for the decapping machinery to form cytoplasmic foci still remains to be elucidated.

Nudt16 was initially identified in Xenopus as a U8 snoRNA binding protein, termed X29, and shown to possess decapping activity (Ghosh et al, 2004). X29 is a nucleolar protein capable of specifically binding and decapping the U8 snoRNA in vitro in the presence of Mg2+ although interestingly possessed a more pleiotropic decapping

activity when Mn2+ was the cation source (Ghosh et al, 2004). Although X29 has been

implicated in nucleolar decapping, a direct role for this protein in cellular U8 snoRNA stability has yet to be addressed. The Nudt16, mammalian ortholog of X29, also possesses decapping activity (Taylor & Peculis, 2008) and has been proposed as a nucleolar decapping enzyme. Interestingly although conserved in metazoans, an obvious ortholog of Nudt16 is lacking in S. cerevisiae, C. elegans and Drosophila (Taylor &

Peculis, 2008). Here we demonstrated that the mammalian homolog, Nudt16, is a cytoplasmic decapping enzyme, and we used immortalized mouse embryonic fibroblast

(MEF) cell lines with reduced levels of Dcp2 or Nutd16 to study the differential utilization of the two decapping enzymes in NMD, ARE-mediated decay and miRNA- mediated silencing. Our results showed that Dcp2 was preferentially used in NMD; Dcp2 and Nudt16 are redundant in miRNA mediated silencing; Dcp2 and Nudt16 are differentially utilized for the decay of ARE-containing mRNAs; and loss of Dcp2 or

Nudt16 did not appear to impact P-body assembly. These data suggest decapping enzymes could be differentially utilized for different cellular mRNA decay processed in mammalian cells.

Results

Nudt16 is a cytoplasmic decapping enzyme in mammalian cells.

Since the X29 protein was reported to be a decapping enzyme, the ability of its mammalian homolog Nudt16 to decap mRNA was tested. His-tagged recombinant proteins were generated and the extent of decapping detected by thin layer chromatography (TLC). As expected, Nudt16 contains decapping activity (Figure 14A) while decapping was not detected with the Nudt16-like 1 (Nudt16L1) protein negative control, which is highly similar to Nudt16 but lacks the critical cation binding residues

(Taylor & Peculis, 2008).

Although the Xenopus homolog of Nudt16 has been reported to be a nucleolar protein (Ghosh et al, 2004), we next asked whether the same is true in mammalian cells.

Human Nudt16 protein containing a Flag epitope tag at the amino terminus was expressed in U2OS cells and its localization determined by immunofluorescence with an anti-Flag antibody by confocal microscopy. Flag-tagged human Nudt16 was localized primarily to the cytoplasm (Figure 14B). Similar results were also obtained when Nudt16 containing the epitope tag at the carboxyl terminus was tested (Figure 14B), indicating that Nudt16, unlike its Xenopus counterpart, is predominantly a cytoplasmic protein.

Collectively, the observations that Nudt16 contains decapping activity and is contained in the cytoplasm suggest Nudt16 is a second cytoplasmic mRNA decapping enzyme.

Establishment of Dcp2 and Nudt16 knockdown MEF cell lines

We generated mice containing a homozygous insertion of the β-galactosidase-

Figure 14. Nudt16 is a cytoplasm mRNA decapping enzyme.

(A) Decapping activity of the recombinant His tagged human Dcp2, Nudt16 and mouse

Nudt16-like 1 proteins are shown. Decapping products resolve on PEI-TLC developed in

0.45 M (NH4)2SO4.

(B) FLAG-hNudt16 (Top) or hNudt16-FLAG (Bottom) was overexpressed in U2OS cells.

The epitope tagged protein was localized by indirect immunofluorescence with an anti-

FLAG antibody by confocal microscopy. The presented images are representative of

>95% of the transfected cells. The Differential Interference Contrast (DIC) images of the

same cells are shown as indicated.

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neomycin resistance fusion gene (β-geo) within intron 1 of the mouse Dcp2 loci

(Dcp2β/β;(Song et al, 2010)). However, the mice are hypomorphic for Dcp2 production

likely due to background alternative splicing that bypasses the β-geo gene to generate low levels of Dcp2 protein despite the homozygous insertion of the β-geo gene. To characterize the role of Dcp2 in distinct mRNA decay processes, we utilized SV40 large

T antigen immortalized mouse embryonic fibroblast (MEF) cells obtained from wild type or Dcp2β/β embryos which contain detectable and undetectable Dcp2 protein levels

respectively (Song et al, 2010). The extensively reduced levels of Dcp2 protein in these

cells provided us with a Dcp2 knock down cell line to assess the significance of Dcp2 in

mRNA decay. No morphological changes were obvious between the wild type and

Dcp2β/β MEF cell lines except for a slow growth phenotype observed in the Dcp2β/β cells

(data not shown).

To investigate the role of both the Nudt16 and Dcp2 decapping proteins in

cellular mRNA decay, we knocked down Nudt16 mRNA and corresponding protein

levels in wild type and Dcp2β/β MEF cells using a lentiviral shRNA expression system.

Greater than 95% reduction of Dcp2 protein level was observed in the Dcp2β/β MEF cells

and approximately 70 % and 90 % reduction of Nudt16 was observed in the Nudt16

knockdown cells within the wild type or Dcp2β/β MEF cells respectively (Figure 15).

These Dcp2 and Nudt16 individual and double knockdown MEF cell lines were used in the subsequent analyses to assess the significance of Dcp2 and Nudt16 in mRNA decay.

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Figure 15. Reduction of Nudt16 protein levels in wild type and Dcp2β/β MEFs.

Extract from immortalized clonal MEF cell lines derived from wild type embryos, or

embryos with a homozygous insertion of the β-galactosidase-neomycin resistance fusion

gene (β-geo) within intron 1 of the mouse Dcp2 loci (Dcp2β/β) expressing lentiviral directed empty vector control (lanes 1 and 3) or Nudt16-specific shRNA (lanes 2 and 4)

(Song et al, 2010) were used in Western blot analysis. Bands corresponding to Dcp2,

Nudt16 or the GAPDH loading control protein are indicated on the right.

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Loss of Dcp2 and Nudt16 had no apparent impact on P-bodies in MEF cells

In addition to a dispersed cytoplasmic localization, Dcp2 protein also localizes in

cytoplasmic foci termed P-bodies (Sheth & Parker, 2003; van Dijk et al, 2002). The

function of P bodies is still unclear although they have been postulated to be sites of

mRNA decay, mRNP storage and miRNA-directed mRNA decay (Franks & Lykke-

Andersen, 2007; Franks & Lykke-Andersen, 2008; Liu et al, 2005b; Sheth & Parker,

2003; Sheth & Parker, 2006). Disruption of the Dcp2 gene in yeast results in an increase

in the size and number of P bodies (Teixeira & Parker, 2007), believed to be a

consequence of aberrantly uncapped mRNA-containing mRNPs that aggregate. However,

surprisingly, siRNA-directed knockdown of Dcp2 in mammalian cells did not lead to

altered P body states (Stoecklin et al, 2006; Yu et al, 2005). Consistent with these

previous observations, the virtually complete knock down of Dcp2 in mammalian cells

also did not reveal appreciable differences in P bodies within the Dcp2β/β MEF cells.

Wild type and Dcp2β/β MEF cells were mixed and cultured on the same coverslip to ensure similar cell confluency and growth stage. P-bodies were stained with antibodies

directed to either Dcp2 or Dcp1a. As expected, both proteins colocalized within P body

foci in the wild type MEF cells and foci were not detected with the Dcp2-specific antisera

in the Dcp2β/β cells (Figure 16A). The number of P-bodies, as determined by Dcp1a localization, remained comparable regardless of the presence or absence of Dcp2, with

20.2 foci and 21.1 foci per cell respectively. Analysis of P-bodies in cells expressing reduced levels of Nudt16 protein was also carried out. The number of P bodies, as determined by anti-Dcp1a immunofluorescence analysis, remained the same in MEF cells with reduced Nudt16 protein or a combination of both Dcp2 and Nudt16 knockdown

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Figure 16. P-body numbers are unaltered in MEF cells with undetectable levels of

Dcp2 and/or reduced levels of Nudt16 protein.

(A) Wild type and Dcp2β/β MEF cells were simultaneously cultured on the same cover slide and distinguished by immunofluorescence using affinity purified Dcp2 specific antibody detected with FITC conjugated secondary antibody. Dcp1a was used as a P- body marker and visualized by an anti-Dcp1a antibody detected with Texas Red conjugated secondary antibody. The number of P-bodies per cell, as determined by

Dcp1a positive foci, were counted and the average numbers obtained from 30 cells are graphed on the right with standard deviations indicated.

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(B) P bodies were visualized by immunofluorescence with anti-Dcp1a antibody detected with FITC conjugated secondary antibody and quantitated as described in (A) except wild type or Dcp2β/β MEF cells expressing a control vector or Nudt16 specific shRNA were

used as indicated.

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(Figure 16B). These data indicate that formation of P-bodies appears independent of both

Dcp2 and Nudt16 decapping enzymes in mammalian cells.

Preferential utilization of Dcp2 in NMD

The demonstrations that both Dcp2 and Nudt16 possess transcript specific decapping properties (Ghosh et al, 2004; Li et al, 2009; Li et al, 2008) prompted us to test whether various mRNA decay processes differentially utilized the two decapping enzymes and assessed their roles in: nonsense-mediated decay (NMD), ARE-mediated decay and noncoding RNA-mediated gene silencing. The effect of Dcp2 on the reduction of nonsense mRNA levels was assessed by transiently transfecting MEF cells with plasmids encoding either a normal β-globin transcript lacking a nonsense mutation

(Norm) or its nonsense mutation-containing version (PTC39). A Renilla luciferase plasmid was cotransfected for normalization of transfection efficiencies. Total RNA was isolated 24 h post-transfection, reverse transcribed, and analyzed by quantitative real-time

PCR. A comparison of nonsense-containing transcript levels to that of the nonsense- lacking transcript levels are presented in Figure 17A. Consistent with Lejeune et al.,

(Lejeune et al, 2003), PTC39 mRNA levels were significantly reduced in the wild type

MEF cells with 43% of the mRNA remaining relative to Norm mRNA. In contrast, a statistically significant increase in PTC39 mRNA levels was detected in Dcp2β/β MEF

cells where 64% of mRNA remained relative to Norm mRNA levels.

Analysis of the same reporter constructs was also carried out in MEF cells

containing reduced levels of Nudt16. Interestingly, a reduction of Nudt16 levels did not

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Figure 17. Contribution of decapping enzymes to NMD.

(A) mRNA levels derived from transgenes expressing a β-globin transcript (Norm) or nonsense codon-containing β-globin transcript (PTC39) in wild type and Dcp2β/β MEF cells transduced with or without Nudt16 specific shRNA or in wild type MEF cells transduced with Upf1 shRNA expressing virus as indicated, were determined by quantitative real-time PCR. mRNA levels were determined 24h post-transfection and normalized to Renilla luciferase mRNA levels expressed from a cotransfected plasmid.

The normalized mRNA value for the sense β-globin transcript was arbitrarily set to 100%.

Average of three independent experiments is shown with standard deviation indicated.

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Asterisks denote p<0.05 (ANOVA followed by Tukey-Kramer Multiple Comparisons

Test).

(B) The level of TCR-β mRNA (Norm) or nonsense codon-containing TCR-β transcript

(PTC68) were determined as described in (A) above. The normalized mRNA value for the sense TCR-β transcript was arbitrarily set to 100%. Average of three independent experiments are shown with standard deviation indicated. Asterisks denote p<0.05 and double asterisks denote p<0.01 (ANOVA followed by Tukey-Kramer Multiple

Comparisons Test).

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result in a detectable difference in PTC39 mRNA levels compared to that observed in

control knockdown cells (Figure 17A). Similarly, knockdown of Nudt16 in the Dcp2β/β

cell background also did not result in altered PTC39 mRNA levels and were

indistinguishable from levels obtained with reduced Dcp2 in the Dcp2β/β cells. Similar

results were observed using a PTC-containing TCR- β reporter. MEF cells were transfected with plasmids encoding a normal TCR-β transcript (Norm) or its nonsense mutation-containing version (PTC68). A two-fold increase in PTC68 mRNA levels was detected in Dcp2β/β MEF cells relative to Norm mRNA, while a reduction of Nudt16,

either in the wild type or in the Dcp2β/β MEF background, did not significantly alter

PTC68 mRNA levels (Figure 17B). Knockdown of Upf1, an NMD core factor, was used

as a positive control for NMD directed decay of each PTC transcript. However, these are

likely underestimation since we were only able to obtain 65% knockdown of Upf1 (data

not shown). Collectively, these data demonstrate that Dcp2 is preferentially utilized over

Nudt16, in the demise of these two PTC-containing transcripts.

Redundancy of Dcp2 and Nudt16 in miRNA-mediated gene silencing

Dcp2 has been implicated in miRNA-mediated gene silencing, where a knockdown of the Dcp1:Dcp2 decapping complex impaired miRNA-mediated silencing of firefly luciferase expression in Drosophila cells (Rehwinkel et al, 2005). To determine the significance of Dcp2 in miRNA and siRNA mediated gene silencing in mammalian cells, we assessed the level of Renilla luciferase activity from transcripts containing or lacking miRNA target sites. Wild type or Dcp2β/β MEF cells were transiently transfected

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with plasmids encoding Renilla luciferase lacking an miRNA target sequence in the

3´UTR or containing either a let-7 miRNA target sequence (3X let-7 bulge) or a let-7

perfectly complementary sequence. The three reporter constructs were each co-

transfected with a plasmid expressing the pri-let7 miRNA and firefly luciferase plasmid for normalization of transfection efficiencies. Cells were harvested and assayed for luciferase activity 24h post-transfection. Renilla luciferase activities are presented in

Figure 18A with the activity obtained from the reporter lacking a let-7 target sequence arbitrarily set to 100%. Interestingly, significant differences were not detected in Renilla luciferase activities with either the let-7 bulged or perfect match sites when comparing the results of wild type and Dcp2β/β MEF cells. An 89% reduction in luciferase activity

was observed in wild type MEF cells while an 87% reduction was detected in the Dcp2β/β

cells when assaying luciferase activities from transcripts harboring a bulged let-7 target

site. Similarly, a 95% and 94% reduction in luciferase activity were detected in the wild

type and Dcp2β/β cells respectively when assaying luciferase activities from transcripts

harboring a perfect match to the let-7 site. Therefore, in contrast to the situation in

Drosophila where Dcp2 was found to contribute to miRNA directed silencing, the almost

complete absence of Dcp2 does not appear to have a significant impact on miRNA or

siRNA mediated gene silencing with this reporter system in mammalian cells.

To assess the role of Nudt16 or the combined effect of Dcp2 and Nudt16, the

same reporter constructs were tested in Nudt16 knockdown MEF cells. As in Figure 18A,

the reduction of Nudt16 alone had no detectable consequence on Renilla luciferase

activity with either the let-7 bulged or perfectly matched sites. Importantly, although a

reduction of either decapping enzyme individually had no consequence, a double knock-

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Figure 18. Dcp2 and Nudt16 are redundant in miRNA mediated gene silencing.

(A) Plasmids expressing mRNAs encoding the Renilla luciferase with three let-7 miRNA

bulge sites, a let-7 perfectly complementary site, or no let-7 site in the 3´ UTR were

transfected into the indicated MEF cell lines and luciferase activity was analyzed 24h

post-transfection. Renilla luciferase activity was normalized to Firefly luciferase activity

derived from a cotransfected plasmid. The normalized value of Renilla luciferase with no

let-7 site was arbitrarily set to 100%. The average of three independent experiments is shown with standard deviation indicated by the error bars. Cell line nomenclature is as described in the legend to Figure 15. An increase in Renilla luciferase activity was

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detected only in cells with a reduction of both Dcp2 and Nudt16 protein. Asterisks denote

p<0.01 (ANOVA followed by Tukey-Kramer Multiple Comparisons Test).

(B) A similar analysis as that presented in (A) is shown except the Dcp2 and Nudt16 double knockdown was complemented with an shRNA-resistant Nudt16 expression construct. Luciferase activity was normalized as in (A) and the average of three independent experiments is shown with standard deviation. Asterisks denote p<0.02

(ANOVA followed by Tukey-Kramer Multiple Comparisons Test).

(C) Renilla luciferase mRNA levels from an analysis as presented in (A) above were determined by real-time quantitative PCR and normalized to mRNA levels derived from a cotransfected Firefly luciferase expression construct. The average of three independent experiments is shown with standard deviation indicated by the error bars.

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down of both Nudt16 and Dcp2 significantly increased Renilla luciferase activity of the

reporter construct containing the let-7 bulged sites. In contrast to the 11% of renilla activity detected in the control virus infected cells containing wild type levels of both

Dcp2 and Nudt16, the activity was increased approximately three fold to 31% in the

double knockdown conditions (Figure 18A). Importantly, this increase can be reversed by

overexpression of Nudt16 (Figure 18B) indicating the presence of at least one of the

decapping enzymes is sufficient to promote maximal silencing. Interestingly, Renilla

luciferase activity of the reporter construct containing a let-7 perfect match target site

showed no significant change despite the low level of both Dcp2 and Nudt16. These data

indicate that although Dcp2 and Nudt16 do not contribute to siRNA-directed gene silencing in our assays, they demonstrate a redundant contribution to miRNA directed gene silencing where an effect was only detected when the level of both proteins were simultaneously reduced. This suggests that miRNA mediated silencing utilizes either the

Dcp2 or Nudt16 decapping proteins.

To determine whether the increased luciferase activity in the Dcp2β/β/sh-Nudt16

MEF cell line was due to an increase in luciferase mRNA levels or an indirect

consequence of reduced Dcp2 and Nudt16, Renilla luciferase mRNA levels were

determined relative to the mRNA from the cotransfected control Firefly luciferase

reporter. As shown in Figure 18C, despite a considerable decrease in luciferase activity

from the let-7 bulge sites containing mRNA, only a modest decrease of the mRNA was

observed. Furthermore, a statistically significant difference in steady state Renilla

luciferase mRNA containing the let-7 bulge sites was not detected in the Dcp2 and

Nudt16 double knockdown MEF cells relative to control cells. These data indicated that

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the detected decrease in luciferase activity in Figure 18A is not a direct consequence of a

corresponding decrease in mRNA levels. Furthermore, the observed increase in

luciferase activity from the let-7 bulge site-containing mRNA in the double knockdown

cells are not a result of increased steady state mRNA levels and therefore appear to be an

indirect consequence of Dcp2 and Nudt16 decapping on translation.

Differential utilization of Dcp2 and Nudt16 in the decay of ARE-containing mRNAs

Dcp2 has been reported to function in ARE-mediated decay where the

destabilizing ARE-binding factors, TTP and BRF-1, can recruit the Dcp2 decapping

complex in vitro to promote ARE-dependent decapping (Fenger-Gron et al, 2005a;

Lykke-Andersen & Wagner, 2005). To examine the consequence of the lack of detectable

Dcp2 on the decay of ARE-containing mRNAs, we investigated the stability of endogenous ARE-containing mRNAs in the Dcp2β/β cells. Wild type and Dcp2β/β cells

were treated with actinomycin D and the decay of specific ARE-containing mRNAs was

analyzed by quantitative real-time PCR for up to two hours following transcriptional

arrest. Six different ARE-containing mRNAs with detectable expression in MEF cells

were tested. As shown in Figure 19, stability of the IFNα2 mRNA, was significantly

increased in Dcp2β/β cells, with a two fold increase in half-life upon reduction of Dcp2.

No significant difference in mRNA half-lives was detected in the stability of c-Fos, IL4, c-Jun and P53 mRNAs when Dcp2 was individually reduced. However, a contribution of

Dcp2 was detected on the stability of c-Myc and c-Jun mRNAs when its levels were reduced in conjunction with Nudt16 (see below). These data indicate that Dcp2 protein is

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Figure 19. Differential utilization of Dcp2 and Nudt16 in ARE-mediated decay. mRNA levels of the indicated mRNAs were followed up to 2 hours post actinomycin D treatment. Levels of the denoted endogenous ARE-containing mRNAs remaining at the indicated time points were determined by quantitative reverse transcription and real-time

PCR. The values of three independent RNA preparations normalized to the β-actin mRNA are shown, with +/- standard deviation. Cell line nomenclature is as described in the legend to Figure 15.

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differentially utilized in the decay of ARE-containing mRNAs where a subset, but not all

ARE-containing mRNAs appear to be responsive to Dcp2 activity.

The same mRNAs were also analyzed in the absence of Nudt16. Interestingly, a reduction of Nudt16 protein levels did not alter the half-lives of the c-Fos, IL4, or P53 mRNAs (Figure 19). However, the c-Myc mRNA was stabilized by 50% from a 30 min to 45 min half-life upon reduction of Nudt16 and importantly, it was further stabilized to a 1 hr half-life in the double knock down conditions, indicating an additive role for

Nudt16 and Dcp2 in decay of the c-Myc mRNA. Redundant roles of Dcp2 and Nudt16 were also observed with the c-Jun mRNA where an individual reduction of either Dcp2 or Nudt16 protein levels had no effect on mRNA half-life while a simultaneous reduction of both proteins lead to an almost doubling of the c-Jun mRNA half-life from 30 min to

50 minutes. These results indicate that Dcp2 and Nudt16 fulfill redundant roles in regulating c-Jun mRNA decay. Collectively, our analysis of ARE-containing mRNAs reveals a complex and transcript specific utilization of the decapping enzymes by ARE- containing mRNAs ranging from no consequence of reduced Dcp2 or Nudt16 levels to synergistic roles of both decapping enzymes.

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Discussions

In this study, we identified a second cytoplasmic decapping enzyme Nudt16 and

investigated the utilization of Dcp2 and Nudt16 decapping enzymes in distinct cellular

mRNA decay events in mouse embryonic fibroblast cell lines. Our results suggest Dcp2

and Nudt16 are selectively involved in specific mRNA decay processes and do not

uniformly function on all mRNAs. The NMD substrate tested preferably used Dcp2;

while an miRNA mediated silencing reporter appear to use both decapping enzymes

redundantly and ARE-containing mRNAs differentially utilized the two decapping

enzymes.

The Xenopus laevis ortholog of Nudt16, X29, was reported to be nucleolar and

specifically bind the U8 snoRNA and preferentially hydrolyzed it at least in vitro (Ghosh

et al, 2004; Tomasevic & Peculis, 1999). However, we detected epitope-tagged Nudt16 predominantly in the cytoplasm with no obvious accumulation in nucleoli. Although the antibody we have generated can detect Nudt16 by Western analysis, we have not been able to detect endogenous Nudt16 by immunofluorescence above background in any cell

type we have tested with either polyclonal sera or affinity purified antibody. Although

the cellular localization in Figure 14B is based on exogenous Nudt16 expression, we

believe the cytoplasmic signal is representative of the endogenous protein. As shown in

Figure 14B, the signal is predominantly cytoplasmic and generally devoid from the

nucleus. It is unlikely that the Flag-tag is interfering with an adjacent nuclear localization

signal on the expressed protein since similar results were obtained regardless of whether

the tag was at the amino terminus or carboxyl terminus.

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NMD has been shown to evoke decapping in both yeast and mammalian cells

(Couttet & Grange, 2004; Lejeune et al, 2003; Muhlrad & Parker, 1994). The key NMD

factor Upf1 can form a complex with the Dcp2 protein in both organisms (He & Jacobson,

1995; Lejeune et al, 2003; Lykke-Andersen, 2002). Knockdown of Dcp2 by siRNA impaired the removal of nonsense codon containing transcript in Hela cells, indicating that Dcp2 decapping is utilized in NMD (Lejeune et al, 2003). Our data with MEF cells containing virtually undetectable levels of Dcp2 was consistent with previous reports where clearing of nonsense containing β–globin or TCRβ reporter mRNA was compromised in the absence of Dcp2 (Figure 17). Interestingly, we did not detect a role for Nudt16 on the same mRNA (Figure 17), suggesting Dcp2 but not Nudt16 is specifically utilized with these NMD substrates. As expected, a knockdown of Upf1 resulted in a more pronounced increase in the PTC containing mRNA levels, however, this is likely an underestimation since only a 65% reduction in Upf1 mRNA levels were obtained (data not shown). Considering PTC containing transcripts are subjected to

NMD-directed clearing by multiple pathways, including deadenylation (Chen & Shyu,

2003; Zheng et al, 2008) and Smg6 endonucleolytic decay (Eberle et al, 2009; Gatfield &

Izaurralde, 2004; Huntzinger et al, 2008), only a modest increase of the PTC transcript would be expected in the Dcp2β/β cells. Thus Dcp2 decapping contributes to, but is not essential for NMD, while Nudt16 decapping may not be a major contributor to NMD.

Decapping has been implicated in miRNA mediated silencing, since depletion of

Dcp1:Dcp2 in Drosophila cells partially relieved miRNA mediated repression of

luciferase activity (Behm-Ansmant et al, 2006; Rehwinkel et al, 2005). However in our analyses, depletion of Dcp2 protein alone in MEF cells did not significantly affect

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miRNA mediated silencing of luciferase activity (Figure 18A). Only when Dcp2 and

Nudt16 were simultaneously reduced did we observe a significant restoration of

luciferase activity (Figure 18A). This indicates Dcp2 and Nudt16 are redundant in miRNA-mediated repression, and the miRNA silencing machinery could use either Dcp2 or Nudt16 in mammals. The lack of an obvious ortholog for Nudt16 in Drosophila

(Taylor & Peculis, 2008) might be one explanation. One surprising finding in our studies was that although we detect a simultaneous dependence of both decapping enzymes, miRNA target reporter mRNA levels were not significantly altered under reduced decapping enzyme conditions. This could either be due to residual levels of decapping enzyme still present in the knockdown conditions or an indication of an indirect role of the decapping enzymes in mammalian miRNA directed silencing in this system. The latter possibility suggests a potential role of Dcp2 and Nudt16 in the regulation of mRNA(s) that encode facilitators of translational silencing. Future efforts will address this possibility. Furthermore, since decapping activators including Dcp1a, Edc4, Edc3,

Lsm1-7 are differentially required for distinct miRNA targets in Drosophila (Eulalio et al,

2007b), analysis of additional miRNA substrates will be necessary to determine whether

Dcp2 and or Nudt16 may also fulfill a direct function in regulating mRNA levels for a subset of miRNA targets in mammal cells. In contrast to the miRNA-directed silencing, siRNA-directed silencing was not impacted by the loss of either Dcp2 or Nudt16 (Figure

18A and 18C) consistent with siRNA predominantly evoking an endonucleolytic cleavage mechanism for mRNA decay (Liu et al, 2004a; Zamore et al, 2000).

An important finding revealed from our data is that ARE-containing transcripts

can differentially utilize the two different decapping enzymes to promote their decay.

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Several lines of evidence have previously implicated decapping in ARE-mediated decay

with the focus exclusively on Dcp2. For example, the ARE-binding factors TTP and

BRF-1 can interact with the decapping complex and TTP could specifically activate Dcp2

decapping in vitro (Fenger-Gron et al, 2005a; Lykke-Andersen & Wagner, 2005),

suggesting ARE-mediated decay involves Dcp2 decapping. In addition Dcp2 was shown

to contribute to decay of the c-Fos mRNA (Murray & Schoenberg, 2007). However, the

first line of evidence suggesting that Dcp2 may not be the only decapping enzyme

involved in ARE-mediated decay was provided by Stoecklin et al. (Stoecklin et al, 2006)

where intriguingly, depletion of Xrn1 significantly stabilized a GM-CSF reporter mRNA

while depletion of Dcp2 had no detectable effect on the decay of the same mRNA. These

finding suggested the GM-CSF mRNA is degraded from the 5' end but in a Dcp2

independent manner.

Our results demonstrate that decapping enzymes could be differentially utilized

for the decay of ARE-containing mRNAs where both Dcp2 and Nudt16 can individually

or synergistically be involved (Figure 19). The IFNα2 mRNA selectively utilizes the

Dcp2 protein for decay while degradation of the c-Myc mRNA involves both Dcp2 and

Nudt16. The c-Jun mRNA was only stabilized when both Dcp2 and Nudt16 were

reduced, suggesting Dcp2 and Nudt16 are redundant in the decay of this mRNA. The

decay of c-Fos, IL4 and P53 mRNAs were insignificantly affected by a depletion of

either Dcp2 or Nudt16. This is not surprising since the ARE also promotes exosome

mediated 3' to 5' decay (Chen et al, 2001; Mukherjee et al, 2002). The differential

requirement of decapping enzymes in ARE-mediated decay is likely a consequence of distinct ARE-binding proteins that interact with each ARE and recruit one or the other

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decapping enzyme. Future efforts to identify Nudt16 binding partners will begin addressing this possibility.

The demonstration that at least two different decapping enzymes can initiate mRNA 5' end decay in mammalian cells and that distinct mRNA decay processes differentially utilize these enzymes, provides new avenues to pursue mRNA turnover.

The utilization of distinct decapping enzymes in specific decay may be a broadly utilized process as evident by a 5´-end quality control mechanism with decapping endonuclease activity on aberrantly capped mRNAs (Jiao et al, 2010). Future efforts addressing how each decay process and each transcript recruits and modulates the decapping proteins will greatly advance our understanding of regulated mRNA decay.

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Chapter III: Regulation of Decapping in Interferon-mediated

Anti-viral Immunity

Summary

Viral-induced transcriptional activation of cytokines, such as type-I interferons

(interferon (IFN)-α and IFN-β), constitutes the first line of antiviral immune response.

The family of interferon regulatory factors (IRFs) play an important role in modulating expression of type I IFNs. Here we show that regulation of decapping is important for type-I IFN mediated anti-viral immune response. In mouse embryonic fibroblasts (MEFs) lacking the decapping enzyme Dcp2, the anti-viral immune response is significantly elevated, evident by increased type-I IFN production and upregulation of other antiviral genes. We identified IRF7 mRNA as the direct target of Dcp2. In Dcp2 knockdown MEF cells, IRF7 mRNA was significantly stabilized, leading to increased IRF7 protein levels.

Our data indicates Dcp2 negatively modulates the anti-viral immune response by promoting the degradation of IRF7 mRNA. Interestingly, Dcp2 expression is also induced in virus infection, suggesting a negative feedback loop in the anti-viral immune response to temper IFN production and promote the return to normal homeostasis.

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Introduction

The interferon (IFN) system is the first line of defense against virus infection in

mammals. Type-I interferons (IFN-α and IFN-β) are widely expressed cytokines that

mediate the anti-viral innate immune response. They are induced in response to viral

exposure and are extracellularly secreted and function on yet uninfected cells to activate a

global antiviral state in which virus replication is inhibited. Type I IFNs can be induced in most cell types and exhibit profound pleiotropic effects on many aspects of cellular

functions, including gene transcription, protein translation, cell growth and cell motility.

At the molecular level, the transcriptional activation of IFN-α/β genes are

mediated by a complex array of pathways involving multiple transcription factors. In

most cases, the ligands of type I IFN-inducing receptors are viral nucleic acids such as

single-stranded (ss)RNA, doublestranded (ds)RNA or dsDNA (Fensterl & Sen, 2009). A

well-characterized pathway is triggered by viral double-stranded RNA generally derived

from viral RNA genomes or viral replication intermediates. It can be sensed in the

cytoplasm by RIG-I (retinoic acid-induced gene-I) and MDA-5 (melanoma

differentiation-associated gene-5) (Andrejeva et al, 2004; Fensterl & Sen, 2009;

Yoneyama et al, 2004), or in the endosome by TLR3 (Alexopoulou et al, 2001). The signaling cascades initiates, either via a TRAF3-containing complex or TRAF6- containing complex, phosphorylates and activates critical transcription factors in the induction of type I IFNs, including IRF-3, IRF-7, NF-κB, ATF-2, and c-JUN (Fitzgerald et al, 2003; Saha et al, 2006; Seth et al, 2005; Sharma et al, 2003; Xu et al, 2005). These factors then bind to the promoter regions of type I IFNs as well as other

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cytokine/chemokine genes, and activate their transcription. IRF3 and IRF7 are members

of a family of transcription factors called IFN regulatory factors (IRF), and they play

crucial roles in mediating the induction of type I IFNs (Nguyen et al, 1997). IRF-3 is

constitutively expressed in all cell types while IRF7 is usually induced by virus infection in many cell types. Most of IFN-α subtypes require the induced IRF-7, whereas IFN-β

can be induced without IRF-7 by the constitutively expressed IRF-3 (Honda et al, 2006).

Although transcriptional regulation of IFN gene expression has been well characterized in the activation of the anti-viral innate immune response, little is known about the significance of regulation of mRNA decay in this process. mRNA decay plays an important role in the control of gene expression and response to regulatory events. In eukaryotic cells, bulk mRNA decay typically initiates with the removal of 3′ poly(A) tail followed by degradation of the mRNA in a 5′ to 3′ direction or a 3′ to 5′ direction (Wilusz et al, 2001b). Degradation from 3′ end is carried out by the cytoplasmic RNA exosome, which is a multisubunit 3′ to 5′ exoribonuclease complex (Liu et al, 2006), and the resulting cap structure is hydrolyzed by the scavenger decapping enzyme DcpS (Liu &

Kiledjian, 2006). In the 5′ to 3′ decay pathway, the monomethyl guanosine (m7G) mRNA cap is cleaved first by the Dcp2 decapping enzyme (Dunckley & Parker, 1999;

Lykke-Andersen, 2002; van Dijk et al, 2002; Wang et al, 2002b) and the monophosphate

RNA is degraded progressively by the 5′ to 3′ exoribonuclease Xrn1 (Decker & Parker,

1993; Hsu & Stevens, 1993).

Decapping is a key step in mRNA decay that’s highly regulated by both positive and negative regulators. To study the significance of decapping in anti-viral immune response, we used mouse embryonic fibroblast cells deficient in Dcp2 decapping enzyme.

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By microarray analysis, we found that after lentivirus infection, a group of anti-viral

genes were significantly upregulated in Dcp2 knockdown MEF cells compared to

wildtype cells. Further studies revealed that Dcp2 directly regulates the mRNA stability

of IRF7, a key transcription factor in the anti-viral immune response. IRF7 protein levels

increased in Dcp2 knockdown MEF cells, which could subsequently lead to elevated expression of various downstream anti-viral effector genes. In addition, Dcp2 expression is also induced upon viral infection providing the potential for a negative feedback regulatory network in the anti-viral immune response.

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Results

Genes involved in anti-viral immune response were upregulated in Dcp2β/β MEF

cells

In an effort to identify genes potentially regulated by Dcp2 and/or Nudt16 regulated mRNAs in cells, the identity of mRNAs with altered expression levels in Dcp2 and/or Nudt16 knockdown MEF cells was determined by microarray analysis. Total

RNAs from the four MEF cell lines with Dcp2 and/or Nudt16 knockdown as described in

Chapter II (Figure 14) were subjected to Illumina Sentrix Mouse 24K Array in duplicate.

Compared to the wildtype MEF, 194 mRNAs were upregulated in Dcp2β/β cells more than 1.5 fold, 297 upregulated in Nudt16 knockdown cells and 279 in the double knockdown cells. In depth analysis of these upregulated mRNAs by Ingenuity Pathway

Analysis (IPA) software revealed that genes involved in anti-viral response pathway were

significantly overrepresented in Dcp2β/β cells or the double knockdown cells, but not

Nudt16 single knockdown cells, indicating they are specifically regulated by Dcp2. As

shown in Figure 20A, among the 30 genes involved in anti-viral immune response

pathway with detectable levels in the microarrays, 14 were significantly upregulated with

more than 1.5 fold in Dcp2β/β cells. Three representative mRNAs, IRF7, CXCL10 and

OAS2 were chosen for further confirmation by real-time PCR. IRF7 is a key

transcription factor that induces expression of type I interferons and other cytokines and

chemokines upon virus infection; CXCL10 is a chemokine whose transcription is

activated by IRF7 (as well as IRF3); OAS2 is an interferon-inducible gene which encodes

2'-5'-oligoadenylate synthetase, a critical effector in the innate immune response to viral

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Figure 20. A subset of genes involved in anti-viral immune response were upregulated in Dcp2β/β MEF cells.

(A) Wildtype and Dcp2β/β MEF cells were infected with lentivirus for 2 days. Total cellular RNAs were extracted and subjected to microarray analysis. 30 genes involved in anti-viral immune response with signals above background were selected, and the fold

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change of each gene in Dcp2β/β MEF compared to wildtype MEF is plotted. Data represent the average of two independent experiments and dark bars indicate genes that were upregulated more than 1.5 fold in Dcp2β/β MEFs.

(B) RNAs used in (A) were reverse transcribed and subjected to real-time PCR analysis.

IRF7, CXCL10 and OAS2 mRNAs were amplified with gene specific primers and normalized to β-actin mRNA level. The mRNA levels in the wild type MEF were arbitratily set to 1. The average of two experiments is shown.

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infection. Consistent with the microarray, the expression of all three genes were

upregulated from 2 to 5 fold in the Dcp2β/β cells relative to wildtype cells by real-time

RT-PCR analysis (Figure 20B).

The microarray assays above were carried out after infection of lentiviruses that

express Nudt16-specific shRNA or a control lacking an shRNA. To examine whether the

increased expression of anti-viral genes in Dcp2β/β MEF cells was dependent on virus

infection, we performed a time course of virus infection. Wildtype and Dcp2β/β MEF cells were infected with control lentiviral particles lacking an encoded shRNA. Total RNAs were isolated from cells at 0 hour (before infection),12 and 24 hours post- infection and the levels of specific mRNAs present at each time point assessed by quantitative RT-PCR.

Importantly, steady state level of IRF7 mRNA was 2.2 fold higher in the Dcp2β/β MEF

cells relative to wild type cells prior to viral infection (Figure 21A). Similarly, OAS2

mRNA levels were also elevated (2.5 fold) in the Dcp2β/β MEF cells at time zero while

no differences were observed for CXCL10 at time zero. IRF3, which is also a critical

transcription factor in anti-viral immune response but not upregulated in Dcp2β/β cells in

the microarray, showed no significant difference between wiletype and Dcp2β/β MEF

cells. These data indicate that Dcp2 influences the expression of IRF7 and OAS2 under

normal untransduced conditions.

An impact of Dcp2 on genes involved in innate immunity is further evident

following viral transduction. As shown in Figure 21B, IRF7 mRNA level increased about

5 fold in Dcp2β/β MEF cells compared to the wildtype cells at 12 and 24 hours post-viral

infection. Similar result was observed for OAS2 mRNA, which had a 6 fold increase in

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Figure 21. IFN mediated anti-viral immune response was elevated in Dcp2β/β MEF cells following lentivirus infection.

(A) Total RNAs of wild type and Dcp2β/β MEF cells were extracted and mRNA levels of specific genes were determined by quantitative real-time RT-PCR, normalized to β-actin

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mRNA. mRNA levels in wild type cells were arbitrarily set to 1. Data represent the

average of three independent experiments with standard deviation denoted by the error

bar.

(B) Wildtype and Dcp2β/β MEF cells were treated with control lentivirus and total RNAs

were extracted at 12 hours and 24 hours after infection. mRNA levels of specific genes

were determined by quantitative real-time RT-PCR, and normalized to β-actin mRNA. mRNA levels in wild type cells at 12 hours post-infection were arbitrarily set to 1. Data represent the average of three independent experiments with standard deviation denoted by the error bar.

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Dcp2β/β cells. A modest increase (1.8 fold) of CXCL10 mRNA level in Dcp2β/β MEF

cells was evident 24 hours after virus treatment. There was no significant difference of

IRF3 mRNA level between wildtype and Dcp2β/β MEF cells after viral infection. Because

type I interferons are key factors in anti-viral immunity, we also tested the expression of

interferon-α and interferon-β in wildtype and Dcp2β/β cells upon viral infection. IFNα2

and IFNβ mRNAs were not detectable before infection (data not shown) and were both

induced upon virus treatment. In Dcp2β/β MEF cells, a 3 fold greater induction was

observed for IFNα2 mRNA compared to wild type cells while a 2 fold more induction was evident for IFNβ. These data indicate that innate immune response to viral infection is generally enhanced in Dcp2β/β MEF cells, with upregulation of a number of , but not all,

key factors in the interferon mediated anti-viral response pathway.

Catalytic activity of Dcp2 is critical for the regulation of anti-viral immune response

To determine whether the Dcp2-directed regulation of the anti-viral immune

response is dependent on the catalytic activity of Dcp2, we introduced wildtype or

catalytically inactive Dcp2, which contains mutations of two conserved glutamic acids at

residues 147 and 148 within the NUDIX motif, into the Dcp2β/β MEF cell. Stable cell

lines that constitutively express exogenous Dcp2 protein were selected and the level of

Dcp2 protein was analyzed by western blot assay. As shown in Figure 22A, the wildtype

and catalytically inactive Dcp2 genes are expressed at comparable levels in the

Dcp2β/β MEF cell. These cells were subjected to viral infection and the induction of IRF7

and IFNα2 mRNAs determined. As shown in Figure 22B, Dcp2β/β MEF cells expressing

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Figure 22. Catalytic activity of Dcp2 is required for the regulation of anti-viral

immune response.

(A) Wildtype or the catalytic mutant Dcp2 (hDcp2 EE/Q) or the control vector (con) were

expressed in indicated MEF cells and stable cell lines were selected. Expression of Dcp2 protein is confirmed by western blot with GAPDH as the loading control. Dcp2β/β MEFs

expressing control vector is used as a negative control and wildtype MEFs expressing

control vector is used as the positive control.

(B) MEF cells in (A) were infected with control lentivirues. Total RNAs before infection

and 12 hours after infection were extracted, and mRNA levels of specific genes were

determined by quantitative real-time RT-PCR and normalized to β-actin mRNA. For

IRF7, mRNA level in wildtype cells before infection were arbitrarily set to 1; for IFNα2,

mRNA levels in wildtype cells after infection were arbitrarily set to 1. Data represent the

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average of three independent experiments with standard deviation denoted by the error bar. Overexpression of wildtype but not the catalytic mutant Dcp2 reversed the increase of both IRF7 and IFNα2 in Dcp2β/β MEF cells.

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control vector lacking Dcp2 consistently demonstrated elevated levels of IRF7 and

IFNα2 upon viral infection compared to wildtype MEF cells. Importantly, the expression

of wildtype Dcp2 in Dcp2β/β background greatly suppressed the expression of IRF7 and

IFN α2 to levels comparable to the wildtype MEF cells. However, the expression of

catalytically inactive Dcp2 did not have significant effect on the expression of IRF7 and

IFN α2 in the Dcp2β/β MEF cells. These results indicate that the catalytic decapping activity of Dcp2 is critical for the regulation of anti-viral immune response in MEF cells.

Dcp2 regulates the mRNA stability and protein expression of IRF7

Since the decapping activity of Dcp2 is required for the regulation of anti-viral immune response, it is reasonable to anticipate that Dcp2 regulates the mRNA stability of a subset of key genes in the anti-viral pathway. In the previous study (Chapter II, Figure

19), IFNα2 was already shown to be a Dcp2 target whose mRNA stability is regulated by

Dcp2. Here we tested several additional genes in the pathway. Wildtype and

Dcp2β/β MEF cells were infected with lentiviruse for 24 hours and then treated with

actinomycin D to block transcription and the decay of specific mRNAs was monitored by

quantitative real-time RT-PCR following a time course up to 8 hours. As shown in Figure

23A, the IRF7 mRNA was significantly stabilized under reduced Dcp2 levels. The half- life of the IRF7 mRNA was 4 hours in the wildtype MEF cells and greater than the 8

hours time course of the experiment in the Dcp2β/β MEF. A more modest difference was

detected for CXCL10 mRNA where the half-life was increased from approximately 2

hours in the wild type MEF to 3 hours in the Dcp2β/β MEF cells. No significant difference

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Figure 23. Dcp2 regulates mRNA stability and protein expression of IRF7.

(A) Wildtype and Dcp2β/β ΜΕF cells were infected with control lentivirus for 24 hours and then treated with actinomycin D. Total RNAs were harvested at indicated time post- treatment and the decay of IRF7, CXCL10 and OAS2 mRNAs were determined by quantitative real-time RT-PCR and normalized to β-actin mRNA. The average of three independent experiments was plotted with standard deviation denoted by the error bar.

IRF7 mRNA was significantly stabilized in Dcp2β/β ΜΕF cells.

(B) Wildtype and Nudt16 knockdown ΜΕF cells were infected with control lentivirus for

24 hours and then treated with actinomycin D. Total RNAs were harvested at indicated time post-treatment and the decay of IRF7 mRNA was determined by quantitative real- time RT-PCR and normalized to β-actin mRNA. The average of three independent experiments was plotted with standard deviation denoted by the error bar.

Total cell extracts before and after infection were harvested and IRF7 protein level is determined by western blot with GAPDH as the internal control. Quantitation of IRF7 protein normalized to GAPDH is shown on the right. The average of three independent experiments was plotted with standard deviation denoted by the error bar. IRF7 protein level increased in Dcp2β/β ΜΕF cells with or without virus infection.

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in the stability of the OAS2 mRNA was detected. As a control, the IRF7 mRNA stability

did not change in MEF cells with reduced Nudt16 levels (Figure 23B). Therefore, these

data demonstrate that the decay of IRF7 mRNA is regulated by Dcp2. Since IRF7 is an

upstream transcription factor in the anti-viral immune response pathway, it could be the

major target of Dcp2 in viral infection that in turn affects expression of a broad range of

downstream genes.

To examine whether the regulation of IRF7 mRNA stability by Dcp2 manifests to

the level of altered IRF7 protein, we analyzed IRF7 protein levels by western blot in

wildtype and Dcp2β/β MEFs with or without virus infection. Consistent with the elevated

mRNA levels, the loss of Dcp2 in the Dcp2β/β MEF cells result in a 3 fold increase in

IRF7 protein levels compared to wildtype MEF prior to virus infection (Figure 23C). As

expected, IRF7 protein levels increased in both cell backgrounds following viral infection,

however a 2.5 fold greater increase was detected in the Dcp2β/β MEF cells. Collectively,

these data demonstrate Dcp2 is a negative regulator of IRF7 mRNA stability and protein

accumulation and may function to attenuate it expression upon viral infection.

Dcp2 expression is induced following viral infection

As a potential modulator of anti-viral immune response gene mRNAs, we next

determined whether the expression of Dcp2 itself was altered during viral infection. Wild

type MEF cells were infected with lentiviral particles and Dcp2 expression followed over time by real-time RT-PCR and western-blot analysis. As in Figure 24A, Dcp2 mRNA levels increased upon treatment with viruses, with 2.5 and 4 fold inductions compared to

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Figure 24. Dcp2 expression is induced in virus infection.

(A) Wildtype MEF cells were treated with lentivirus or ploy(IC) (50 µg/ml) for indicated time, or growth media containing no virus for 24 hours. Total RNAs before and after treatment were harvested and Dcp2 mRNA level was determined by real-time RT-PCR and normalized to β-actin mRNA. Dcp2 mRNA level in untreated MEF was arbitrarily set to 1. The average of three independent experiments was plotted with standard deviation denoted by the error bar. Dcp2 mRNA significantly increased with virus infection or poly (IC) treatment.

(B) Wildtype MEF cells were treated with lentivirus, poly (IC) (50 µg/ml) or growth media containing no virus for 24 hours. Total cell extracts before and after treatment were harvested and Dcp2 protein levels were determined by western blot analysis with

GAPDH as internal control. Dcp2 protein was induced in virus infection or poly(IC) treatment.

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the untreated cell at 12 and 24 hours post-infection respectively. Similar induction was also observed when cells were treated with the viral dsRNA mimic, poly (I:C). Dcp2 levels were unchanged in control cells treated with growth media lacking virus, suggesting Dcp2 expression is induced by virus infection. Consistent with the mRNA data, Dcp2 protein levels also significantly increased upon viral infection, or poly (I:C) treatment (Figure 24B). These data imply that when anti-viral immune response is elicited, Dcp2 expression is also induced, consistent with a negative feedback, which may function as a failsafe mechanism to prevent over response of the immune system.

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Discussion

In this section, we report a novel role for the Dcp2 decapping enzyme in regulating anti-viral immunity. Microarray assays revealed that a group of genes involved in anti-viral immune response were upregulated in Dcp2β/β MEF cells upon virus infection. Further mechanistic analysis identified IRF7 as the direct target of Dcp2. With reduced Dcp2 level, the mRNA stability of IRF7 was greatly enhanced. As a result IRF7 protein level also increased in the Dcp2β/β MEF cells compared to wildtype MEF cells with or without virus infection. IRF7 is a key transcription factor in the anti-viral immune response pathway which regulates the expression of IFNs and many other cytokines and chemokines required for the body to establish an anti-viral state. Regulation of IRF7 at the level of mRNA stability defines a previously unknown mechanism of regulating the immune response upon viral infection and potentially provides a new therapeutic target to improve immune response in humans.

In the microarray, we identified 194 mRNAs upregulated in Dcp2β/β MEF cells compared to the wildtype MEFs. However, when we compare the upregulated mRNAs with Dcp2-bound mRNAs (microarray in Chapter I, Table IV), there is very little correlation. One explanation for this is the difference in species. The microarray for

Dcp2-bound mRNAs was carried out with human 293T cells, and the microarray in this chapter was performed with mouse embryonic fibroblast-derived mRNA. Although many mRNAs were highly conserved between human and mouse, the conserved region usually concentrate in the open reading frame of mRNAs (IS THERE A REFERENCE FOR

THIS?), while the 5' UTR regions are more variable. Considering the 5' sequences are

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critical for Dcp2 binding and decapping, it is reasonable that Dcp2 may regulate different

mRNAs in human and mouse cells. In addition, the association of mRNAs with Dcp2

suggests a role of Dcp2 in the decay of specific mRNA substrates, but this does not

necessarily lead to a change in the steady state level of these RNAs. First, Dcp2-directed decapping is only one aspect of the decay process since there are additional decapping enzymes like Nudt16 that may function, as well as a role for the 3'-end decay pathway to also compensate for impaired decapping. Second, the steady state level of an mRNA is determined by the rate of its transcription and decay, and the rate of transcription may be altered in a compensatory mechanism that may overcome alterations in decay.

The Illumina Sentrix Mouse 24K Array contained 72 known genes involved in anti-viral immunity, of which 30 were detected with a level above background in our experiment. mRNA levels for almost half of these genes (14) were elevated in

Dcp2β/β MEF cells by greater than 1.5 fold. Interestingly, most of the upregulated genes

are downstream effectors that function in directly eliminating viral RNA or inhibiting

viral replication. Only one of the 14, IRF7, is involved in upstream viral-sensing and signaling. Our result that Dcp2 regulates the mRNA stability of IRF7 suggests a model where Dcp2 knockdown causes an increase in IRF-7 expression, which results in elevated type-I IFN production in virus infection, and subsequently evokes an increase of IFN-

inducible anti-viral effector genes. Consistent with this premise, we showed that IRF-7

protein levels were upregulated in uninfected Dcp2β/β MEF cells (Figure 23C), and the

expression of IFN-α and IFN-β were enhanced (Figure 21B) in Dcp2β/β MEF cells upon

exposure to virus. This is dependent on Dcp2 decapping activity since exogenous expression of wildtype but not catalytically inactive Dcp2 mutant abolished the enhanced

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expression of IRF7 and IFN-α in Dcp2β/β MEF cells (Figure 22). Therefore, Dcp2 could

mediate a broad anti-viral immune response by targeting a critical upstream transcription

factor.

Expression of type I IFNs leads to induction of anti-viral effector genes and

establishes an anti-viral state on host cell. This process needs to be precisely controlled.

Positive regulators of type I IFN response have been extensively studied. For instance,

mice lacking two virus sensors RIG-I and MDA5 exhibit increased sensitivity to infection

of various RNA viruses (Kato et al, 2006). Knockout of transcription factor STAT1

renders mice susceptible to virus infection due to impaired IFN response (Durbin et al,

1996). Disruption of Pkr gene, the dsRNA-activated protein kinase, caused reduced anti-

viral response to EMCV and VSV infection in mice (Balachandran et al, 2000). Negative

regulators of type I IFN response have also been described. Knockout mice of the

translational repressor 4E-BP are highly resistant to infection by a variety of viruses due

to increased type-I IFN production (Colina et al, 2008). Interestingly, the enhanced type-I

IFN response in 4E-BP knockout mouse is caused by upregulation of IRF7 messenger

RNA translation, indicating translational control of IRF7 expression by 4E-BP protein

(Colina et al, 2008). In this study, we demonstrated negative regulation of IRF7 by the

Dcp2 decapping enzyme at the mRNA stability level. The half-life of IRF7 mRNA

increased more than two fold in Dcp2β/β MEF cells (Figure 23A) compared to that of

wildtype MEF cells, indicating IRF7 mRNA degradation proceeds through Dcp2- mediated decapping. IRF7 is believed to be an important regulator of anti-viral immunity, since a deficiency of IRF7 in mice resulted in increased susceptibility to viral infection and lethality in addition to blunted systemic type I IFN response (Daffis et al, 2008).

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Modulation of IRF7 levels by mRNA decay provides a new avenue of post- transcriptional regulation of anti-viral innate immunity.

It is interesting that the decay of IRF7 mRNA specifically utilizes Dcp2 decapping enzyme but not Nudt16, since IRF7 mRNA did not accumulate in Nudt16 knockdown MEF cells and its stability was not changed either (Figure 23B). In chapter II, we showed Dcp2 and Nudt16 could be differentially employed in different mRNA decay pathways. Here we demonstrate another example of transcript specific utilization of decapping enzymes. In chapter I, we identified a stem-loop structure (DBDE) which could activate Dcp2 decapping when placed at 5' terminus of an mRNA. Examination of the 5' UTR of mouse IRF7 mRNA revealed that it could potentially form a stable stem- loop structure that starts within 10nt from the 5' end. Although the length of the predicted stem is longer than the prototype DBDE in Rrp41 mRNA (12 base pairs), it still fits the criteria of a functional DBDE, which should start within 10nt from the 5' terminus of an mRNA and contain more than 8 base pairs in the stem region. Human IRF7 mRNA is also predicted to form a stem-loop structure at the 5' end and the length of the stem region is more comparable to that of the prototype DBDE in Rrp41 mRNA. As the

DBDE element could directly recruit Dcp2 onto an RNA and activate decapping, the

IRF7 mRNA could be regulated by Dcp2 through a DBDE mediated mechanism.

The importance of regulated mRNA stability has long been established with the control of cytokine and chemokine mRNAs during an immune response. These mRNAs usually contain AU-rich elements (AREs) in their 3'UTRs which confer rapid degradation by a mechanism called ARE medicated decay (AMD). Various ARE-binding proteins are involved in regulating the stability of these mRNAs. For instance, Tristetraprolin (TTP)

144

binds to AREs in various mRNAs (such as TNFα and GM-CSF) and can promote

deadenylation and subsequent degradation of these transcripts (Carballo et al, 1998;

Carballo et al, 2000); while another well characterized ARE-binding protein, HuR (Hu

antigen R) is implicated in the stabilization of many ARE containing mRNAs, including

TNFα, VEGF, GM-CSF and c-fos (Dean et al, 2001; Fan & Steitz, 1998; Levy et al,

1998; Peng et al, 1998). AREs could differentially activate both 5′ to 3′ and 3′ to 5′ decay

pathways, including Dcp2 decapping (Chen et al, 2001; Gao et al, 2001; Lykke-Andersen

& Wagner, 2005; Mukherjee et al, 2002; Murray & Schoenberg, 2007; Stoecklin et al,

2006). In our study, we identified two mRNAs in the anti-viral immune response, IRF7

and IFNα2, whose stability is regulated by Dcp2. IFNα2 is an ARE-containing mRNA, while the IRF7 mRNA has a short 3'UTR that does not contain any obvious ARE element.

The half-life of IRF7 mRNA (4 hours) is considerably longer than that of a typical ARE-

mRNA (about 0.5 hour for c-myc and c-fos mRNAs). Therefore, IRF7 represents an

example of non-ARE mRNA which undergoes regulation of mRNA stability in the

immune system.

Induction of mRNA decay factors in the immune response has already been

demonstrated. For instance, TTP levels increase in lipopolysaccharide (LPS)-stimulated, bone marrow–derived macrophages, leading to instability of TNF-α mRNA and inhibition of TNF-α secretion (Cao et al, 2004; Carballo et al, 1998). Therefore, TTP participates in a negative feedback loop, in which the level of TTP is increased by the same stimuli that induces expression of TNF-α in macrophages. A similar scenario may also be functional with Dcp2: viral infection induces expression of type I IFNs and IRF7,

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and also increases the level of Dcp2. Dcp2 promotes decay of IRF7 and IFNα2, forming

a negative feedback loop in the anti-viral immune response. This might help to prevent

excess production of IFNs, which could manifestate autoimmune pathologies, since

therapeutic type I IFN administration is associated with the appearance of autoimmunity

(Biggioggero et al, 2010). Therefore, negative regulation of the anti-viral immune

response by Dcp2 could be a protective mechanism against deleterious overexpression of

IFNs.

In summary, in this chapter we uncovered a novel role of the Dcp2 decapping

enzyme in anti-viral immunity. It decreases the level of IRF7, a key transcription factor in

signaling of virus infection, by promoting degradation of IRF7 mRNA. Consistently a

wide range of IFN-inducible anti-viral genes were upregulated in the Dcp2β/β MEF cells.

Therefore we propose Dcp2 is a negative regulator of the anti-viral immune response.

Interestingly, Dcp2 expression is also induced upon viral infection, forming a negative

feedback loop to possibly restrict the amplitude of IFN-mediated immune response. This

regulation of IRF7 by mRNA degradation opens a new avenue of monitoring anti-viral

immunity and should permit the development of screening assays for compounds to

potentiate or decrease IFN-mediated immune response.

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Concluding remarks

mRNA degradation from 5′ end involves cleavage of the m7G cap structure, which can be carried out by Dcp2 or Nudt16 decapping enzymes. Dcp2 is an RNA binding protein that specifically binds and decaps a subset of mRNA targets. We biochemically isolated RNAs that copurify with Flag-Dcp2 and identified sequence element at the 5′ terminus of Rrp41 mRNA as a specific substrate of Dcp2. This element

(termed DBDE) directly recruits Dcp2 protein onto the 5′ end of an mRNA and enhances decapping activity both in vitro and in cells. Mutational analysis of the DBDE element revealed a stem-loop secondary structure which is required for both Dcp2 binding and decapping. Importantly, mRNAs that contain DBDE-like structures also showed enhanced decapping activity in vitro, indicating the 5′ terminal stem-loop as a general mechanism of promoting Dcp2 decapping, which can be exploited as a predictive tool to identify Dcp2 target mRNAs.

The recent identification of Nudt16 as a second cytoplasmic decapping enzyme in mice opened a new area of regulation of mRNA decapping in mammalian cells. Utilizing the hypomophic Dcp2 knockout MEF cell lines and lentiviruses that express shRNA aginst Nudt16, we studied the differential involvement of Dcp2 and Nudt16 in cellular mRNA decay pathways. Our data suggest that non-sense mediated decay (NMD) specifically uses Dcp2, while miRNA mediated gene silencing could employ Dcp2 and

Nudt16 redundantly, and Dcp2 and Nudt16 are differentially utilized in ARE-mediated decay. These facts indicate that rather than a default decapping enzyme that decaps every mRNA, Dcp2 and Nudt16 are involved in the degradation of specific mRNA substrates.

147

Moreover, Dcp2 and Nudt16 together regulate only a fraction of total cellular transcripts

(Song et al, 2010), implying that decapping is a minor aspect of mRNA decay or that it is

a highly redundant process involving multiple decapping enzymes. Consistent with the

latter possibility, we recently identified a novel decapping enzyme Rai1 with unique

catalytic activity in yeast (Jiao et al, 2010). Therefore, it appears that a number of distinct

decapping complexes exist, each with their own substrate specificities and regulations.

With a number of regulatory proteins identified for Dcp2, it is interesting to investigate

how Nudt16 decapping activity is regulated. Does it require accessory proteins? If so,

what proteins does it interact with? Is Nudt16 activity temporally and spatially regulated

in cells? Are there more decapping proteins in mammalian cells? Future experiments will

answer these questions.

The interesting finding of enhanced anti-viral immune response in Dcp2β/β MEF

cells has led us to identify IRF7, a key transcription factor in anti-viral immunity, as a

direct Dcp2 target. IRF7 mRNA was significantly stabilized with reduced Dcp2 level,

which manifested to increased IRF7 protein level in Dcp2β/β MEF cells. Consequently, genes downstream of IRF7 signaling, including IFNs, CXCL10 and OAS2 were upregulated in Dcp2β/β MEF cells upon virus infection. These facts indicate Dcp2 as a

post-transcriptional modulator of anti-viral immunity in MEF cells. Interstingly, Dcp2

level is also induced by viral infection, implying a negative feedback loop in the IFN-

mediated immune response. The importance of Dcp2 as a regulator of anti-viral immune

response needs to be further tested in the immune system and the Dcp2β/β mice. Whether

Dcp2 functions as a universal immunomodulatory factor in all cell types or is involved in cell-type specific control of anti-viral immune response remains unknown. Nevertheless,

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the regulation of innate anti-viral immunity by mRNA degradation opens a new path of

modulating IFN-mediated immune response and provides a potential target for anti-viral therapeutics.

149

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Curriculum Vita

2005 B.S. Tsinghua University, P. R. China, Biological Sciences. 2011 Ph.D. Rutgers University, Cell and Developmental Biology

Publications Li Y., Song M. and Kiledjian M. Differential utilization of decapping enzymes in mammalian mRNA decay pathways. RNA. 2011 Mar; 17(3):419-428.

Song M., Li Y. and Kiledjian M. Multiple mRNA decapping enzymes in mammalian cells. Mol. Cell 2010 Nov; 40:423-432

Li Y. and Kiledjian M. The regulation of eukaryotic mRNA decapping. Wiley Interdisciplinary Reviews RNA. 2010 Sept/Oct; 1(2):253-265.

Li Y, Ho ES, Gunderson SI, Kiledjian M. Mutational analysis of a Dcp2-binding element reveals general enhancement of decapping by 5'-end stem-loop structures. Nucleic Acids Res. 2009 Apr; 37(7):2227-37.

Li Y., Song M. and Kiledjian M. Transcript-specific decapping and regulated stability by the human Dcp2 decapping protein. Mol. Cell. Biol. 2008 Feb; 28(3):939-48.