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REGULATION OF MRNA DECAY IN S. CEREVISIAE BY THE SEQUENCE-SPECIFIC RNA-BINDING PROTEIN VTS1
by
Laura Marie Rendl
A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Graduate Department of Biochemistry
University of Toronto
© Copyright by Laura Marie Rendl 2009
ii
Regulation of mRNA Decay in S. cerevisiae by the Sequence-Specific RNA-
Binding Protein Vts1
Laura Marie Rendl
Doctor of Philosophy
Graduate Department of Biochemistry University of Toronto
2009
Abstract
Vts1 is a member of the Smaug protein family, a group of sequence-specific RNA-
binding proteins that regulate mRNA translation and degradation by binding to consensus stem-
loop structures in target mRNAs. Using RNA reporters that recapitulate Vts1-mediated decay in
vivo as well as endogenous mRNA transcripts, I show that Vts1 regulates the degradation of
target mRNAs in Saccharomyces cerevisiae. In Chapter Two, I focus on the mechanism of Vts1-
mediated mRNA decay. I demonstrate that Vts1 initiates mRNA degradation through
deadenylation mediated by the Ccr4-Pop2-Not deadenylase complex. I also show that Vts1
interacts with the Ccr4-Pop2-Not deadenylase complex suggesting that Vts1 recruits the
deadenylase machinery to target mRNAs, resulting in transcript decay. Following poly(A) tail
removal, Vts1 target transcripts are decapped and subsequently degraded by the 5’-to-3’
exonuclease Xrn1. Taken together these data suggest a mechanism of mRNA degradation that
involves recruitment of the Ccr4-Pop2-Not deadenylase to target mRNAs. Previous work in
Drosophila melanogaster demonstrated that Smg interacts with the Ccr4-Pop2-Not complex to
regulate mRNA stability, suggesting Smaug family members employ a conserved mechanism of
mRNA decay.
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In Drosophila, Smg also regulates mRNA translation through a separate mechanism
involving the eIF4E-binding protein Cup. In Chapter Three, I identify the eIF4E-associated
protein Eap1 as a component of Vts1-mediated mRNA decay in yeast. Interestingly Cup and
Eap1 share no significant homology outside of the seven amino acid eIF4E-binding motif. In
eap1Δ cells mRNAs accumulate as deadenylated capped species, suggesting that Eap1 stimulates
mRNA decapping. I demonstrate that the Eap1 eIF4E-binding motif is required for efficient
degradation of Vts1 target mRNAs and that this motif enables Eap1 to mediate an interaction
between Vts1 and eIF4E. Together these data suggest Vts1 influences multiple steps in the
mRNA decay pathway through interactions with the Ccr4-Pop2-Not deadenylase and the
decapping activator Eap1.
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Acknowledgments
I offer my sincerest gratitude to my supervisor Craig Smibert for his essential encouragement
and guidance throughout my graduate career. Thanks for introducing a second model organism
to the lab and seeing the value of yeast as more than fly food.
I also thank Grant Brown and Henry Krause for serving on my advisory committee and
providing helpful insight throughout the course of this project.
I thank members of the Smibert lab for their technical assistance and suggestions.
To Ben Pinder, who was there through the challenges and triumphs, thank-you for the helpful
scientific discussions, endless support, and, most of all, friendship that extends beyond the walls
of MSB.
I am grateful to members of the Brown and Segall labs for reagents, strains and yeast advice
along the way.
I appreciate the financial support from the Ontario Graduate Scholarship Program.
I could not have sustained the mental, physical and emotional energy to complete this project
without the tireless encouragement I received from family and friends.
A big thank-you to Tania Roberts and Jessica Vaisica for commiserating with me after failed
experiments, keeping me entertained during seminars, and, most of all, their friendship.
A very big thank-you to my sister Lianne and my parents whose faith in me and support of my
education has been both astonishingly generous and unwavering. Even if you don’t understand
what it all means I dedicate this work to you.
Lastly to my girls – Andrea, Angela, Caroline, Cesca, Liliana, Nancy and Deanne – your
encouragement has kept me smiling and believing. Thank-you for your collective support and
friendship, which has enriched my life and provided me with the motivation I needed to finish
this work.
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Table of Contents
ABSTRACT ................................................................................................................................. ii
ACKNOWLEDGMENTS ............................................................................................................... iv
TABLE OF CONTENTS................................................................................................................. v
LIST OF FIGURES ...................................................................................................................... ix
LIST OF ABBREVIATIONS .......................................................................................................... xi
CHAPTER 1: INTRODUCTION...................................................................................................... 1
1.1 PATHWAYS OF MRNA DECAY ......................................................................................... 2
1.1.1 Deadenylation-Dependent Exonucleolytic Digestion ................................................ 4
1.1.1.1 Enzymes that Catalyze Deadenylation .............................................................. 4
1.1.1.2 mRNA Decapping............................................................................................. 6
1.1.1.3 Enhancers of Decapping .................................................................................. 6
1.1.1.4 5’-to-3’ Exonucleolytic Decay .......................................................................... 7
1.1.1.5 3’-to-5’ Exonucleolytic Decay .......................................................................... 7
1.1.1.6 Processing Bodies ............................................................................................ 8
1.1.1.7 The Competition Between Translation and Decapping ..................................... 9
1.1.1.8 Dhh1 and Pat1 Influence mRNA Translation and Decapping ..........................10
1.1.2 mRNA Decay Via Endonucleolytic Cleavage..........................................................12
1.1.3 Decay Pathways for mRNA Quality Control ...........................................................12
1.1.3.1 Nonsense-Mediated Decay ..............................................................................13
1.1.3.2 Nonstop Decay ................................................................................................14
1.1.3.3 No-Go Decay ..................................................................................................15
1.1.3.4 Ribosome Extension-Mediated Decay..............................................................16
1.2 REGULATION OF MRNA STABILITY BY SEQUENCE-SPECIFIC TRANS-ACTING FACTORS ....16
1.2.1 Regulation of Endonucleolytic Cleavage .................................................................17
1.2.1.1 Vigilin Regulates mRNA Cleavage in Xenopus laevis ......................................17
1.2.1.2 mRNA Regulation by the Iron-Regulatory Element..........................................18
1.2.1.3 RNAi Induced Endonucleolytic Cleavage ........................................................18
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1.2.2 Elements that Recruit the Deadenylase Complex .....................................................19
1.2.3 miRNA Regulation of mRNA Decay.......................................................................21
1.2.4 The Stabilizing and Destabilizing Effects of A+U-Rich Elements............................22
1.3 POST-TRANSCRIPTIONAL REGULATION BY THE SMG PROTEIN FAMILY.............................23
1.3.1 Drosophila Smg ......................................................................................................25
1.3.1.1 Post-Transcriptional Regulation of nos mRNA ................................................25
1.3.1.2 Smg-Mediated Regulation of nos mRNA..........................................................27
1.3.1.3 Post-Transcriptional Regulation of Hsp83 mRNA ...........................................30
1.3.1.4 Smg-Mediated Regulation of Hsp83 mRNA .....................................................30
1.3.1.5 Smg is a Major Regulator of mRNA Stability...................................................31
1.3.2 Mammalian Smg.....................................................................................................31
1.3.3 Vts1, the Smg Homolog in Saccharomyces cerevisiae .............................................33
1.4 THESIS RATIONALE ........................................................................................................35
CHAPTER 2: S. CEREVISIAE VTS1 INDUCES DEADENYLATION-DEPENDENT TRANSCRIPT
DEGRADATION AND INTERACTS WITH THE CCR4-POP2-NOT DEADENYLASE COMPLEX..........36
2.1 ABSTRACT .....................................................................................................................37
2.2 INTRODUCTION...............................................................................................................38
2.3 MATERIALS AND METHODS ............................................................................................40
2.3.1 Yeast Strains and Media..........................................................................................40
2.3.2 mRNA Analysis ......................................................................................................40
2.3.3 Immunoprecipitations..............................................................................................41
2.3.4 Confocal Microscopy ..............................................................................................42
2.4 RESULTS ........................................................................................................................42
2.4.1 Ccr4 is Required for Vts1-Mediated mRNA Decay .................................................42
2.4.2 Vts1 Target mRNAs Undergo Ccr4-Dependent Deadenylation................................43
2.4.3 Vts1 Interacts with the Ccr4-Pop2-Not Deadenylase Complex ................................47
2.4.4 Vts1-Mediated Decay Requires mRNA Decapping and 5’-to-3’ Degradation ..........47
2.4.5 Vts1-Mediated Decay of an Endogenous Target mRNA Requires Ccr4 and Xrn1....51
2.4.6 Vts1 Localizes to P-Bodies in xrn1Δ Cells ..............................................................55
2.5 DISCUSSION ...................................................................................................................57
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2.5.1 Smg Family Members Employ a Conserved Mechanism to Induce Transcript
Degradation.......................................................................................................................57
2.5.2 The Role of Foci in the Function of Smg Family Members......................................58
CHAPTER 3: THE EIF4E-BINDING PROTEIN EAP1 FUNCTIONS IN VTS1-MEDIATED
TRANSCRIPT DECAY .................................................................................................................60
3.1 ABSTRACT .....................................................................................................................61
3.2 INTRODUCTION...............................................................................................................62
3.3 MATERIALS AND METHODS ............................................................................................64
3.3.1 Yeast Strains and Media..........................................................................................64
3.3.2 mRNA Analysis ......................................................................................................64
3.3.3 Immunoprecipitations..............................................................................................66
3.4 RESULTS ........................................................................................................................67
3.4.1 Eap1 is Required for Efficient Decay of Vts1 Target mRNAs..................................67
3.4.2 Eap1 is Not a Core Component of the mRNA Decay Machinery .............................70
3.4.3 Eap1 Does Not Function in mRNA Deadenylation ..................................................72
3.4.4 Eap1 Stimulates mRNA Decapping.........................................................................74
3.4.5 Eap1 Mediates an Interaction Between Vts1 and eIF4E...........................................75
3.4.6 Binding to eIF4E is Required for Eap1 to Stimulate Transcript Decay.....................78
3.4.7 Eap1 Enhances the Degradation of Another Endogenous mRNA.............................80
3.5 DISCUSSION ...................................................................................................................82
3.5.1 The Role of Eap1 in mRNA Decapping...................................................................82
3.5.2 The Function of Eap1 in Yeast ................................................................................83
3.5.3 The Role of eIF4E-Binding Proteins in mRNA Decay.............................................84
CHAPTER 4: CONCLUSIONS AND FUTURE DIRECTIONS.............................................................85
4.1 CONCLUSIONS ................................................................................................................86
4.2 FUTURE DIRECTIONS ......................................................................................................88
4.2.1 Testing Direct Recruitment of the Deadenylase to Target mRNAs...........................88
4.2.2 Testing Recruitment of Eap1 to Vts1 Target mRNAs ..............................................92
4.2.3 Determining the Eap1 Mechanism of mRNA Degradation.......................................94
4.2.3.1 Confirming Eap1 as an Activator of Decapping ..............................................94
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4.2.3.2 Does Eap1 Influence eIF4E Cap-Affinity?.......................................................95
4.2.3.3 Does Eap1 Recruit Decapping Activators?......................................................96
4.2.4 Identification and Characterization of Eap1 Target mRNAs ....................................97
4.2.4.1 The Role of Eap1 in PUF Target Regulation ...................................................99
REFERENCES ..........................................................................................................................102
COPYRIGHT ACKNOWLEDGMENTS .........................................................................................117
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List of Figures
Figure 1-1 Eukaryotic mRNA Decay Pathways ......................................................................... 3
Figure 1-2 The Smg Family of RNA-Binding Proteins .............................................................24
Figure 1-3 Model for Smg-Mediated Repression of nos mRNA Translation .............................29
Figure 1-4 Model for Smg-Mediated Decay of Hsp83 mRNA ..................................................32
Figure 2-1 Vts1 Stimulates the Decay of Target mRNAs ..........................................................44
Figure 2-2 Vts1 Induces the Deadenylation of Target mRNAs..................................................46
Figure 2-3 Vts1 Interacts with Pop2 .........................................................................................48
Figure 2-4 Vts1-Mediated mRNA Decay Requires Components of the Major mRNA Decay
Pathway ....................................................................................................................................50
Figure 2-5 Deadenylation is the First Step in Vts1-Mediated mRNA Decay .............................52
Figure 2-6 Stability of YIR016W mRNA is Regulated by Vts1 .................................................54
Figure 2-7 Vts1 Localizes to P-Bodies in xrn1Δ Cells ..............................................................56
Figure 3-1 Eap1 is Required for Efficient Degradation of Vts1 Target mRNAs ........................68
Figure 3-2 Eap1 is Not Required for the Degradation of GFP-SRE- mRNA..............................71
Figure 3-3 Eap1 is Not Required for Transcript Deadenylation.................................................73
Figure 3-4 Eap1 Stimulates Transcript Decapping ....................................................................76
Figure 3-5 Eap1 Mediates an Indirect Interaction Between Vts1 and eIF4E..............................77
Figure 3-6 The Eap1/eIF4E Interaction is Required for Efficient Decay of Vts1 Target
mRNAs.....................................................................................................................................79
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Figure 3-7 The Stability of COX17 mRNA is Regulated by Eap1 .............................................81
Figure 4-1 Model for Vts1-Mediated mRNA Decay .................................................................87
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List of Abbreviations
4E-T.................................................................................................................. eIF4E-transporter
ACT1.................................................................................................................................. actin 1
Ago............................................................................................................................... argonaute
ARE................................................................................................................. A+U-rich element
bcd ......................................................................................................................................bicoid
cycB ................................................................................................................................. cyclin B
DOX ..........................................................................................................................doxycycline
dsRNA ........................................................................................................double-stranded RNA
DTT ......................................................................................................................... dithiothreitol
Eap1.................................................................................................... eIF4E-associated protein 1
EDBR ................................................................................................. Eap1/Dhh1 binding region
EDTA ...................................................................................... ethylene-diamine-tetra-acetic acid
eIF ...................................................................................................... eukaryotic initiation factor
EJC ........................................................................................................... exon junction complex
ELAV .......................................................................................embryonic lethal abnormal vision
EVIR................................................................................................ Eap1/Vts1 interaction region
FBP............................................................................................................ FUSE-binding protein
GFP ....................................................................................................... green fluorescent protein
glo.................................................................................................................................... glorund
GST ....................................................................................................... glutathione S-transferase
hb.................................................................................................................................hunchback
HEPES..........................................................4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
hnRNP ...........................................................................heterogeneous nuclear ribonucleoprotein
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hsp83 .......................................................................................................... heat shock protein 83
HuR .........................................................................................................................Hu-antigen R
IRE .............................................................................................................iron response element
IRE-BP ......................................................................................................... IRE-binding protein
m7G-cap....................................................................................................7-methylguanosine cap
m7GDP.........................................................................................7-methylguanosine diphosphate
mCRD............................................................................major protein coding-region determinant
miRNA ...................................................................................................................... micro RNA
mRFP...................................................................................... monomeric red fluorescent protein
mRNA ................................................................................................................ messenger RNA
mRNP .............................................................................................. messenger ribonucleoprotein
NGD ..........................................................................................................................no-go decay
NMD..................................................................................................... nonsense-mediated decay
nos ...................................................................................................................................... nanos
NP-40 .......................................................................................................................nonidet P-40
NSD....................................................................................................................... nonstop decay
nt................................................................................................................................. nucleotides
ORF ................................................................................................................open reading frame
osk ....................................................................................................................................... oskar
PABP.......................................................................................................poly(A) binding protein
PAGE .....................................................................................polyacrylamide gel electrophoresis
PAN................................................................................................................... poly(A) nuclease
PARN .......................................................................................................... poly(A) ribonuclease
P-bodies ............................................................................................................ processing bodies
PEBR....................................................................................................Puf3/Eap1 binding region
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PTC .................................................................................................premature termination codon
PUF ................................................................................................. Pumillio/FBP protein family
pum..................................................................................................................................pumillio
RBD............................................................................................................ RNA binding domain
REMD ................................................................................... ribosome extension-mediated decay
RISC........................................................................................... RNA-induced silencing complex
RNA ..................................................................................................................... ribonucleic acid
RNAi ................................................................................................................. RNA interference
RNase ........................................................................................................................ ribonuclease
RNP ................................................................................................................... ribonucleoprotein
rump ....................................................................................................................... rumpelstiltskin
SAM ..................................................................................................................sterile alpha motif
SCR1......................................................................................................small cytoplasmic RNA 1
SDS ........................................................................................................... sodium dodecyl sulfate
siRNA..........................................................................................................short interfering RNA
smg ......................................................................................................................................smaug
SMG .................................................................suppressor with morphogenetic effect on genitalia
SPR..................................................................................................... surface plasmon resonance
SRE ................................................................................................... Smaug recognition element
SSR......................................................................................................... Smaug similarity region
TCA................................................................................................................ trichloroacetic acid
TCE ................................................................................................. translational control element
Tet ..............................................................................................................................tetracycline
TfR ................................................................................................................. transferrin receptor
TLC .....................................................................................................thin layer chromatography
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Tris ....................................................................................... tris (hydroxymethyl) aminomethane
tRNA .......................................................................................................................transfer RNA
TTP........................................................................................................................ Tristetraprolin
UTR................................................................................................................untranslated region
uv..........................................................................................................................ultraviolet light
VDIR .................................................................................... Vts1 deadenylase interaction region
VDR ....................................................................................................Vts1 deadenylation region
Vts1 .............................................................................................................. VTI1-2 suppressor 1
1
Chapter 1
Introduction
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Chapter 1
Introduction
1.1 Pathways of mRNA Decay
The conservation of several mRNA degradation pathways in eukaryotes highlights the
importance of mRNA turnover as a means to control gene expression. Regulation of mRNA
stability is a way to effect rapid change in response to environmental cues, block the expression
of mutant genes or regulate the subcellular localization of a particular gene product. Given these
functions, it is important to understand how mRNAs are targeted for destruction. In eukaryotes,
the mRNA 5’ 7-methylguanosine (m7G) cap and the 3’ poly(A) tail play important roles in the
regulation of mRNA stability. In general, both structures protect mRNAs from exonucleolytic
decay and as such their removal is often an important step in transcript degradation. In addition,
both play essential roles in translation initiation through their ability to recruit translation factors
to transcripts. The essential roles of the 5’ m7G-cap and poly(A) tail in the regulation of
transcript stability and mRNA translation highlights the fact that these processes are tightly
coupled.
In eukaryotes the degradation of mRNAs can occur through four general mechanisms:
deadenylation-dependent 5’-to-3’ decay, deadenylation-dependent 3’-to-5’ decay,
endonucleolytic cleavage, or by specialized quality control pathways (Figure 1-1). Although the
major pathway for degradation may vary among species, all organisms have multiple
mechanisms in place for mRNA turnover.
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Figure 1-1 Eukaryotic mRNA Decay Pathways
(A) Shown is a schematic representation of the deadenylation-dependent pathways of mRNA
decay. The initial step in this pathway is shortening of the mRNA poly(A) tail. Following
deadenylation, the transcript is decapped and subsequently degraded by a 5’-to-3’
exoribonuclease (left). This pathway is known as the major mRNA decay pathway in yeast.
Alternatively following deadenylation, the transcript is degraded 3’-to-5’ by a complex of
exoribonucleases known as the exosome (right). (B) Sequence-specific endoribonucleases cleave
mRNAs internally. This cleavage event generates free 3’ and 5’ ends that are rapidly degraded
by cellular exoribonucleases.
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1.1.1 Deadenylation-Dependent Exonucleolytic Digestion
Two pathways by which mRNAs can be degraded both begin with shortening of the
poly(A) tail. Deadenylation can trigger mRNA decapping followed by 5’-to-3’
exoribonucleolytic decay of the message (Hsu & Stevens, 1993; Muhlrad et al., 1994, 1995;
Caponigro & Parker, 1996a). Deadenylation-dependent decapping followed by 5’-to-3’
exonucleolytic decay is considered the major mRNA decay pathway in Saccharomyces
cerevisiae (Decker & Parker, 1993; Hsu & Stevens, 1993; Muhlrad et al., 1994, 1995; Caponigro
& Parker, 1996a). In an alternative pathway, following poly(A) tail shortening, transcripts can be
degraded 3’-to-5’ by a complex of exonucleases known as the exosome (van Hoof & Parker,
1999; Mitchell & Tollervey, 2000; Butler, 2002).
1.1.1.1 Enzymes that Catalyze Deadenylation
mRNA degradation often begins with shortening of the poly(A) tail. To date three
enzyme complexes have been identified as mRNA deadenylases [reviewed in (Parker & Song,
2004)]. In yeast, the major deadenylase complex contains two nucleases, Ccr4 and Pop2, and
several accessory proteins, Not1-Not5, Caf4, Caf16, Caf40, and Caf130 (Bai et al., 1999; Chen
et al., 2001b; Liu et al., 2001; Tucker et al., 2001; Denis & Chen, 2003). Ccr4 is a member of the
Mg2+-dependent endonuclease-related protein family, which is conserved among eukaryotes
(Dlakic, 2000). In yeast, disruption of the CCR4 gene or mutation of key residues in the protein
active site show a strong defect in poly(A) tail removal, implicating Ccr4 as the major catalytic
component of the Ccr4-Pop2-Not deadenylase complex (Daugeron et al., 2001; Tucker et al.,
2001; Tucker et al., 2002). Pop2 is a member of the RNase D family of 3’-to-5’ exonucleases,
however the yeast protein contains non-canonical residues at three of the five residues involved
in formation of the catalytic site (Daugeron et al., 2001; Tucker et al., 2001). Pop2 is considered
5
an essential component of the major yeast deadenylase complex since cells lacking Pop2
accumulate mRNAs with extended poly(A) tails, but the catalytic activity of Pop2 in vivo
remains unclear (Daugeron et al., 2001; Chen et al., 2002; Tucker et al., 2002). Currently Pop2 is
thought to provide structural integrity to the deadenylase complex and stimulate the catalytic
activity of Ccr4.
Cytoplasmic mRNA deadenylation can also be catalyzed by the poly(A) nuclease (PAN)
complex, which consists of the Pan2 and Pan3 proteins. The catalytic subunit Pan2 is a member
of the RNase D superfamily (Moser et al., 1997), while the role of Pan3 in this complex remains
to be determined. In yeast, the function of the PAN complex in poly(A) tail removal is revealed
in ccr4Δ cells where the predominant deadenylase is absent and residual deadenylation is
dependent on Pan2 (Tucker et al., 2001). The complete block of deadenylation in a ccr4Δ/pan2Δ
double mutant identifies PAN as a second cytoplasmic deadenylase in yeast (Tucker et al.,
2001). Similar studies in mammals have illustrated biphasic kinetics of deadenylation, with PAN
initiating deadenylation followed by Ccr4-mediated shortening (Yamashita et al., 2005). PAN is
also involved in an early step of yeast poly(A) metabolism in which an initially long tail is
trimmed to the standard length of 60 to 80 nucleotides in the cytoplasm (Brown & Sachs, 1998).
Poly(A) ribonuclease (PARN) has been purified and characterized from mammalian cells
and Xenopus laevis oocytes (Parker & Song, 2004). PARN is a member of the RNase D
superfamily and mutation of predicted catalytic residues inhibit deadenylase activity (Ren et al.,
2002; Lai et al., 2003). PARN homologs are found in many eukaryotes but are notably absent
from Drosophila and S. cerevisiae (Parker & Song, 2004). Biochemical studies have shown that
PARN binds directly to the mRNA cap structure, and that this interaction stimulates its
deadenylase activity (Dehlin et al., 2000). Cap analogues inhibit PARN deadenylase activity but
6
stimulate the mRNA decapping enzyme (Dehlin et al., 2000; Gao et al., 2000). In the current
model of PARN activity, the interaction of PARN with the mRNA cap inhibits decapping during
poly(A) shortening (Parker & Song, 2004). Once deadenylation is complete, PARN dissociates
from the transcript to allow the decapping enzyme access to the mRNA (Parker & Song, 2004).
1.1.1.2 mRNA Decapping
Two proteins, Dcp1 and Dcp2, function together as the decapping enzyme in yeast
[reviewed in (Coller & Parker, 2004)]. Dcp2 is the catalytic subunit of the decapping complex,
and Dcp1 functions as a critical cofactor to enhance Dcp2 activity. Dcp1 strongly stimulates
Dcp2 catalytic activity while having little effect on the interaction of Dcp2 with the m7G-cap
(Deshmukh et al., 2008). Structural evidence suggests that this process involves the
transformation of Dcp2 from an inactive open conformation to an active closed conformation,
which orients the Dcp2 N-terminus toward the catalytic site and renders Dcp2 catalytically active
(Deshmukh et al., 2008; She et al., 2008). Removal of the m7G-cap structure by the decapping
enzyme yields a m7GDP product and a 5’-monophosphate mRNA (Stevens, 1980; LaGrandeur &
Parker, 1998).
1.1.1.3 Enhancers of Decapping
Several proteins, including Edc1, Edc2, Edc3, the Lsm1-7 complex, Pat1 and Dhh1, have
been identified as enhancers of decapping [reviewed in (Coller & Parker, 2004)]. While not
required for decapping per se, these factors are important for efficient decapping as mutations in
the genes encoding them can alter the decay rate of specific mRNAs. Edc1 and Edc2, which are
specific to yeast, appear to specifically stimulate Dcp2 cap-binding/catalysis because, like Dcp1,
they stimulate Dcp2 decapping in vitro (Schwartz et al., 2003; Steiger et al., 2003). Edc3 is
conserved throughout eukaryotes and most likely stimulates Dcp2 recruitment (Franks & Lykke-
7
Andersen, 2008). Cellular depletion of Edc3 impairs decapping of a subset of mRNAs (Badis et
al., 2004; Kshirsagar & Parker, 2004; Coller & Parker, 2005). Edc3 interacts directly with
multiple decapping factors, including Dcp1 and Dcp2, suggesting that it directly recruits or
activates the decapping complex on target mRNAs (Decker et al., 2007; Tritschler et al., 2007).
The specific roles of the Lsm1-7 complex, Pat1 and Dhh1 in decapping are discussed below.
1.1.1.4 5’-to-3’ Exonucleolytic Decay
Following decapping, the resulting mRNA with an exposed 5’-monophosphate is rapidly
degraded by the 5’-to-3’ exonuclease Xrn1 (Stevens & Maupin, 1987). This protein is a member
of a large family of conserved exonucleases, although little is known about the catalytic
mechanism of its members (Parker & Song, 2004). Xrn1 has been shown to degrade uncapped
mRNAs in vitro (Larimer & Stevens, 1990). In yeast, xrn1Δ cells accumulate uncapped mRNA
decay intermediates and the presence of a strong secondary structure near the 5’ end of a
transcript inhibits exonucleolytic decay (Hsu & Stevens, 1993; Muhlrad et al., 1994; Poole &
Stevens, 1997). From these data it was concluded that following decapping Xrn1 degrades
mRNAs in a 5’-to-3’ direction.
1.1.1.5 3’-to-5’ Exonucleolytic Decay
Deadenylation is a major precursor for the decapping of yeast mRNAs (Muhlrad &
Parker, 1994), however in higher organisms the link between deadenylation and decapping is
less clearly defined (Franks & Lykke-Andersen, 2008). In an alternative pathway following
deadenylation mRNAs can be degraded 3’-to-5’ by the exosome, a large protein complex
containing multiple 3’-to-5’ exonucleases (van Hoof & Parker, 1999; Mitchell & Tollervey,
2000; Butler, 2002). The nine core exosome subunits are predicted to form a ringlike structure
that must interact with the heterotrimeric complex of Ski2, Ski3 and Ski8 to degrade cytoplasmic
8
mRNAs [reviewed in (Parker & Song, 2004)]. The 3’-to-5’ mechanism of mRNA decay is
considered a second general pathway of mRNA turnover in yeast. This is supported by the
synthetic lethal phenotype observed when mutations that inactivate the 5’-to-3’ pathway are
combined with mutations that inactivate the 3’-to-5’ pathway, arguing that mRNA decay by
these pathways is essential for yeast viability (Anderson & Parker, 1998). Although 5’-to-3’
decay is traditionally considered the major pathway, microarray experiments suggest that <20%
of yeast mRNAs are significantly up-regulated in dcp1Δ or xrn1Δ strains, suggesting other
mechanisms like the exosome may play a more substantial role in global mRNA turnover (He et
al., 2003).
1.1.1.6 Processing Bodies
Destruction of translating mRNAs seems to involve transition from an actively
translating form to one consistent with mRNA decay. This transition is thought to involve the
aggregation of mRNPs into distinct cytoplasmic foci, known as processing bodies (P-bodies)
(Sheth & Parker, 2003; Coller & Parker, 2005; Teixeira et al., 2005). Although the exact protein
composition of P-bodies is yet to be determined, a core group of proteins found in P-bodies is
conserved from yeast to mammals [reviewed in (Parker & Sheth, 2007)]. In this group are the
decapping proteins Dcp1 and Dcp2, the decapping activators Dhh1, Pat1, Edc3 and the Lsm1-7
complex, as well as the 5’-to-3’ exonuclease Xrn1 [reviewed in (Parker & Sheth, 2007)]. The
Ccr4-Pop2-Not deadenylase complex also accumulates in mammalian P-bodies (Cougot et al.,
2004; Andrei et al., 2005) and in certain yeast mutants that are defective in mRNA decay
(Teixeira & Parker, 2007). P-bodies also contain RNA, as treatment with RNase in vitro or in
permeabilized cells causes P-body disassembly (Teixeira et al., 2005; Eulalio et al., 2007b).
9
P-bodies are highly dynamic structures formed by the assembly and disassembly of
translationally repressed mRNPs [reviewed in (Franks & Lykke-Andersen, 2008)]. The direct
relationship between P-body formation and the concentration of translationally repressed mRNPs
in the cytoplasm is supported by several observations. The entrapment of mRNAs in polysomes
by cycloheximide treatment causes rapid disappearance of P-bodies (Cougot et al., 2004;
Teixeira et al., 2005). Also in yeast, glucose deprivation, which causes the release of mRNAs
from polysomes, increases P-body formation (Brengues et al., 2005; Teixeira et al., 2005).
Once assembled into P-bodies mRNAs can undergo two possible fates: they can re-enter
the translational pool or they can be degraded [reviewed in (Franks & Lykke-Andersen, 2008)].
Observations in yeast indicate that mRNAs assembled into P-bodies during glucose starvation
are released back into the translating pool once glucose is added back to cells (Brengues et al.,
2005). In addition, yeast cells in stationary phase release mRNAs from P-bodies when growth
resumes (Brengues et al., 2005). The localization of mRNA decay factors and mRNA decay
intermediates to P-bodies, suggests that some mRNAs concentrated in P-bodies are being
degraded (Sheth & Parker, 2003). The increase in size and abundance of P-bodies when the
catalytic steps of decapping or 5’-to-3’ decay are inhibited, further supports the role of P-bodies
in mRNA decay (Sheth & Parker, 2003; Andrei et al., 2005). Further investigation is required to
understand how P-bodies discriminate between mRNAs that are destined for degradation and
mRNAs that evade decay for re-entry into the translational pool.
1.1.1.7 The Competition Between Translation and Decapping
Transcript destabilization is often tightly coupled to translational control (Jacobson &
Peltz, 1996). Both the poly(A) tail and 5’ cap structure are required for translation and also
inhibit mRNA degradation. The poly(A) tail inhibits decapping through the interaction of
10
poly(A) binding protein (PABP) with the translation initiation factor eIF4G (Coller & Parker,
2004). In yeast, mutation of initiation factors eIF4E, the cap-binding protein, and eIF4G lead to
an increase in decapping (Schwartz & Parker, 1999). At the restrictive temperature, temperature
sensitive mutations in eIF4E or deletion of eIF4G result in faster mRNA decay rates (Schwartz
& Parker, 1999). The addition of purified yeast eIF4E inhibits decapping in vitro (Schwartz &
Parker, 1999; Wilusz et al., 2001a), while the addition of cap analog to mammalian extracts
enhances the mRNA decapping rate (Gao et al., 2001). This suggests that the cap-binding
complex can block access of the decapping enzyme to the mRNA 5’ cap structure. Decapping
requires remodeling of the mRNP to remove the translation initiation cap-binding complex and
recruit mRNA decapping factors (Tharun et al., 2000; Coller & Parker, 2004; Parker & Song,
2004; Teixeira et al., 2005). The first step of decapping involves repression of mRNA
translation. However, since mRNAs can be kept stably in a translationally repressed state, not all
mRNAs that are translationally repressed are decapping substrates (Brengues et al., 2005;
Teixeira et al., 2005). It is thought that only the translational repression events that cause
destabilization of the translation initiation complex will stimulate decapping (Franks & Lykke-
Andersen, 2008). Currently little is known about how translation initiation factors are exchanged
for the decapping enzyme.
1.1.1.8 Dhh1 and Pat1 Influence mRNA Translation and Decapping
In yeast, Dhh1 and Pat1 appear to activate decapping and promote translational
repression. Both proteins can function as translational repressors in response to various
environmental stresses (Coller & Parker, 2005). In addition, overexpression of Dhh1 or Pat1
causes general repression of yeast mRNA translation, and codepletion of Pat1 and Dhh1 prevents
the general translational repression observed upon glucose starvation (Coller & Parker, 2005).
11
Studies on dhh1Δ/pat1Δ double mutants revealed severe defects in mRNA decapping and
translational repression in response to glucose deprivation (Coller & Parker, 2005).
Dhh1 belongs to a conserved family of DexD/H box ATPases (Coller et al., 2001). The
function of Dhh1 in mRNA translation and its localization to P-bodies is conserved from yeast to
mammals (Sheth & Parker, 2003; Cougot et al., 2004; Coller & Parker, 2005; Barbee et al.,
2006). It has been previously proposed that Dhh1 might dissociate eIF4E from the mRNA cap
structure as a means of both promoting translational repression and mRNA decay (Fischer &
Weis, 2002). However, Dhh1 can repress the translation of both capped and uncapped mRNAs,
arguing that Dhh1 inhibits translation independent of cap recognition (Coller & Parker, 2005). In
dhh1Δ cells, insertion of a strong secondary structure into the 5’UTR of an unstable mRNA
exhibits a wild-type rate of decay (Coller & Parker, 2005). This demonstrates that repressing
mRNA translation via the stem-loop structure bypasses the need for Dhh1 and illustrates that the
role of Dhh1 as a translational repressor is functionally linked to its role in mRNA decapping.
Currently, there are two possible mechanisms proposed in yeast by which Dhh1 might
inhibit translation. In the simplest model Dhh1 promotes the assembly of a translation repression
complex which sequesters the mRNA to P-bodies (Coller & Parker, 2005). The movement of
translationally repressed mRNPs to P-bodies may also facilitate mRNA decapping. Alternatively
there is some evidence that suggests Dhh1 may directly affect the association of the 40S
ribosome with mRNAs (Coller & Parker, 2005).
Pat1 is also a conserved protein that localizes to P-bodies in yeast, Drosophila and
mammalian cells (Coller & Parker, 2005; Eulalio et al., 2007a; Scheller et al., 2007). Yeast
strains lacking Pat1 show the strongest defects in decapping of any mutant besides defects in the
12
decapping enzyme itself (Pilkington & Parker, 2008). Structure-function analysis has identified
two functional domains within Pat1: one that mediates mRNA decapping and another that
functions in translational repression (Pilkington & Parker, 2008). Pat1 together with a complex
of Lsm proteins, the Lsm1-7 complex, promotes decapping of deadenylated yeast mRNAs
(Bouveret et al., 2000; Tharun et al., 2000; Tharun & Parker, 2001). Unlike Dhh1, the role of
Pat1 in mRNA decapping does not depend on translational repression (Coller & Parker, 2005).
The interaction of Pat1 with multiple decapping factors, including Edc3, Dcp1 and Dcp2,
suggests that Pat1 may assist in recruiting the decapping complex to deadenylated mRNAs that
have accumulated in P-bodies (Tharun et al., 2000; Tharun & Parker, 2001; Pilkington & Parker,
2008). A region required for Pat1-mediated translational repression and P-body association was
identified by structure-function analysis (Pilkington & Parker, 2008). The mechanism of Pat1-
mediated translational repression awaits further investigation.
1.1.2 mRNA Decay Via Endonucleolytic Cleavage
Eukaryotic mRNAs can be degraded by endonucleolytic cleavage independent of
deadenylation. Sequence-specific endonuclease target sites in transcripts are recognized and
cleaved, and the resulting fragments readily degraded by cytoplasmic exonucleases (Beelman &
Parker, 1995). In some cases endonucleolytic cleavage is regulated by proteins that bind near the
cleavage site, rendering the site inaccessible to nucleolytic attack (Jacobson & Peltz, 1996).
Specific examples of this type of regulation are discussed below.
1.1.3 Decay Pathways for mRNA Quality Control
Eukaryotic cells employ many quality control systems to regulate cytoplasmic mRNAs
[reviewed in (Doma & Parker, 2007)]. These systems are essential to prevent the accumulation
and misexpression of nonfunctional mRNAs. Aberrant RNAs are generated at many points
13
during the biogenesis of a mature mRNA. They are produced as a result of inaccurate
transcription, improper processing or mutations in the DNA template (van Hoof et al., 2002).
The various pathways of mRNA quality control depend on specific proteins that target aberrant
transcripts for degradation.
1.1.3.1 Nonsense-Mediated Decay
Nonsense-mediated mRNA decay (NMD) selectively degrades mRNAs harboring
premature termination (nonsense) codons (PTCs) [reviewed in (Chang et al., 2007)]. These
mRNAs are degraded to prevent their translation and the production of truncated proteins with
potentially deleterious effects. The core NMD machinery consists of three proteins: UPF1,
UPF2, and UPF3. Deletion or silencing of genes encoding these proteins results in the
stabilization of PTC-containing mRNAs in all organisms in which NMD has been identified
(Hodgkin et al., 1989; Leeds et al., 1991; Cui et al., 1995; He et al., 1997; Page et al., 1999;
Lykke-Andersen et al., 2000; Serin et al., 2001; Mendell et al., 2002; Gatfield et al., 2003). Four
additional factors, known as suppressor with morphogenetic effect on genitalia (SMG) proteins,
assist in NMD by regulating the phosphorylation of UPF1: SMG1, SMG5, SMG6, and SMG7
(Cali et al., 1999; Page et al., 1999; Denning et al., 2001; Pal et al., 2001; Yamashita et al., 2001;
Anders et al., 2003; Chiu et al., 2003; Ohnishi et al., 2003; Grimson et al., 2004).
UPF1, is a group I helicase recruited to mRNAs by the translation machinery upon
recognition of PTCs (Czaplinski et al., 1998; Kashima et al., 2006). Commitment of the UPF1-
bound mRNA for degradation by NMD is triggered by the SMG1-mediated phosphorylation of
UPF1, which is dependent on UPF2 and UPF3. SMG7 then binds phosphorylated UPF1 and
recruits decay enzymes to degrade the PTC-containing transcript (Anders et al., 2003; Ohnishi et
al., 2003; Unterholzner & Izaurralde, 2004; Fukuhara et al., 2005). In yeast and mammals, both
14
the 5’-to-3’ and 3’-to-5’ pathway readily degrade PTC-bearing mRNAs [reviewed in (Garneau et
al., 2007:Parker, 2004 #2156)]. However in Drosophila, which lack a SMG7 ortholog, NMD
substrates are initially cleaved by an unidentified endonuclease (Gatfield & Izaurralde, 2004).
The resulting 3’ and 5’ fragments are subsequently degraded by Xrn1 and the exosome
respectively (Gatfield & Izaurralde, 2004).
In mammalian cells UPF2 and UPF3 are part of the Exon Junction Complex (EJC), a
large protein complex deposited upstream of exon-exon junctions during RNA splicing.
Recognition of PTCs results from the cross-talk between terminating ribosomes and a
downstream EJC comprising UPF2 and UPF3 (Baker & Parker, 2004; Maquat, 2004). According
to the current model, if translating ribosomes encounter a stop codon upstream of an EJC, UPF1
is recruited by translation release factors and interacts with the UPF2 and UPF3 proteins bound
to the downstream EJC (Baker & Parker, 2004; Maquat, 2004). In both Drosophila and S.
cerevisiae, PTC definition occurs independently of exon-exon boundaries (Gatfield et al., 2003;
Baker & Parker, 2004; Maquat, 2004). Currently it is thought that the main NMD determinant in
these organisms is the distance between the stop codon and the poly(A) tail (Amrani et al.,
2004).
1.1.3.2 Nonstop Decay
Eukaryotic mRNAs that do not contain a termination codon, referred to as “nonstop”
transcripts, are rapidly degraded (Frischmeyer et al., 2002). The rapid decay of these transcripts
is referred to as nonstop decay (NSD) and requires translation of the mRNA (Frischmeyer et al.,
2002). In yeast, nonstop transcripts are rapidly degraded in ccr4Δ, xrn1Δ, dcp1Δ and upf1Δ cells,
indicating that degradation of nonstop transcripts does not require any of the factors involved in
the major mRNA decay pathway or NMD (Frischmeyer et al., 2002). Instead, nonstop transcripts
15
are stabilized in cells depleted for core components of the exosome (van Hoof et al., 2002).
Treatment with cycloheximide or depletion of charged tRNAs significantly increased the
stability of nonstop transcripts, suggesting that translation is required for nonstop decay
(Frischmeyer et al., 2002). Ski7, a paralog of eukaryotic translation elongation factors eRF1A
and eRF3, is an important component of NSD. Recognition of nonstop mRNAs requires the
binding of Ski7 to an empty aminoacyl-(RNA-binding) site (A site) on the ribosome (van Hoof
et al., 2002). Binding of Ski7 recruits the exosome to a transcript with a ribosome stalled near the
3’ end (van Hoof et al., 2002).
1.1.3.3 No-Go Decay
Yeast mRNAs with strong pauses in translation elongation are targeted for
endonucleolytic cleavage in a process known as No-Go decay (NGD) [reviewed in (Doma &
Parker, 2007)]. Insertion of a stable stem-loop structure into the coding region of reporter
mRNAs causes ribosome stalling and subsequent transcript decay (Doma & Parker, 2006). NGD
is initiated by cleavage of the mRNA close to the ribosome stall site (Doma & Parker, 2006).
This break generates entry sites for Xrn1 and the exosome, which degrade the remainder of the
transcript (Doma & Parker, 2006). Cleavage triggered by NGD requires translation because the
termination of translation before the obstruction protects the mRNA from degradation (Doma &
Parker, 2006). Initial cleavage of the mRNA is promoted by Hbs1 and Dom34, which are similar
to the translation termination factors eRF1 and eRF3 (Doma & Parker, 2006, 2007). It is
hypothesized that Hbs1 and Dom34 interact with the stalled ribosome, however the mechanism
of mRNA cleavage in NGD is unknown.
16
1.1.3.4 Ribosome Extension-Mediated Decay
Ribosome extension-mediated decay (REMD) represses the expression of C-terminally
extended proteins. These proteins are generated when ribosomes inappropriately translate and
then terminate within the mRNA 3’UTR (Doma & Parker, 2007). In humans, REMD represses
protein synthesis from the human α-globin gene containing an antitermination mutation (Kong &
Liebhaber, 2007). Antitermination transcripts are stabilized by mutation of the initiation codon,
which blocks translation (Weiss & Liebhaber, 1994). This indicates that, like other mRNA
quality control pathways, REMD is dependent on translation. REMD degrades these transcripts
by accelerated shortening of the poly(A) tail through an unknown mechanism (Kong &
Liebhaber, 2007). Currently REMD has only been observed in erythroid cells (Kong &
Liebhaber, 2007) and future investigations are needed to understand the cell-type specificity of
this pathway.
1.2 Regulation of mRNA Stability by Sequence-Specific Trans-Acting Factors
The decay of many mRNAs is regulated by cis-acting elements found within the
transcript body. These elements are recognized by trans-acting factors and can have a stabilizing
or destabilizing effect on their targets. In some cases the recruitment of trans-acting factors has a
direct effect, such as blocking access of an endonuclease cleavage site. In other cases these
factors regulate mRNA turnover by recruiting other trans-acting factors, like components of the
degradation machinery, to the transcript. Examples of the different mechanisms used by RNA-
binding proteins to regulate mRNA decay are discussed in this section.
17
1.2.1 Regulation of Endonucleolytic Cleavage
As mentioned previously, some mRNAs are degraded by endonucleases that recognize
specific sequence elements within the transcript and cleave the mRNA internally [reviewed in
(Wilusz et al., 2001b)]. The cleavage event generates exposed 3’ and 5’ ends that are rapidly
degraded by exonucleases (Wilusz et al., 2001b). Endonuclease-catalyzed mRNA decay can be
regulated by RNA-binding proteins interacting with cis-acting stability elements to block
cleavage by site-specific mRNA endonucleases (Binder et al., 1994).
1.2.1.1 Vigilin Regulates mRNA Cleavage in Xenopus laevis
In Xenopus, estrogen induces the stabilization of vitellogenin mRNA, which encodes an
egg yolk precursor protein, and the destabilization of albumin mRNA (Brock & Shapiro, 1983;
Pastori et al., 1991). This process corresponds with upregulation of the sequence-specific mRNA
endonuclease PMR-1 and the hnRNP K homology-domain RNA-binding protein Vigilin
(Dodson & Shapiro, 1997). PMR-1 preferentially cuts RNA at a single-stranded consensus
pentamer APyrUGA (Chernokalskaya et al., 1997). Multiple copies of this consensus pentameter
are found in the vitellogenin mRNA 3’UTR and the albumin 3’UTR (Nielsen & Shapiro, 1990;
Cunningham et al., 2000). Expression of PMR-1 leads to the rapid degradation of albumin
mRNA by endonucleolytic cleavage at these sites (Nielsen & Shapiro, 1990). The upregulation
of Vigilin is required to protect vitellogenin mRNA from PMR-1 cleavage. Vigilin binds this
region of the vitellogenin 3’UTR and occludes recognition of the site and mRNA cleavage by
PMR-1 (Cunningham et al., 2000). Although PMR-1 cleavage sites are also found in the albumin
3’UTR, the affinity of Vigilin for these sites is insufficient to disrupt PMR-1 recognition and
cleavage (Cunningham et al., 2000). The differential binding affinity of Vigilin for PMR-1
cleavage sites regulates the stability of specific mRNAs during estrogen stimulation.
18
1.2.1.2 mRNA Regulation by the Iron-Regulatory Element
In higher eukaryotes, the stability of transferrin receptor (TfR) mRNA is regulated by
cellular iron concentration [reviewed in (Jacobson & Peltz, 1996)]. Iron regulation of TfR
mRNA stability is dependent on highly conserved stem-loop structures termed iron response
elements (IREs) in the 3’UTR [reviewed in (Klausner et al., 1993)]. A functional endonuclease
cleavage site is embedded within the IREs (Binder et al., 1994). When cellular iron levels are
low, IRE-binding protein (IRE-BP) binds with high affinity to IREs and thus occludes nuclease
access to the cleavage site (Binder et al., 1994). When iron levels are elevated, the interaction of
IRE-BP with IREs is weakened and endonucleolytic cleavage occurs (Binder et al., 1994). The
nuclease responsible for TfR mRNA cleavage remains to be identified.
1.2.1.3 RNAi Induced Endonucleolytic Cleavage
Small RNAs can trigger different types of gene silencing that are collectively referred to
as RNA silencing or RNA interference (RNAi). RNAs are processed into small ~21-26
nucleotides (nt) RNA molecules known as micro RNAs (miRNAs) and short interfering RNAs
(siRNAs). miRNAs are produced from transcripts that form stem-loop structures while siRNAs
are produced from long double-stranded (dsRNA) precursors [reviewed in (Valencia-Sanchez et
al., 2006)]. Once processed siRNAs and miRNAs are incorporated into the RNA-induced
silencing complex (RISC) which is the effector of RNAi (Gregory et al., 2004). A key
component of RISC is an Argonaute protein. The Argonaute protein family is diverse, with all
members containing a PAZ domain, involved in miRNA/siRNA binding, and a PIWI domain,
related to RNase H endonucleases (Lingel & Sattler, 2005). miRNA and siRNA serve to target
RISC to specific mRNAs based on sequence complementarity. RNAi can regulate both mRNA
translation and RNA degradation. The mechanism of RNAi-mediated regulation is typically
19
determined by base-pairing between the siRNA/miRNA and the mRNA, and the protein
composition of RISC.
One manner in which siRNAs control post-transcriptional gene expression is by directing
endonuclease cleavage of target mRNAs. This cleavage requires Ago2, the only Argonaute
capable of endonuclease cleavage in humans and Drosophila (Liu et al., 2004; Okamura et al.,
2004). Endonucleolytic cleavage by RISC is generally favored when perfect base-pairing
between the siRNA and the mRNA target occurs, although some mismatches can be tolerated
and still allow cleavage (Mallory et al., 2004; Yekta et al., 2004). The products of RNAi-
mediated cleavage are subsequently degraded by general mRNA decay factors (Valencia-
Sanchez et al., 2006). miRNAs do not usually exhibit full complementarity to their target
mRNAs, and induce mRNA degradation via an endonuclease-independent mechanism (discussed
below).
1.2.2 Elements that Recruit the Deadenylase Complex
The regulation of mRNA decay is often mediated by cis-acting sequence elements that
enhance deadenylation. These elements are bound by proteins that recruit the deadenylase
machinery and enhance degradation via poly(A) tail removal. This method of mRNA
destabilization is characteristic of the Drosophila proteins Smaug, Nanos and Bicaudal C; the
cold-shock-domain-containing protein UNR; and members of the PUF protein family, named for
its founders Drosophila Pumillio (Pum) and C. elegans FUSE-binding protein (FBP) (Chang et
al., 2004; Goldstrohm et al., 2006; Chicoine et al., 2007; Goldstrohm et al., 2007; Hook et al.,
2007; Kadyrova et al., 2007).
20
The mammalian protein UNR is a cold-shock-domain-containing RNA-binding protein
involved in the regulation of mRNA stability. UNR binds the c-fos major protein coding-region
determinant of instability (mCRD) and also poly(A) binding protein (PABP) (Chang et al.,
2004). The interaction between UNR and PABP is necessary for full mRNA destabilization
(Chang et al., 2004). UNR also interacts with Ccr4 and is proposed to recruit the Ccr4-Pop2-Not
deadenylase complex to initiate mRNA decay by poly(A) tail shortening (Chang et al., 2004).
Interestingly, blocking either translation initiation or elongation greatly impairs deadenylation
and mRNA decay, demonstrating that UNR-mediated degradation is coupled to translation
(Chang et al., 2004). UNR is regarded as a “landing/assembly” platform for the formation of a
deadenylation/decay complex on a mCRD-containing transcript (Chang et al., 2004). The
interaction between UNR and PABP renders associated mRNAs resistant to poly(A) shortening
(Grosset et al., 2000). Ribosome transit through the mCRD disrupts the UNR-PABP interaction,
leading to the recruitment of the Ccr4-Pop2-Not complex and deadenylation-mediated mRNA
decay (Grosset et al., 2000; Chang et al., 2004). This mechanism illustrates that mRNA
degradation can be tightly coupled to translation.
PUF proteins are characterized by the presence of eight consecutive PUF repeats that are
required for RNA binding [reviewed in (Wickens et al., 2002)]. Individual PUF proteins
regulate mRNAs by binding to sequence-specific control elements in the transcript 3’UTR. In S.
cerevisiae the PUF protein Mpt5 binds and regulates HO mRNA, which encodes a DNA
endonuclease required for mating-type switching (Tadauchi et al., 2001; Gerber et al., 2004).
Mpt5 regulates the degradation of HO mRNA by recruiting the Ccr4-Pop2-Not deadenylase
complex and initiating poly(A) tail removal (Goldstrohm et al., 2006; Goldstrohm et al., 2007).
HO mRNA is also regulated by Puf4 which binds to a distinct site in the HO 3’UTR (Hook et al.,
21
2007). Like Mpt5, Puf4 stimulates HO mRNA deadenylation by directly recruiting the Ccr4-
Pop2-Not deadenylase complex (Hook et al., 2007). By binding to HO mRNA simultaneously,
Puf4 and Mpt5 collaborate to regulate transcript decay through deadenylase recruitment.
In Drosophila, the PUF protein Pumillio (Pum) binds to specific elements in the Cyclin B
(CycB) mRNA (Kadyrova et al., 2007). The main function of Pum is to recruit the RNA-binding
protein Nanos (Nos), as regulation by Pum is absolutely dependent on its ability to bind Nos
(Kadyrova et al., 2007). Together Nos and Pum recruit the Ccr4-Pop2-Not deadenylase complex
to CycB mRNA, resulting in translational repression of the transcript via deadenylation
(Kadyrova et al., 2007). Similarly, the Drosophila RNA-binding protein Bicaudal C (Bic-C)
negatively regulates its own expression by binding specific sequences in the Bic-C 5’UTR
(Chicoine et al., 2007). Bic-C directly binds Not3/5, a component of the Ccr4-Pop2-Not
deadenylase, to promote mRNA decay via deadenylation (Chicoine et al., 2007).
1.2.3 miRNA Regulation of mRNA Decay
The upregulation of mRNA targets when the miRNA pathway is inhibited, demonstrates
that miRNAs can induce mRNA degradation (Behm-Ansmant et al., 2006; Giraldez et al., 2006;
Eulalio et al., 2007b). In animal cells, Argonaute proteins do not usually degrade target mRNAs
through endonucleolytic cleavage, except when the miRNA is fully complementary to the target.
Rather miRNAs induce mRNA decay via the general mRNA decay machinery. In Drosophila S2
cells, mRNA degradation by miRNAs is inhibited by depletion of the Ccr4 deadenylase or
decapping machinery (Behm-Ansmant et al., 2006). Similarly in zebrafish, miRNAs accelerate
the deadenylation of target mRNAs while mutation of the miRNA target site delayed mRNA
deadenylation (Giraldez et al., 2006). These data demonstrate that miRNAs can accelerate
mRNA degradation via deadenylation and subsequent decapping.
22
1.2.4 The Stabilizing and Destabilizing Effects of A+U-Rich Elements
A signal for rapid mRNA degradation in mammalian cells is the A+U-rich element
(ARE) found in the 3’UTR of some mRNAs encoding cytokines, proto-oncogenes and growth
factors [reviewed in (Wilusz et al., 2001b)]. A number of ARE-binding proteins have been
identified and shown to have either a positive or negative effect on mRNA stability (Wilusz et
al., 2001b). Tristetraprolin (TTP) binds to AREs through two CCCH-type zinc-finger domains
and induces their destabilization (Lai et al., 2000). Following TTP binding, transcripts are
deadenylated and subsequently degraded (Chen et al., 2001a). In contrast, binding of AREs by
Hu-antigen R (HuR), a member of the embryonic lethal abnormal vision (ELAV) protein family,
leads to mRNA stabilization (Peng et al., 1998). Although the mechanism of HuR-mediated
mRNA stabilization is poorly understood, in overexpressing cells HuR appears to protect the
mRNA body from degradation rather than slow the rate of deadenylation (Peng et al., 1998).
These data suggest that TTP and HuR have antagonistic effects on the stability of ARE-
containing mRNAs.
The mechanism by which TTP destabilizes ARE-containing mRNAs is not well
understood. TTP can target a tethered non-ARE reporter mRNA for rapid decay (Lykke-
Andersen & Wagner, 2005). In addition, TTP interacts with a number of mRNA decay enzymes
responsible for deadenylation, decapping, 5’-to-3’ decay and 3’-to-5’ decay (Lykke-Andersen &
Wagner, 2005). Currently TTP is hypothesized to deliver ARE-containing mRNAs to P-bodies.
TTP can localize with tethered mRNAs to P-bodies and this localization is strongly enhanced
when the mRNA decay machinery is impaired (Franks & Lykke-Andersen, 2007). In contrast,
depletion of endogenous TTP or overexpression of a dominant-negative mutant TTP protein,
23
impairs the localization of ARE-containing mRNAs to P-bodies (Franks & Lykke-Andersen,
2007).
Recent work suggests that microRNAs are also involved in the regulation of ARE-
mediated mRNA decay. Components of the RNAi machinery are required for the rapid decay of
ARE-containing mRNAs in Drosophila S2 cells and HeLa cells (Jing et al., 2005). In addition
human miR16, containing a sequence that is complementary to the ARE sequence, was shown to
be required for ARE-mediated mRNA decay (Jing et al., 2005). TTP interacts with the human
Argonaute protein eIF2C2, a component of the RISC effector complex (Jing et al., 2005). These
data argue that TTP may assist in the targeting of RISC to ARE mRNAs, and suggest that
cooperation of miRNA and ARE-binding proteins is required for efficient ARE mRNA decay
(Jing et al., 2005).
1.3 Post-Transcriptional Regulation by the Smg Protein Family
The Smg family is a class of proteins, conserved from yeast to humans, that regulate gene
expression post-transcriptionally (Figure 1-2) (Smibert et al., 1996; Dahanukar et al., 1999;
Smibert et al., 1999; Aviv et al., 2003; Baez & Boccaccio, 2005; Semotok et al., 2005; Rendl et
al., 2008). Smg proteins bind RNA through a conserved sterile alpha motif (SAM) domain that
interacts directly with stem-loop structures known as Smaug Recognition Elements (SREs)
(Smibert et al., 1996; Dahanukar et al., 1999; Smibert et al., 1999; Crucs et al., 2000; Aviv et al.,
2003; Green et al., 2003). In addition to similarities within their SAM domains, two other
regions are common to Drosophila Smg and many of its homologs (Aviv et al., 2003). The two
regions, designated Smg similarity regions 1 and 2 (SSR1 and SSR2), have been previously
identified in the Drosophila, mouse and human Smg proteins (Smibert et al., 1999).
24
Figure 1-2 The Smg Family of RNA-Binding Proteins
Cladogram representing overall sequence similarity and domain architecture of the Smg
homologs. Zif represents a CCHC zinc-finger domain. Species abbreviations: dm, Drosophila
melanogaster; ag, Anopheles gambiae; hs, Homo sapiens; mm, Mus musculus; ce,
Caenorhabditis elegans; ca, Candida albicans; sp, Schizosaccharomyces pombe; sc,
Saccharomyces cerevisiae. Figure adapted from Aviv et al. (2003).
25
SAM domains in general are best characterized to function as protein-protein interaction
domains [reviewed in (Qiao & Bowie, 2005)]. However the Smg SAM domain represents a
subclass of SAM domains with RNA binding abilities (Aviv et al., 2003; Green et al., 2003). The
structures of the Smg SAM domain in S. cerevisiae and Drosophila conform to the consensus
SAM architecture of five helices that are packed by a hydrophobic core (Aviv et al., 2003; Green
et al., 2003; Aviv et al., 2006; Oberstrass et al., 2006). The SAM domain of Smg family
members differ from other SAM domains in that they also carry a cluster of basic residues on
their surface which mediate RNA binding (Green et al., 2003; Aviv et al., 2006; Oberstrass et al.,
2006). The consensus for Smg RNA binding is defined as a nonspecific stem displaying a loop
up to 7 nucleotides (nt) long, CNGGN0-3 (Aviv et al., 2006). Structural analysis demonstrates
that the sequence specificity of SRE binding is limited to recognition of the G residue at position
3 of the loop, in combination with recognition of the SRE tertiary structure (Aviv et al., 2006;
Johnson & Donaldson, 2006; Oberstrass et al., 2006). All Smg proteins are predicted to bind
RNA with the same specificity and regulate target mRNAs post-transcriptionally. This has been
confirmed by studies of Smg in Drosophila, mammals and yeast.
1.3.1 Drosophila Smg
The Smg-mediated post-transcriptional regulation of two target mRNAs, nanos (nos) and
heat shock protein 83 (Hsp83), has been well characterized in Drosophila. In the following
sections I will discuss the importance of nos and Hsp83 regulation to Drosophila development,
and more specifically the mechanisms of Smg-mediated regulation.
1.3.1.1 Post-Transcriptional Regulation of nos mRNA
Posterior development of the Drosophila embryo requires Nos, a translational regulator
of hunchback (hb) and bicoid (bcd) mRNA. Embryos lacking Nos activity exhibit abdominal
26
defects (Wang & Lehmann, 1991), and ectopic expression of Nos at the anterior leads to lethal
head defects (Wharton & Struhl, 1991). Posterior expression of Nos is the result of local
translation of nos mRNA which has been localized to the embryo posterior (Wang & Lehmann,
1991). However localization of nos transcripts is an inefficient process with only 4% of nos
mRNA estimated to be posteriorly localized while the remainder is dispersed throughout the bulk
cytoplasm of the embryo (Bergsten & Gavis, 1999). To avoid misexpression of Nos, unlocalized
nos mRNA is translationally repressed. Both localization and translational regulation of nos are
controlled by regions of the nos 3’UTR.
Posterior localization of nos mRNA is mediated by a complex signal of several partially
redundant elements in the 3’UTR (Gavis et al., 1996a). Rumpelstiltskin (Rump), a heterogeneous
nuclear ribonucleoprotein (hnRNP) M homolog, recognizes two CGUU motifs within the nos
localization element (Jain & Gavis, 2008). Mutation of either of these motifs disrupts Rump
binding and nos mRNA localization (Jain & Gavis, 2008). The fact that sequences required for
Rump binding are also required for posterior localization supports a direct role for Rump in
mediating nos localization through an interaction with the nos 3’UTR.
A region of the nos 3’UTR, known as the translational control element (TCE) is required
for translational repression of nos in the bulk cytoplasm (Dahanukar & Wharton, 1996).
Repression of nos translation is mediated by cis-acting stem-loop structures within the TCE.
Glorund (Glo), a member of the hnRNP F/H family, binds to a double-stranded U/A-rich motif
within the TCE to repress unlocalized nos translation in the Drosophila oocyte (Kalifa et al.,
2006). Mutations within this motif that disrupt Glo binding correlate to a loss of nos repression
(Kalifa et al., 2006). The TCE also contains two functionally redundant RNA stem-loop
elements, that share the common loop sequence of CUGGC (Dahanukar & Wharton, 1996;
27
Gavis et al., 1996a; Gavis et al., 1996b; Smibert et al., 1996). These stem-loops are recognized
by the RNA-binding protein Smg, which functions as a translational repressor in the Drosophila
embryo (Dahanukar & Wharton, 1996; Smibert et al., 1996; Smibert et al., 1999). The
mechanism of Smg-mediated repression is discussed below.
Posteriorly localized nos mRNA must be translated to direct Drosophila development.
Since both Glo and Smg are ubiquitously expressed throughout the oocyte and embryo
respectively, a mechanism must be in place to activate nos translation at the posterior
(Dahanukar & Wharton, 1996; Smibert et al., 1996; Smibert et al., 1999; Kalifa et al., 2006). The
mechanism of nos translational activation at the oocyte posterior remains to be elucidated. In the
embryo, repression of nos translation at the posterior is relieved by Oskar (Osk) (Gavis &
Lehmann, 1994; Wang et al., 1994). While the molecular mechanisms that underlie the role of
Osk in nos translational activation are unknown, one experiment suggests that Osk prevents Smg
binding to nos mRNA, which allows localized transcripts to be translated at the posterior
(Zaessinger et al., 2006).
1.3.1.2 Smg-Mediated Regulation of nos mRNA
Smg binds to SRE stem-loop structures within the nos TCE to regulate the translation of
unlocalized nos mRNA in the Drosophila embryo (Dahanukar & Wharton, 1996; Smibert et al.,
1996; Smibert et al., 1999). Mutations to the CUGGC loop sequence at positions one and three
disrupt Smg binding in vitro and lead to developmental head defects in vivo, consistent with the
disruption of unlocalized nos repression (Dahanukar & Wharton, 1996; Smibert et al., 1996).
Mutations to the SRE stem also disrupt nos repression however compensatory mutations that
restore formation of a stem rescue nos repression, indicating that the formation but not sequence
of the SRE stem is required for Smg binding (Crucs et al., 2000).
28
Smg binds unlocalized nos mRNA and recruits the eIF4E-binding protein Cup to repress
translation in the bulk cytoplasm of the embryo (Nelson et al., 2004). Cup is an eIF4E-binding
protein capable of binding the translation initiation factor through a consensus eIF4E-binding
motif (YXXXXLΦ, where Φ denotes a hydrophobic residue) (Nelson et al., 2004). Using
biochemical approaches, Cup was identified as a Smg-binding protein that mediates an indirect
interaction between Smg and eIF4E (Nelson et al., 2004). Formation of the Smg/Cup/eIF4E
complex on nos mRNA antagonizes binding of the translation initiation factor eIF4G (Nelson et
al., 2004). By recruiting Cup, Smg disrupts formation of the translation initiation complex and
prevents misexpression of unlocalized nos mRNA (Figure 1-3). In addition to repression of
translation initiation by Smg, the observation that translationally repressed nos transcripts remain
polysome associated suggests that additional mechanisms may contribute to the translational
repression of unlocalized nos (Clark et al., 2000).
Unlocalized nos mRNAs are translationally repressed in the bulk cytoplasm and
subsequently degraded in the first two to three hours of embryonic development (Bashirullah et
al., 1999). While Smg has a well documented role in regulating transcript stability (see below),
conflicting results about its specific role in regulating nos mRNA stability have been reported
(Semotok et al., 2005; Zaessinger et al., 2006).
29
Figure 1-3 Model for Smg-Mediated Repression of nos mRNA Translation
(A) Shown is a schematic representation of translation initiation. The mRNA 5’ m7G-cap is first
recognized by the cap-binding protein eIF4E, which recruits the translation initiation factor
eIF4G to the transcript though the eIF4E-binding motif (YXXXXLΦ) present in eIF4G. Binding
of eIF4G to eIF4E facilitates formation of the translation intitation complex on the transcript and
recognition of the mRNA by the 40S ribosomal subunit. (B) Smg binds to SREs and represses
translation of unlocalized nos mRNA in the Drosophila embryo. Upon binding to nos mRNA,
Smg recruits the eIF4E-binding protein Cup, which mediates an indirect interaction between
Smg and eIF4E. Formation of the Smg/Cup/eIF4E complex blocks binding of eIF4G and
represses mRNA translation, preventing misexpression of unlocalized nos mRNA.
30
1.3.1.3 Post-Transcriptional Regulation of Hsp83 mRNA
In Drosophila, early embryo development is programmed by maternally provided
mRNAs. Maternal Hsp83 transcripts are initially distributed uniformly within the maturing
oocyte (Ding et al., 1993). After fertilization, Hsp83 mRNA localizes to the posterior pole plasm
via a degradation-protection mechanism (Ding et al., 1993; Bashirullah et al., 1999; Bashirullah
et al., 2001). Posterior-localized transcripts are protected while Hsp83 mRNA in the bulk
cytoplasm is rapidly degraded (Bashirullah et al., 1999; Bashirullah et al., 2001). Cis-acting
elements within Hsp83 mRNA are required for both degradation and protection of the transcript
(Bashirullah et al., 1999; Semotok et al., 2008).
1.3.1.4 Smg-Mediated Regulation of Hsp83 mRNA
Smg binds specifically to multiple SREs in Hsp83 mRNA and induces transcript
degradation in the bulk cytoplasm of the embryo (Semotok et al., 2005; Semotok et al., 2008). At
the posterior pole Hsp83 mRNA escapes Smg-mediated decay through an unknown mechanism.
In smg mutants, Hsp83 mRNA is stabilized throughout the embryo and is not preferentially
localized to the posterior, supporting a role for Smg in the degradation and localization of Hsp83
(Semotok et al., 2005). Mutation of SREs within the Hsp83 mRNA or mutation of Smg to block
RNA binding, also stabilize the transcript (Semotok et al., 2008). Upon binding to Hsp83
mRNA, Smg physically recruits the Ccr4-Pop2-Not deadenylase complex and initiates transcript
decay via poly(A) tail shortening (Semotok et al., 2005). In ccr4 mutants Hsp83 mRNA is
stabilized, and in smg mutants Hsp83 persists as a stable transcript with a long poly(A) tail.
When targeted to a heterologous mRNA, Smg is sufficient to induce transcript decay via
deadenylation (Semotok et al., 2005). Although Smg can also function as a translational
repressor, Smg does not repress Hsp83 mRNA translation (Semotok et al., 2005). Smg
31
physically interacts with the Ccr4-Pop2-Not deadenylase and this complex appears to be distinct
from the repressor complex it forms with Cup (Semotok et al., 2005). Therefore Smg appears to
employ two molecularly distinct mechanisms to regulate mRNAs post-transcriptionally. In the
case of mRNA decay, Smg triggers transcript destabilization through recruitment of the Ccr4-
Pop2-Not deadenylase (Figure 1-4).
1.3.1.5 Smg is a Major Regulator of mRNA Stability
Early Drosophila development is controlled by RNAs and proteins that are maternally
loaded into the oocyte. Upon egg activation a subset of maternal mRNAs are targeted for
degradation [reviewed in (Tadros & Lipshitz, 2005)]. Smg is a major regulator of mRNA
stability, required for the elimination of two-thirds of the unstable maternal mRNAs during
embryo development (Tadros et al., 2007). Transcripts regulated by Smg are enriched in gene
ontology annotations for cell cycle functions, suggesting a relationship between the increased
stability of these transcripts and the cell cycle defects observed in smg mutant embryos
(Dahanukar et al., 1999; Tadros et al., 2007).
1.3.2 Mammalian Smg
Two Smg homologs are present in the mammalian genome (Smibert et al., 1999; Aviv et
al., 2003). Smg 1 located in human and mouse chromosome 14, shares all three regions of
conservation with Drosophila Smg: SSR1, SSR2 and SAM domain (Aviv et al., 2003; Baez &
Boccaccio, 2005). When transfected in BHK cells, Smg 1 represses the translation of SRE-
containing mRNAs without affecting their stability (Baez & Boccaccio, 2005). The mechanism
of repression by Smg 1 awaits further investigation however it is assumed that, like other Smg
proteins, Smg 1 can regulate mRNAs by binding to SREs via the SAM domain.
32
Figure 1-4 Model for Smg-Mediated Decay of Hsp83 mRNA
In the Drosophila embryo, Smg binds to SREs in Hsp83 mRNA and induces transcript
degradation. Upon binding to Hsp83 mRNA, Smg physically recruits the Ccr4-Pop2-Not
deadenylase complex and initiates transcript deay via poly(A) tail shortening.
33
When expressed in fibroblasts, Smg 1 forms cytoplasmic foci that contain polyadenylated
RNA and are in equilibrium with polysomes (Baez & Boccaccio, 2005). These Smg 1 foci lack
GW182, a P-body marker, but contain several proteins that are found in stress granules. Stress
granules are induced by environmental cues and contain nontranslating mRNAs, a subset of
translation initiation factors, 40S ribosomal subunits, and a number of RNA-binding proteins
(Anderson & Kedersha, 2006). Whether the human Smg 1 foci represent a novel function for this
protein or are in some way related to the function of other Smg family members awaits further
investigation.
1.3.3 Vts1, the Smg Homolog in Saccharomyces cerevisiae
In S. cerevisiae, VTS1 is a nonessential gene whose protein product has been implicated
in vesicular transport based on its ability to suppress the growth and vacuolar-transport defects in
vti1-2 cells, which carry a temperature-sensitive allele of the Q-SNARE VTI1 (Dilcher et al.,
2001). Vts1 is a member of the Smg protein family based on the presence of a conserved Smg
SSR1 region and SAM domain (Aviv et al., 2003). The Vts1 SAM domain can bind RNA with
the same specificity for SRE stem-loop structures as other Smg family members (Aviv et al.,
2003). A point mutation within the SAM domain, known to block Smg RNA binding, disrupts
Vts1 RNA binding demonstrating a direct interaction between the Vts1 SAM domain and RNA
(Aviv et al., 2003).
Vts1 can regulate the expression of a reporter mRNA harboring SREs in the 3’UTR of
the transcript (Aviv et al., 2003). Reporter expression is regulated at the level of mRNA stability
and requires Vts1 and functional SREs. Vts1-mediated mRNA degradation appears to require the
Ccr4-Pop2-Not deadenylase complex since reporter steady-state levels are increased in ccr4Δ
34
cells (Aviv et al., 2003). Given that both Drosophila Smg and Vts1 require Ccr4 to degrade
mRNAs, a conserved mechanism may be employed by all Smg proteins to regulate mRNA
stability.
To gain a better understanding of Vts1 function, RNA targets that seem to be regulated
by Vts1 have been identified by several methods. In two studies, potential endogenous targets of
Vts1 were identified by immunoprecipitating Vts1 and identifying copurifying mRNAs via
microarray (Aviv et al., 2006; Hogan et al., 2008). These targets were largely nuclear-encoded
mRNAs and enriched for SRE sites. To confirm the microarray results, selected transcripts co-
purified with Vts1 but not a version in which the SAM domain was mutated to disrupt RNA
binding (Aviv et al., 2006). The steady state levels of selected transcripts was elevated in vts1Δ
cells compared to wild-type, further confirming that some are regulated by Vts1 (Aviv et al.,
2006). Some of the identified transcripts are required for sporulation and germination, cellular
functions that are enhanced in the vts1Δ strain (Deutschbauer et al., 2002). Interestingly, a
significant proportion of the targets encode genes involved in carbohydrate transport (Hogan et
al., 2008). A third approach to identify Vts1 target transcripts employed microarray gene
profiling to identify genes strongly upregulated in vts1Δ cells compared to wild-type as potential
targets of Vts1 regulation (Oberstrass et al., 2006). Characterization of these genes by
bioinformatics showed that many contain one to several copies of the SRE. As validation of the
microarray data, the expression of four predicted targets was analyzed by Northern blot and
found to be expressed at higher levels in vts1Δ cells compared to wild-type (Oberstrass et al.,
2006). While potential endogenous targets of Vts1 have been identified, the mechanism that
underlies Vts1-mediated regulation of these transcripts has not been explored.
35
1.4 Thesis Rationale
At the inception of my thesis the role of Vts1 in mRNA degradation was relatively
uncharacterized. Since Vts1 was identified as a member of the Smg protein family capable of
specifically binding and regulating mRNA (Aviv et al., 2003; Green et al., 2003; Aviv et al.,
2006; Oberstrass et al., 2006) the goal of my project was to characterize the molecular
mechanism of Vts1-mediated mRNA decay. To this end, I performed experiments using RNA
reporters that recapitulate Vts1-mediated degradation in vivo and endogenous mRNA transcripts
to elucidate the pathway of transcript decay in S. cerevisiae. I demonstrate that Vts1 target
mRNAs are degraded by the major decay pathway of deadenylation-dependent decapping
followed by 5’-to-3’ exonucleolytic digestion. Together with data illustrating an interaction
between Vts1 and a component of the deadenylase complex Pop2, I propose that Vts1 recruits
the Ccr4-Pop2-Not deadenylase to induce degradation of target transcripts.
I also identified the eIF4E-binding protein Eap1 as a novel component of the RNA
turnover machinery and characterized its role in Vts1-mediated mRNA decay. Using both RNA
reporters and endogenous transcripts I demonstrate that Vts1 target mRNAs accumulate as
deadenylated capped species in eap1Δ cells, suggesting that Eap1 stimulates mRNA decapping. I
also provide data that Eap1 mediates an indirect interaction between Vts1 and eIF4E. I show that
the interaction between Vts1 and eIF4E requires the Eap1 eIF4E-binding motif. I propose that
the Vts1/Eap1/eIF4E complex increases the susceptibility of target mRNAs to decapping.
Together these data illustrate that Vts1 employs multiple mechanisms to induce rapid
degradation of target mRNAs.
36
Chapter 2
S. cerevisiae Vts1 Induces Deadenylation-Dependent Transcript
Degradation and Interacts with the Ccr4-Pop2-Not Deadenylase
Complex
The work described in this chapter was published in:
Rendl, L.M, Bieman, M.A., and Smibert, C.A. (2008). S. cerevisiae Vts1p induces
deadenylation-dependent transcript degradation and interacts with the Ccr4p-Pop2p-Not
deadenylase complex. RNA 14: 1328-1336.
Contribution of work: I performed the majority of the experimental work presented in this
chapter.
M. Bieman performed the coimmunoprecipitation of Vts1 and Pop2 (Figure 2-3).
37
Chapter 2
S. cerevisiae Vts1 Induces Deadenylation-Dependent Transcript Degradation
and Interacts with the Ccr4-Pop2-Not Deadenylase Complex
2.1 Abstract
The Smaug family of sequence-specific RNA-binding proteins regulates mRNA
translation and degradation by binding to consensus stem/loop structures in target mRNAs. Vts1
is a member of the Smaug protein family that regulates the stability of target transcripts in S.
cerevisiae. Here we focus on the mechanism of Vts1-mediated mRNA decay. Using RNA
reporters that recapitulate Vts1-mediated decay in vivo we demonstrate that Vts1 stimulates
mRNA degradation through deadenylation mediated by the Ccr4-Pop2-Not deadenylase
complex. We also show that Vts1 interacts with the Ccr4-Pop2-Not complex suggesting that
Vts1 recruits the Ccr4-Pop2-Not deadenylase complex to target mRNAs resulting in transcript
decay. Following deadenylation Vts1 target transcripts are decapped and subsequently degraded
by the 5’-to-3’ exonuclease Xrn1. Decapping and 5’-to-3’ decay is thought to occur in foci
known as P-bodies, and we provide evidence that Vts1 function may involve P-bodies. Taken
together with previous work, these data suggest that Smaug family members employ a conserved
mechanism to induce transcript degradation that involves recruitment of the Ccr4-Pop2-Not
deadenylase to target mRNAs.
38
2.2 Introduction
The identification of several conserved mRNA degradation pathways in eukaryotes
highlights the importance of mRNA turnover as a means of controlling gene expression. In
Saccharomyces cerevisiae the major mRNA decay pathway initiates with deadenylation by the
Ccr4-Pop2-Not deadenylase complex (Decker & Parker, 1993; Hsu & Stevens, 1993; Muhlrad et
al., 1995). Following poly(A) removal the transcript is decapped and then degraded 5’-to-3’ by
the exonuclease Xrn1 (Hsu & Stevens, 1993; Muhlrad et al., 1994, 1995; Caponigro & Parker,
1996b). In addition to the general pathway of mRNA decay, sequence-specific RNA-binding
proteins add an additional level of complexity to the regulation of transcript stability (Garneau et
al., 2007). In general these proteins bind target sequences in a given mRNA and can affect
transcript stability by interacting with components of the mRNA decay machinery.
The Smaug (Smg) family is a class of proteins, conserved from yeast to humans, that
regulate gene expression post-transcriptionally through their ability to bind directly to target
mRNAs (Smibert et al., 1996; Dahanukar et al., 1999; Smibert et al., 1999; Aviv et al., 2003;
Baez & Boccaccio, 2005; Semotok et al., 2005). Family members bind RNA with similar
specificity through a conserved sterile alpha motif (SAM) domain that is able to interact with
stem-loop structures termed Smg recognition elements (SREs) (Smibert et al., 1996; Dahanukar
et al., 1999; Smibert et al., 1999; Crucs et al., 2000; Aviv et al., 2003; Green et al., 2003).
Drosophila Smg, the founding member of this family, regulates mRNA translation and
destabilization through separate mechanisms. Smg distinguishes between these roles by
recruiting specific trans-acting factors that aid in transcript regulation. Smg represses mRNA
translation by recruiting the eIF4E-binding protein Cup to target transcripts (Nelson et al., 2004).
In contrast, Smg triggers transcript destabilization by recruiting the Ccr4-Pop2-Not deadenylase
39
complex to target mRNAs (Semotok et al., 2005). We have proposed a similar mechanism of
mRNA destabilization by Vts1, the Smg homolog in S. cerevisiae (Aviv et al., 2003). Vts1 can
regulate the expression of a reporter mRNA harboring SREs in the 3’UTR of the transcript. This
work also showed that reporter expression is regulated at the level of mRNA stability and that
both Vts1 and functional SREs are required for this effect. Vts1-mediated mRNA degradation
appears to require Ccr4-Pop2-Not deadenylase complex since reporter mRNA steady-state levels
are increased in cells lacking Ccr4 (Aviv et al., 2003), which is the catalytic subunit of this
complex (Chen et al., 2002; Tucker et al., 2002; Parker & Song, 2004). While potential
endogenous targets of Vts1 have been identified by immunoprecipitating Vts1 and identifying
co-purifying mRNAs via microarray (Aviv et al., 2006), the mechanism that underlies Vts1-
mediated regulation of these transcripts has not been explored.
Here we show that Vts1 induces mRNA decay via the major mRNA decay pathway of
deadenylation-dependent decapping and subsequent 5’-to-3’ decay. Using reporter mRNAs we
show that Vts1 and Ccr4 are required for the rapid decay and deadenylation of SRE-containing
transcripts in vivo and that Vts1 associates with the Ccr4-Pop2-Not deadenylase complex,
suggesting that Vts1-mediated decay results from recruitment of this deadenylase to target
mRNAs. We demonstrate that deadenylation is the initial step in Vts1-mediated mRNA decay
and that following poly(A) tail removal transcripts are decapped and then degraded in a 5’-to-3’
direction by the exonuclease Xrn1. Decapping and 5’-to-3’ decay of transcripts is thought to
occur in cytoplasmic foci known as P-bodies (Sheth & Parker, 2003), and we provide additional
evidence that suggests that Vts1 function might involve P-bodies. Together, these data
specifically outline the molecular mechanism Vts1 employs to degrade mRNAs. They also
provide evidence suggesting that Smg family members employ a conserved mechanism to induce
40
transcript decay, which involves recruitment of the Ccr4-Pop2-Not deadenylase to target
mRNAs.
2.3 Materials and Methods
2.3.1 Yeast Strains and Media
Yeast strains used in this study were derivatives of the wild-type BY4741 (Brachmann et
al., 1998). All deletion strains are as described by Winzeler et al. (1999), with the exception of
the ccr4Δ strain, which is described by Woolstencroft et al. (2006). The TetO7-DCP2 strain is
described by Mnaimneh et al. (2004). The strains were transformed by standard techniques and
plasmids were maintained by growth in selective media.
2.3.2 mRNA Analysis
Transcriptional pulse-chase experiments employed GFP-SRE+ and GFP-SRE- reporters
described by Aviv et al. (2003). Both reporters express the GFP open reading frame in
p413GAL1 with approximately 160 nt of the CYC1 3’UTR containing the 3xSRE+ or 3xSRE–
sequence, which is described in Smibert et al. (1996). For these experiments cells were grown at
30oC to mid-log phase in selective medium containing 2% raffinose and cooled to 20oC, with the
exception of Figure 2-6 in which transcriptional shut-off was performed at 30oC. An accurate
measure of reporter mRNA stability at 30oC was not possible due to the short half-life of the
GFP-SRE+ reporter in wild-type cells. Cooling to 20oC slowed the decay of GFP-SRE+ mRNA to
allow for measurement of reporter mRNA half-lives. GFP reporter transcription was initiated by
the addition of galactose to a final concentration of 2%. After 16 min transcription was repressed
by the addition of glucose to a final concentration of 4%. Total RNA was isolated by glass bead
lysis in LET buffer (100 mM LiCl, 20 mM EDTA, 25 mM Tris-HCl [pH 8.0]) and LET-
equilibrated phenol at the indicated time points and analyzed by Northern blot. SCR1 or ACT1
41
RNA was used for normalization of reporter transcript levels as indicated. Where indicated,
thiolutin (Carbosynth Limited) was added to cultures to a final concentration of 20 µg/mL at
time 0 of the time course. Where indicated, TetO7-DCP2 cells were treated with 10 µg/mL
doxycycline for 16 hours before RNA reporter transcription was induced with galactose. RNase
H cleavage assays were performed as described by Decker et al. (1993), using an oligonucleotide
that hybridized ~330 nt upstream of the GFP reporter poly(A) site. To control for differences in
RNA concentration in these experiments an amount of each RNA sample was analyzed via a
standard Northern blot and probed for SCR1 RNA. Northern blots were exposed to
PhosphorImager screens and analyzed with ImageQuant Software with the exception of TetO7-
DCP2 in Figure 2-5 which was exposed to X-ray film. The GFP-YIR016W reporter was under
the control of the GAL1 promoter and expressed a GFP N-terminal fusion to the YIR016W ORF
plus 597 nt of YIR016W downstream sequence.
2.3.3 Immunoprecipitations
The plasmid expressing Vts1-Flag was generated in pRS316 with 574 bases of genomic
sequence upstream and 396 nt downstream of the VTS1 ORF. Vts1-Flag also harbors a VSV
epitope which was used for protein detection by Western blot. The Pop2-HA plasmid was
created in pRS313 with a C-terminal 3X HA tag and 549 nt of genomic sequence upstream and
530 nt downstream of the POP2 ORF. Cells were harvested and lysed in KHT buffer (150 mM
KCl, 30 mM Hepes pH 7.4, 0.1% Tween) by glass bead lysis and clarified at 15,000 rpm for 10
min. Anti-Flag M2 affinity gel (Sigma) was added to the supernatant and bound for 3 h at 4oC.
Beads were washed four times at 4oC with KHT buffer and then twice in 100 mM KCl, 100 mM
Hepes pH 7.4 at room temperature for 10 min. Vts1-VSV-Flag and associated proteins were
eluted with Flag-peptide (Sigma) for 10 min at room temperature. Where indicated 0.35 µg/µL
42
RNase A (Fermentas) was added during 3 h incubation at 4oC. VSV antibodies (Bethyl
Laboratories) and HA antibodies (Abcam) were used to detect VSV-tagged and HA-tagged
proteins, respectively.
2.3.4 Confocal Microscopy
The Vts1-mRFP plasmid was generated by C-terminally tagging Vts1 with monomeric
RFP in pRS316 with 574 nt of genomic sequence upstream and 396 nt downstream of the VTS1
ORF. The Dhh1-GFP plasmid was generated by C-terminally tagging Dhh1 with GFP in pRS315
with 517 nt of genomic sequence upstream and 430 nt downstream of the DHH1 ORF. Cells
expressing fluorescent proteins were grown to mid-log phase in selective media and visualized
with a Zeiss LSM 510 confocal microscope using the 100X objective. Images shown are of a
single focal plane and were generated with Zeiss LSM browser software.
2.4 Results
2.4.1 Ccr4 is Required for Vts1-Mediated mRNA Decay
We previously proposed that Vts1 regulates mRNA stability through a mechanism that
requires the Ccr4-Pop2-Not deadenylase complex (Aviv et al., 2003). This model was based on
experiments that showed that the steady-state levels of a Vts1-regulated reporter mRNA increase
in a ccr4Δ strain. To further test this model we first compared the stability of a reporter mRNA
in wild-type cells to cells carrying a deletion in either the VTS1 gene or CCR4 gene, which
encodes the catalytic subunit of the deadenylase (Chen et al., 2002; Tucker et al., 2002; Parker &
Song, 2004). This reporter, which has been previously shown to recapitulate Vts1-mediated
decay in vivo (Aviv et al., 2003), encodes green fluorescent protein (GFP) under the control of a
galactose-inducible promoter and has three SREs in its 3’UTR (GFP-SRE+). A similar reporter
43
bearing SREs in which the loop sequences are mutated to block Vts1 binding (GFP-SRE-) serves
as a control.
To measure the stability of the reporter mRNA we used a transcriptional pulse-chase
approach where reporter gene transcription was induced by the treatment of cells with galactose
for 16 min and then inhibited by the addition of glucose. The level of the GFP reporter mRNA
was then measured at 2-min intervals by Northern blotting with the zero time point
corresponding to the glucose addition. As we have previously reported the GFP-SRE+ mRNA is
rapidly degraded (half-life of ~ 4 min) in a manner that is dependent on both Vts1 and wild-type
SREs (Figure 2-1; (Aviv et al., 2003)). Here we show that rapid degradation of GFP-SRE+
mRNA requires Ccr4 as it is significantly stabilized in the ccr4Δ strain. These data confirm our
initial hypothesis that Ccr4 is required for Vts1-mediated mRNA degradation.
2.4.2 Vts1 Target mRNAs Undergo Ccr4-Dependent Deadenylation
To determine if degradation of the GFP-SRE+ transcript involved deadenylation by Ccr4,
we analyzed the poly(A) tail length of each GFP reporter in wild-type, vts1Δ and ccr4Δ cells. We
used an RNase H cleavage method to measure the poly(A) tail length of the GFP reporter
mRNAs over an 8-min transcriptional pulse-chase time course. A specific antisense
oligonucleotide which hybridizes to the GFP open reading frame was added to total RNA from
each time point and cleaved with RNase H to produce a short 3’-end fragment of the GFP
reporter mRNA. The poly(A) tail length of this fragment was measured by performing high
resolution polyacrylamide gel electrophoresis and Northern blotting. In one sample reporter
specific oligonucleotide and oligo(dT) was included and, as such, treatment with RNase H
produced a 3’-end fragment without a poly(A) tail serving as a marker for the deadenylated 3’-
end fragment.
44
Figure 2-1 Vts1 Stimulates the Decay of Target mRNAs
GFP reporter transcription was induced with galactose and then shut off with glucose and GFP
mRNA levels were assayed at the times indicated after transcriptional shutoff by Northern blot.
Levels of GFP mRNA were measured in wild-type (WT), vts1Δ, and ccr4Δ cells. SCR1 RNA
serves as a loading control. Half-lives (T1/2) for each strain are as indicated. The results of three
independent experiments were quantitated and normalized using the levels of SCR1 RNA and
graphed with error bars representing standard deviation.
45
In wild-type cells, at time zero there were two distinct populations of the GFP SRE+
mRNA with one having poly(A) tail lengths ranging from 30 to 50 nucleotides (nt) and another
population having very short poly(A) tails (Figure 2-2). This latter population presumably
represents transcripts that accumulated and were deadenylated during the 16 min induction with
galactose, suggesting rapid deadenylation of the transcript. If the GFP-SRE+ mRNA is subject to
rapid deadenylation followed by decay, this would predict that during a transcriptional pulse-
chase experiment the longer poly(A) tail species would be converted to the shorter tail species,
which at subsequent time points would disappear. However, the short half-life of the GFP-SRE+
transcript might make it difficult to observe these intermediates. Nonetheless, soon after the
addition of glucose we reproducibly detect a decrease in amount of the longer poly(A) tail
species and an increase in the amount of the shorter tail population (compare the distribution of
the poly(A) tail lengths of transcripts at time equals zero to the 2-min time point) while at later
time points the shorter tail population disappears.
In contrast, the majority of the GFP-SRE+ mRNA in vts1Δ cells and ccr4Δ cells and the
GFP-SRE- mRNA in wild-type cells retained a long poly(A) tail distribution of 30 to 50 nt that
exhibited little or no accumulation of short poly(A) tail transcripts during the 8-min time course
(Figure 2-2). Taken together these data suggest that recruitment of Vts1 to GFP mRNA via SREs
stimulates rapid Ccr4-dependent deadenylation. In addition, they demonstrate a strong
correlation between defects in deadenylation (observed in either the vts1Δ or ccr4Δ strains or
when the SREs are mutated) and stabilization of GFP-SRE+ mRNA, indicating that
deadenylation plays a key role in Vts1-mediated transcript decay. Additional data described
below lend further support to the central role of deadenylation in Vts1-mediated transcript decay.
46
Figure 2-2 Vts1 Induces the Deadenylation of Target mRNAs
GFP reporter transcription was induced with galactose and then shut off with glucose and an
RNase H cleavage assay was used to measure the poly(A) tail length of GFP reporter mRNAs in
wild-type (WT), vts1Δ, and ccr4Δ strains at the times indicated after transcriptional shutoff.
Where indicated oligo (dT) was added to remove the poly(A) tail by RNase H treatment,
providing a marker for deadenylated mRNA (dT). The distribution of poly(A) tail lengths at the
indicated times after transcriptional shutoff are displayed on the graphs. SCR1 RNA serves as a
loading control.
47
2.4.3 Vts1 Interacts with the Ccr4-Pop2-Not Deadenylase Complex
Our previous work suggests that Drosophila Smg induces mRNA degradation by
physically recruiting the Ccr4-Pop2-Not deadenylase complex to target transcripts (Semotok et
al., 2005). Given the requirement of Ccr4 for Vts1-mediated mRNA decay in S. cerevisiae, we
sought to determine whether Vts1 physically interacts with a component of the Ccr4-Pop2-Not
deadenylase complex in vivo, Pop2. We performed coimmunoprecipitation experiments using
whole-cell lysates from cells expressing Vts1 tagged with a Flag epitope and Pop2 tagged with
an HA epitope. Lysates from cells expressing only one of the epitope tagged proteins served as
negative controls. Vts1-Flag was immunoprecipitated using anti-Flag resin, and the
immunoprecipitates were analyzed by Western blot. Pop2-HA was present in the anti-Flag
immunoprecipitate when the Vts1-Flag protein was present (Figure 2-3). As a control, Pop2-HA
was not immunoprecipitated from an extract lacking Vts1-Flag. The co-immunoprecipitation of
Vts1 with Pop2 was RNA-independent, as it was observed in the presence of RNase A and when
Vts1 harbored an amino acid change (A498Q) that blocks its ability to bind RNA (Vts1RBD-)
(Aviv et al., 2003). This suggests that Vts1 physically associates with a component of the Ccr4-
Pop2-Not deadenylase complex and supports a model in which Vts1 recruits the deadenylase to a
target transcript to initiate deadenylation-dependent mRNA degradation.
2.4.4 Vts1-Mediated Decay Requires mRNA Decapping and 5’-to-3’ Degradation
In yeast mRNA deadenylation can trigger decapping and subsequent 5’-to-3’
exonucleolytic decay (Caponigro & Parker, 1996b). Our data suggest that Vts1 mediates
transcript degradation by recruiting the Ccr4-Pop2-Not deadenylase complex to initiate mRNA
deadenylation. To determine if Vts1 target transcripts are decapped and degraded 5’-to-3’, we
48
Figure 2-3 Vts1 Interacts with Pop2
Vts1-Flag and Pop2-HA are expressed in yeast cells individually or in combination as indicated,
and crude extracts were subjected to immunoprecipitation using an anti-Flag resin. Starting crude
extracts (input) and the resulting immunoprecipitates (IP: αFLAG) were analyzed by Western
blot. Input samples represent 5% for Vts1 blots and 0.5% for Pop2 blots of material used in the
immunoprecipitations. The addition of RNase A or use of the Vts1 RNA-binding mutant
(Vts1RBD-) is as indicated.
49
measured the stability of our GFP reporters in strains defective for decapping or 5’-to-3’ decay.
To test whether Vts1 target transcripts are decapped, we examined the stability of the GFP
reporters in a strain deficient for DCP2, the catalytic component of the decapping enzyme
(Steiger et al., 2003). As DCP2 is essential in our strain background, we employed a strain
expressing DCP2 under a tetracycline (tet)-regulatable promoter (TetO7-DCP2) (Mnaimneh et
al., 2004). With this promoter-shutoff system, repression of DCP2 transcription is achieved by
the addition of doxycycline (DOX), a tetracycline analog, to the growth medium. In the absence
of DOX, TetO7-DCP2 is expressed and the GFP-SRE+ transcript decays at a rate similar to wild-
type (Figure 2-4). When TetO7-DCP2 transcription is repressed by the addition of DOX, the
stability of GFP-SRE+ is significantly increased, indicating that decapping is required for rapid
turnover of the GFP-SRE+ mRNA. Also, consistent with the role of decapping in Vts1-mediated
decay, we find that GFP-SRE+ mRNA is stabilized in a strain deleted for the decapping activator
PAT1 (Hatfield et al., 1996; Zhang et al., 1999; Bonnerot et al., 2000; Bouveret et al., 2000;
Tharun et al., 2000). Interestingly, degradation of the GFP-SRE+ mRNA does not require another
decapping activator Dhh1 (Figure 2-4) suggesting that Dhh1 is not required for the decapping of
all mRNAs.
Following deadenylation-dependent decapping, the body of a transcript is typically
degraded in a 5’-to-3’ direction by the exonuclease Xrn1 (Hsu & Stevens, 1993; Muhlrad et al.,
1994, 1995). To determine whether Vts1-mediated mRNA degradation requires 5’-to-3’ decay,
we measured the stability of the GFP reporters in an xrn1Δ strain. We found that GFP-SRE+
mRNA was significantly stabilized in the xrn1Δ strain over the 8-min time course (Figure 2-4).
We conclude that 5’-to-3’ decay by the exonuclease Xrn1 is required for Vts1-mediated mRNA
decay.
50
Figure 2-4 Vts1-Mediated mRNA Decay Requires Components of the Major mRNA Decay
Pathway
GFP reporter transcription was induced in the indicated strains with galactose and then shut off
with glucose and GFP mRNA levels were assayed at the times indicated after transcriptional
shutoff by Northern blot. Transcriptional pulse-chase was performed with TetO7-DCP2 in the
absence or presence of DOX as indicated. SCR1 RNA serves as a loading control. Half-lives
(T1/2) for each strain are as indicated. The results of three independent experiments were
quantitated and normalized using the levels of SCR1 RNA and graphed with error bars
representing standard deviation.
51
Thus far we have demonstrated that deadenylation, decapping and 5’-to-3’ decay are
important for Vts1-mediated mRNA degradation. However after deadenylation a transcript can
also be degraded by a complex of 3’-to-5’ exonucleases known as the exosome (Anderson &
Parker, 1998). To determine whether 3’-to-5’ decay is required for Vts1-mediated decay, we
examined the stability of the GFP reporters in a ski2Δ strain that is defective for exosome
function. The half-life of the GFP-SRE+ reporter in ski2Δ cells was similar to wild-type (Figure
2-4) indicating that the exosome does not play a significant role in Vts1-mediated decay.
Our data have shown that deadenylation, decapping and 5’-to-3’ decay are all required
for Vts1-mediated mRNA degradation, leading us to hypothesize that Vts1 targets transcripts for
deadenylation, which triggers decapping and subsequent 5’-to-3’ decay. To illustrate that
deadenylation is the first step in Vts1-mediated decay, which in turn triggers decapping and
subsequent 5’-to-3’ decay of the message, we examined the poly(A) tail length of the GFP-SRE+
mRNA in TetO7-DCP2 and xrn1Δ cells. We find that inhibition of DCP2 expression and
deletion of the XRN1 gene results in the accumulation of a stable deadenylated form of GFP-
SRE+ mRNA (Figure 2-5) indicating that deadenylation precedes decapping and 5’-to-3’
exonucleolytic decay. These data are therefore consistent with Vts1 triggering decay via
deadenylase recruitment.
2.4.5 Vts1-Mediated Decay of an Endogenous Target mRNA Requires Ccr4 and Xrn1
We have shown that Vts1 induces the degradation of artificial reporter transcripts via
deadenylation followed by decapping and 5’-to-3’ decay. To validate our model we set out to
determine if Vts1 uses this mechanism to degrade a bona fide endogenous mRNA target. We
52
Figure 2-5 Deadenylation is the First Step in Vts1-Mediated mRNA Decay
GFP reporter transcription was induced with galactose and then shut off with glucose and an
RNase H cleavage assay was used to measure the poly(A) tail length of the GFP-SRE+ reporter
mRNA in TetO7-DCP2 cells in the presence of DOX (+DOX) and in an xrn1Δ strain at the times
indicated after shutoff. Where indicated oligo(dT) was added to remove the poly(A) tail by
RNase H treatment, providing a marker for deadenylated mRNA (dT). The distribution of
poly(A) tail lengths at the indicated times after transcriptional shutoff are displayed on the
graphs. SCR1 RNA serves as a loading control.
53
previously identified 79 potential Vts1 target transcripts through a pulldown/microarray
approach and validated the interaction of Vts1 with a subset of these mRNAs by showing that we
could detect these transcripts in Vts1 immunoprecipitates via Northern blot (Aviv et al., 2006).
Of these validated transcripts a subset showed an increase in steady-state levels in vts1Δ cells
compared to wild-type, making them potential targets of Vts1-mediated decay. YIR016W mRNA
exhibited the greatest increase, and thus we examined the effect of Vts1 on YIR016W mRNA
stability by measuring the half-life of YIR016W mRNA after treatment of cells with the
transcriptional inhibitor thiolutin. Northern blotting revealed that the transcript had a half-life of
~2 min in wild-type cells while in the vts1Δ strain the transcript was significantly stabilized with
a half-life of ~6 min (Figure 2-6A). To rule out the possibility that thiolutin treatment was having
unanticipated effects on YIR016W mRNA we also generated a construct in which GFP is fused to
the YIR016W ORF under the control of the GAL1 promoter (GFP-YIR016W) and measured the
stability of this reporter mRNA after inducing reporter gene transcription with galactose and then
shutting off transcription with glucose. In these experiments GFP-YIR016W mRNA had a half-
life of <4 min in wild-type cells, and its stability was increased in the vts1Δ strain with a half-life
of ~6 min (Figure 2-6B). Taken together these data indicate that Vts1 is required for the rapid
decay of YIR016W mRNA in vivo.
Having confirmed that Vts1 is required to stimulate rapid degradation of YIR016W
mRNA, we examined the mechanisms involved by assaying the stability of both endogenous
YIR016W mRNA using thiolutin and GFP-YIR016W mRNA via transcriptional pulse-chase in
ccr4Δ and xrn1Δ cells. We found that both endogenous YIR016W mRNA and GFP-YIR016W
mRNA exhibited increased stability in these strains, indicating that Ccr4 and Xrn1 are required
for rapid decay of these transcripts (Figure 2-6A,B). Since transcripts must be decapped to be
54
Figure 2-6 Stability of YIR016W mRNA is Regulated by Vts1
(A) Transcription was inhibited by the addition of thiolutin at time zero. Endogenous YIR016W
mRNA levels were measured in wild-type (WT), vts1Δ, ccr4Δ, and xrn1Δ cells by Northern blot.
ACT1 mRNA serves as a loading control. The results of four independent experiments were
quantitated and normalized using the levels of ACT1 mRNA and graphed with error bars
representing standard deviation. The half-life (T1/2) of the mRNA in the different strains is as
indicated. (B) Transcription of the GFP-YIR016W reporter was induced with galactose and then
shut off with glucose. Reporter mRNA levels were measured in wild-type (WT), vts1Δ, ccr4Δ,
and xrn1Δ cells by Northern blot. ACT1 mRNA serves as a loading control. The results of at
least two independent experiments were quantitated and normalized using the levels of ACT1
mRNA and graphed with error bars representing standard deviation. The half-life (T1/2) of the
mRNA in the different strains is as indicated.
55
susceptible to 5’-to-3’ decay by Xrn1 (Hsu & Stevens, 1993; Muhlrad et al., 1994, 1995;
Caponigro & Parker, 1996b), we can infer that decapping is likely also required. Thus we have
confirmed that the mechanism of Vts1-mediated mRNA decay outlined using an artificial
reporter transcript holds true for an in vivo mRNA target.
2.4.6 Vts1 Localizes to P-Bodies in xrn1Δ Cells
In yeast, mRNAs undergoing degradation can be localized along with the mRNA decay
machinery, including the decapping enzymes and the 5’-to-3’ exonuclease Xrn1, to distinct
cytoplasmic foci known as processing or P-bodies (Sheth & Parker, 2003). We have shown that
deadenylation-dependent decapping and 5’-to-3’ decay are required for Vts1-mediated mRNA
degradation, suggesting that Vts1 may associate with these sites during transcript degradation.
We constructed a monomeric red fluorescent protein (mRFP) C-terminal fusion of Vts1 and
examined its localization in live cells. To visualize P-bodies we used a GFP C-terminal fusion of
Dhh1, a known P-body component (Sheth & Parker, 2003). In wild-type cells Vts1-mRFP does
not co-localize with Dhh1 (Figure 2-7). Given the apparent rapid rate of Vts1-mediated decay,
we theorized that the association of Vts1 with P-bodies may be too transient to observe in wild-
type cells under standard conditions. We therefore examined the localization of Vts1-mRFP in an
xrn1Δ strain which is blocked for 5’-to-3’ decay and therefore retains degrading transcripts in P-
bodies (Sheth & Parker, 2003). In xrn1Δ cells, Vts1-mRFP was concentrated in foci that co-
localize extensively with Dhh1-GFP (Figure 2-7). These results indicate that Vts1-mRFP
localizes to P-bodies in an xrn1Δ strain and suggest that Vts1-mediated mRNA degradation may
take place in P-bodies.
56
Figure 2-7 Vts1 Localizes to P-Bodies in xrn1Δ Cells
Dhh1-GFP (left panel) and Vts1-mRFP (middle panel) were coexpressed in wild-type or xrn1Δ
cells and visualized by confocal microscopy. The merged images are shown in the right panel.
57
2.5 Discussion
Here we detail the molecular mechanism of Vts1-mediated mRNA decay using both an in
vivo reporter mRNA and a bona fide target of Vts1. Our data are consistent with a model
whereby Vts1 induces their degradation via Ccr4-mediated deadenylation. Vts1
coimmunoprecipitates with Pop2 in an RNA-independent manner, suggesting that Vts1 interacts
with the Ccr4-Pop2-Not deadenylase complex. Whether Vts1 interacts directly with one or more
components of the deadenylase or indirectly through another protein remains to be determined.
While our data do not rule out other possibilities, they are consistent with a model whereby Vts1
bound to an mRNA facilitates deadenylase recruitment to that transcript, which in turn triggers
mRNA destabilization.
Deadenylation-dependent decapping followed by 5’-to-3’ decay by the exonuclease Xrn1
has been identified as the major mRNA decay pathway in yeast, and our work demonstrates that
Vts1 functions within this pathway. However, microarray experiments suggest that fewer than
20% of yeast mRNAs are significantly up-regulated in dcp1Δ or xrn1Δ strains, suggesting other
mechanisms play significant roles (He et al., 2003). Clearly additional work is required to
determine the extent to which different mechanisms contribute to global regulation of mRNA
degradation.
2.5.1 Smg Family Members Employ a Conserved Mechanism to Induce Transcript Degradation
Previous work suggests that Drosophila Smg employs a similar mechanism, involving
deadenylation mediated by the recruitment of the Ccr4-Pop2-Not complex, to initiate the
degradation of target mRNAs (Semotok et al., 2005). While Smg initiates transcript decay
through a similar mechanism to that of Vts1, the precise mechanism that underlies decay after
58
deadenylation is not known. Nonetheless, the mechanistic similarities raise the strong possibility
that regions conserved between Vts1 and Smg are responsible for deadenylase binding. Both
Vts1 and Smg carry an as yet uncharacterized region designated Smg similarity region 1 as well
as a SAM domain, either of which could be responsible for deadenylase binding. As these
regions are also present in Smg homologs in other species, this would support a model whereby
destabilization of mRNAs by Ccr4-Pop2-Not recruitment is evolutionarily conserved among the
Smg family. However, human Smg 1 represses the translation of a cotransfected artificial
reporter gene without inducing transcript degradation (Baez & Boccaccio, 2005). Either this
protein has lost the ability to induce transcript degradation or the cell type used in these
experiments lacks a factor required to destabilize mRNAs.
Several other RNA-binding proteins have been suggested to induce transcript decay via
recruitment of the Ccr4-Pop2-Not deadenylase complex. These include PUF family members,
A+U-rich element RNA binding proteins TTP and BRF-1, the cold-shock-domain-containing
protein UNR, and the Drosophila protein Bicaudal C (Chang et al., 2004; Lykke-Andersen &
Wagner, 2005; Goldstrohm et al., 2006; Chicoine et al., 2007; Goldstrohm et al., 2007; Hook et
al., 2007). Thus, deadenylase recruitment may be a common mechanism employed to regulate
the stability of specific mRNAs.
2.5.2 The Role of Foci in the Function of Smg Family Members
Vts1 does not localize to P-bodies in wild-type cells but can be found in these structures
in an xrn1Δ strain. While these data could suggest that Vts1-mediated mRNA degradation takes
place in P-bodies, it is also possible that P-bodies play no role in Vts1 function and more work
will be required to address this issue. Nonetheless, the localization of Vts1 to cytoplasmic foci is
not a unique phenomenon among the Smg protein family. The intracellular distribution of
59
Drosophila Smg is highly punctate with foci of varying size that show a modest colocalization
with P-bodies (Zaessinger et al., 2006). Neither the composition of these Smg foci nor their
functional significance is currently known. When expressed in fibroblasts, human Smg 1 forms
cytoplasmic foci that lack GW182, a P-body marker, but contain several proteins that are found
in stress granules (Baez & Boccaccio, 2005). Stress granules are induced by environmental stress
and contain nontranslating mRNAs, a subset of translation initiation factors, 40S ribosomal
subunits and a number of RNA-binding proteins (Anderson & Kedersha, 2006). While P-bodies
and stress granules are distinct structures, they share various components and under some
circumstances physically interact with one another. Whether the human Smg foci represent a
novel function for this protein or are in some way related to the function of the yeast and
Drosophila family members awaits further investigation.
60
Chapter 3
The eIF4E-Binding Protein Eap1 Functions in Vts1-Mediated
Transcript Decay
Contribution of work: I performed the majority of the experimental work presented in this
Chapter.
H. Vari performed the transcriptional pulse-chase experiments to measure the stability of the
GFP-SRE- reporter (Figure 3-2).
61
Chapter 3
The eIF4E-Binding Protein Eap1 Functions in Vts1-Mediated Transcript
Decay
3.1 Abstract
Sequence-specific RNA-binding proteins can induce the degradation of mRNAs through
their ability to recruit proteins that trigger transcript destabilization. For example, Vts1, the S.
cerevisiae member of the Smaug family of RNA-binding proteins, is thought to induce transcript
decay by recruiting the Ccr4-Pop2-Not deadenylase complex to target mRNAs. The resulting
deadenylation triggers transcript decapping followed by 5’-to-3’ exonucleolytic decay. Here we
show that the eIF4E-binding protein, Eap1, is required for efficient degradation of Vts1 target
transcripts, and that this role involves the ability of Eap1 to interact with eIF4E. Furthermore
Eap1 does not function in deadenylation but is instead involved in transcript decapping. We also
show that Eap1 interacts with Vts1 and mediates an indirect interaction between Vts1 and eIF4E.
Taken together these data suggest a model whereby Vts1-mediated recruitment of Eap1 to target
mRNAs stimulates decapping. While Eap1 is not a component of the general mRNA decay
machinery, it also functions in Puf3-triggered destabilization of COX17 mRNA. Thus, Eap1
might function with other RNA-binding proteins to stimulate transcript decay.
62
3.2 Introduction
The regulation of mRNA degradation has an important role in the control of gene
expression. In Saccharomyces cerevisiae the major mRNA decay pathway is initiated through
transcript deadenylation mediated by the Ccr4-Pop2-Not complex (Decker & Parker, 1993; Hsu
& Stevens, 1993; Muhlrad et al., 1995). After deadenylation, the transcript is decapped by a
heterodimeric complex composed of Dcp1 and Dcp2 [reviewed in (Coller & Parker, 2004;
Parker & Song, 2004)]. In yeast, numerous factors that positively regulate mRNA decapping
have been identified, including Pat1, Dhh1, Edc1, Edc2, Edc3 and the Lsm1-7 complex
[reviewed in (Coller & Parker, 2004; Parker & Song, 2004)]. Although mutations in genes
encoding these factors inhibit decapping, the mechanism through which they regulate this step of
the mRNA decay process remains to be determined. After decapping, the body of the transcript
is degraded 5’-to-3’ by the exonuclease Xrn1 (Hsu & Stevens, 1993; Muhlrad et al., 1994).
Sequence-specific RNA-binding proteins can add another level of control to the
regulation of mRNA stability (Garneau et al., 2007). Typically these proteins bind mRNA target
sequences and interact with other trans factors that influence the rate of mRNA decay. The
Smaug (Smg) family of post-transcriptional regulators, which are conserved from yeast to
humans, bind RNA through a conserved sterile alpha motif (SAM) domain that interacts with
stem-loop structures termed Smg recognition elements (SREs) (Smibert et al., 1996; Dahanukar
et al., 1999; Smibert et al., 1999; Crucs et al., 2000; Aviv et al., 2003; Green et al., 2003; Baez &
Boccaccio, 2005; Semotok et al., 2005; Aviv et al., 2006; Semotok et al., 2008). Vts1, the Smg
family member in S. cerevisiae, stimulates mRNA degradation through deadenylation mediated
by the Ccr4-Pop2-Not deadenylase complex (Aviv et al., 2003; Rendl et al., 2008). Following
deadenylation Vts1 target transcripts are decapped and then degraded by the 5’-to-3’ exonuclease
63
Xrn1 (Rendl et al., 2008). A similar mechanism of deadenylation-dependent mRNA decay is
employed by Smg in Drosophila (Semotok et al., 2005; Zaessinger et al., 2006; Semotok et al.,
2008). Both Vts1 and Smg interact with the Ccr4-Pop2-Not complex, suggesting a model
whereby these proteins induce transcript decay by recruiting the deadenylase to target mRNAs.
Smg also regulates mRNA translation through a separate mechanism involving an interaction
with the eIF4E-binding protein Cup (Nelson et al., 2004). Cup binds to the mRNA cap binding
protein eIF4E through a canonical eIF4E-binding motif (YXXXXLΦ, where Φ is a hydrophobic
amino acid). Cap-dependent translation initiation involves eIF4E recruiting eIF4G to an mRNA,
which indirectly mediates recruitment of the 40S ribosome (Gingras et al., 1999). eIF4G also
interacts with eIF4E through an eIF4E-binding motif, and thus recruitment of Cup to an mRNA
inhibits translation by blocking the eIF4E/eIF4G interaction (Mader et al., 1995; Nelson et al.,
2004).
The role of Cup in Smg function led us to speculate that Vts1 might also regulate target
mRNAs through an eIF4E-binding protein. While there is no Cup homolog in yeast, two eIF4E-
binding proteins, Caf20 and Eap1, have been identified (Altmann et al., 1997; de la Cruz et al.,
1997; Cosentino et al., 2000). In addition, global genetic analysis revealed synthetically lethal
interactions between Eap1 and two deadenylase components, Ccr4 and Pop2 (Pan et al., 2006),
suggesting a functional relationship, either direct or indirect, among the gene products. This
genetic interaction, combined with the role of the Ccr4-Pop2-Not deadenylase in Vts1-mediated
regulation, prompted us to test if Eap1 might function with Vts1 to regulate target mRNAs.
Using two different Vts1 target mRNAs, we demonstrate that Eap1 is required for efficient Vts1-
mediated transcript degradation. We show that Eap1 is not a basic component of the RNA decay
machinery and does not function at the level of deadenylation but is instead required for efficient
64
removal of the 5’ cap. In addition, Eap1-mediated stimulation of transcript decay requires
binding to eIF4E. We also find that Eap1 biochemically interacts with Vts1 and it is able to
mediate an indirect interaction between Vts1 and eIF4E. Taken together these data suggest a
model whereby formation of the Vts1/Eap1/eIF4E complex stimulates transcript decapping.
While Eap1 is not a general RNA decay factor, it is required for efficient degradation of COX17
mRNA, whose degradation is induced by another sequence-specific RNA-binding protein, Puf3.
Thus, Eap1 may function with other RNA-binding proteins, in addition to Vts1, to stimulate
transcript decay of a subset of mRNAs recognized by these, and perhaps other, sequence-specific
RNA-binding proteins.
3.3 Materials and Methods
3.3.1 Yeast Strains and Media
Yeast strains used in this study were derivatives of the wild-type BY4741 (Brachmann et
al., 1998). All deletion strains are as described by Winzeler et al. (1999), with the exception of
the eap1Δ strain which was generated for this study by directed PCR-mediated gene deletion
(MATa eap1Δ::KANMX6 leu2Δ0 his3Δ1 ura3Δ0 met15Δ0). The TetO7-DCP2 strain is described
in Mnaimneh et al. (2004). The strains were transformed by standard techniques, and plasmids
were maintained by growth in selective media.
3.3.2 mRNA Analysis
Transcriptional pulse-chase experiments employed GFP-SRE+, GFP-SRE-, and GFP-
YIR016W reporters described in Rendl et al. (2008). For these experiments cells were grown in
selective medium containing 2% raffinose at 30oC to mid-log phase and then cooled to 20oC,
with the exception of Figure 3-1C and 3-7, in which transcriptional shut-off was performed at
65
30oC. Due to the short half-life of the GFP-SRE+ reporter in wild-type cells, an accurate measure
of reporter mRNA stability at 30oC was not possible. Cooling cells to 20oC slowed the decay of
GFP-SRE+ mRNA to allow for measurement of reporter mRNA half-lives. GFP reporter
transcription was initiated by the addition of galactose to a final concentration of 2% for 16 min
and then repressed by the addition of glucose to a final concentration of 4%. In Figure 3-4 TetO7-
DCP2 cells were treated with 10 µg/mL doxycycline for 16 hours before RNA reporter
transcription was induced with galactose.
Total RNA was isolated by glass bead lysis in LET buffer (100 mM LiCl, 20 mM EDTA,
25 mM Tris-HCl at pH 8.0) and LET-equilibrated phenol at the indicated time points and
analyzed by Northern blot. SCR1 or ACT1 RNA was used for normalization of transcript levels
as indicated. Where indicated, thiolutin (Carbosynth Limited) was added to cultures to a final
concentration of 20 µg/mL at time 0 of the time course.
RNase H cleavage assays were performed as described by Decker and Parker (1993),
using an oligonucleotide that hybridized ~330 nt upstream of the GFP reporter poly(A) site. To
control for differences in RNA concentration, a portion of each RNA sample was analyzed via a
standard Northern blot and probed for SCR1 RNA. Northern blots were exposed to
PhosphorImager screens and analyzed with ImageQuant Software.
For RNA immunoprecipitations total RNA was isolated from cells expressing the GFP-
SRE+ reporter during transcriptional pulse-chase experiments 16 min after transcriptional shut-
off with glucose. Immunoprecipitations were performed as described in Muhlrad et al. (1994)
using monoclonal anti-m3G-/m7G-cap (Synaptic Systems) with the following modifications.
Anti-m3G-/m7G-cap antibodies were prebound at 4oC to protein G-Agarose (Roche) in IPPH
66
(500 mM NaCl, 10 mM Tris-Cl at pH 7.5, 0.1% NP-40) for 2 h. The beads were extensively
washed in IPPL (150 mM NaCl, 10 mM Tris-Cl at pH 7.5, 1 mM EDTA, 0.05% NP-40) and then
incubated at 4oC with 5 µg total RNA in IPPL supplemented with 1 mM DTT and 0.1 U/µL
RNasin (Promega) for 2 h. The supernatant was recovered and incubated for 2 h at 4oC a second
time with antibody prebound to beads. The pellets were washed twice with IPPL and combined.
Bound and unbound mRNA were then isolated from the pellet and supernatant fractions and
analyzed by Northern blot.
For Figure 3-6, transcriptional pulse-chase experiments were performed to analyze
reporter mRNA stability as described above. For analysis of protein expression a sample of cells
at time 0 were denatured by trichloroacetic acid (TCA) precipitation, lysed with glass beads and
the resulting extracts fractionated on SDS-PAGE and analyzed by Western blot using HA
antibodies (Santa Cruz) and PSTAIR antibodies (Sigma) as indicated.
3.3.3 Immunoprecipitations
Immunoprecipitation was performed essentially as described previously in Rendl et al.
(2008). The plasmid expressing Vts1-Flag was generated in pRS315 with a C-terminal Flag tag
and 574 bases of genomic sequence upstream and 396 bases downstream of the VTS1 ORF. The
plasmid expressing Vts1RBD--Flag was generated in pRS313 and is identical in sequence to
Vts1-Flag with the exception of the A498Q amino acid change. All Vts1 constructs also harbor
VSV epitopes that were used for protein detection by Western blot. The Eap1-HA plasmid was
created in pRS316 with a C-terminal 3X HA tag and 490 bases of genomic sequence upstream
and 492 bases downstream of the EAP1 ORF. The Eap1mt-HA construct was generated in
pRS316 and is identical in sequence to Eap1-HA with the exception of the Y109A;L114A amino
acid changes. Preparation of protein extracts by glass bead lysis and subsequent
67
immunoprecipitation experiments were preformed as previously described in Rendl et al. (2008).
Briefly anti-Flag M2 affinity gel (Sigma) was used to immunoprecipitate Flag-tagged proteins
for 3 h at 4oC. After extensive washing Vts1-Flag and associated proteins were eluted with Flag-
peptide (Sigma). VSV antibodies (Bethyl Laboratories), HA antibodies (Santa Cruz) and eIF4E
antibodies (a generous gift from N. Sonenberg (Cosentino et al., 2000)) were used to detect
eluted proteins by Western Blot.
3.4 Results
3.4.1 Eap1 is Required for Efficient Decay of Vts1 Target mRNAs
To assess the role of Eap1 in Vts1 function we first examined the stability of a reporter
mRNA that recapitulates Vts1-mediated decay in vivo (Aviv et al., 2003). The GFP-SRE+
reporter encodes green fluorescent protein (GFP) under the control of the inducible galactose
promoter, and has three SREs in its 3’ untranslated region (UTR). A transcriptional pulse-chase
approach was used to measure the stability of reporter mRNAs by Northern blotting after
transcriptional induction by galactose and subsequent repression by the addition of glucose. We
previously reported that GFP-SRE+ mRNA is rapidly degraded (half-life of ~4 min), but is
stabilized in a vts1Δ strain (half-life of >8 min) (Rendl et al., 2008). Here we show that rapid
degradation of GFP-SRE+ mRNA was also compromised in eap1Δ cells, resulting in a doubling
of the half-life to ~8 min (Figure 3-1A and B). The fact that the GFP-SRE+ mRNA was
stabilized more in a vts1Δ strain than in an eap1Δ strain suggests that, while Eap1 plays a role in
the decay of this mRNA, it is not absolutely required for Vts1 function.
To further confirm the importance of Eap1 in the degradation of Vts1 target mRNAs, we
measured the stability of YIR016W mRNA in eap1Δ cells, having previously shown that Vts1
68
Figure 3-1 Eap1 is Required for Efficient Degradation of Vts1 Target mRNAs
Reporter gene transcription, either GFP-SRE+ (A) or GFP-YIR016W (C), was induced in eap1Δ
cells with galactose and then shut off with glucose, and mRNA levels were assayed by Northern
blot at the times indicated after transcriptional shutoff. SCR1 RNA serves as a loading control.
The results of at least three independent experiments were quantitated and normalized using the
levels of SCR1 RNA and plotted with error bars representing standard deviation. Half-lives (T1/2)
of the mRNAs in the different strains are as indicated. The result of t tests comparing the percent
of RNA remaining, either GFP-SRE+ (B) or GFP-YIR016W (D), in wild-type (WT) and eap1Δ
cells at the end of the time course is indicated, with error bars representing standard deviation.
Note that wild-type and vts1Δ data from Figure 2-1 and Figure 2-6 is included for comparison.
69
binds to this mRNA, and regulates its stability through deadenylation, decapping and 5’-to-3’
exonucleolytic decay (Aviv et al., 2006; Rendl et al., 2008). We used a reporter construct in
which GFP is fused to the YIR016W ORF under the control of the GAL1 promoter (GFP-
YIR016W). GFP-YIR016W reporter transcription was induced by adding galactose to eap1Δ
cells and then shut off once again by the addition of glucose. Similar to our findings using the
GFP-SRE+ reporter, we found that the stability of GFP-YIR016W mRNA was increased in the
eap1Δ strain as compared to wild-type resulting in significantly more reporter mRNA present in
the final time point compared to wild-type cells (Figure 3-1C and D). In addition, the stability of
YIR016W mRNA was significantly enhanced in eap1Δ cells as assayed after treatment of cells
with the transcriptional inhibitor thiolutin (data not shown). Taken together these data indicate
that, although not absolutely essential, Eap1 is required for the rapid decay of Vts1 target
mRNAs.
In principle Eap1 could function with Vts1 to repress the translation of target mRNAs
similar to the role of Cup in Smg function. However, we have been unable to document a role
for Vts1 in regulating translation that is independent of transcript decay. For example, we have
shown that GFP protein levels from the GFP-SRE+ mRNA are repressed approximately four fold
compared to GFP-SRE- mRNA and that this repression is eliminated in a vts1Δ strain ((Rendl et
al., 2008), and data not shown). However, when GFP protein levels were corrected using the
levels of GFP mRNA we did not see a statistically significant difference in the ratio of GFP
protein levels from the GFP-SRE+ and GFP-SRE- mRNAs in wild-type, vts1Δ and eap1Δ cells
(data not shown). These results suggest that repression of the GFP-SRE+ reporter expression is
mediated largely through degradation of target mRNAs, however we cannot rule out some role
for translational repression mediated by Vts1 and/or Eap1 that our assays are not sensitive
70
enough to detect. Alternatively, since the poly(A) tail is known to stimulate translation, Vts1-
mediated deadenylation of target mRNAs could simultaneously repress translation and induce
transcript decay; if so, the two processes would not be separable.
3.4.2 Eap1 is Not a Core Component of the mRNA Decay Machinery
The role of Eap1 in the degradation of Vts1 target mRNAs could indicate that Eap1 is a
core component of the major mRNA decay pathway, which involves deadenylation by the Ccr4-
Pop2-Not complex, followed by decapping and 5’-to-3’ decay. Alternatively, the role of Eap1
could be more specific, perhaps reflecting a direct function in Vts1-mediated decay. To explore
these alternative possibilities, we assessed the stability of a GFP reporter mRNA (GFP-SRE-),
which is identical to the GFP-SRE+ reporter with the exception that it carries SREs in which the
loop sequences are mutated to block Vts1 binding (Aviv et al., 2003). Transcriptional pulse-
chase experiments in wild-type and eap1Δ cells have demonstrated that Eap1 does not play a role
in regulating the stability of GFP-SRE- mRNA (Figure 3-2A). Instead, the GFP-SRE- mRNA is
destabilized through the major mRNA decay pathway, as degradation required Ccr4 (the
catalytic subunit of the Ccr4-Pop2-Not deadenylase) and the 5’-to-3’ exonuclease Xrn1 (Figure
3-2B) in a Vts1-independent manner (Figure 3-2A). Thus, Eap1 is not an integral component of
the major mRNA decay pathway. Instead, a role for Eap1 in the degradation of our GFP-SRE+
and GFP-YIR016W reporters is a consequence of Vts1 targeting the mRNAs for degradation,
suggesting that Eap1 plays a specific role in mRNA degradation.
Interestingly, these experiments demonstrate that GFP-SRE- mRNA is less stable in a
vts1Δ strain (Figure 3-2A). We speculate that the physical interaction between Vts1 and the
Ccr4-Pop2-Not deadenylase complex (Rendl et al., 2008) in wild-type cells sequesters some
fraction of the deadenylase into a pool that is unable to act on mRNAs that are not targeted by
71
Figure 3-2 Eap1 is Not Required for the Degradation of GFP-SRE- mRNA
GFP-SRE- gene transcription was induced in wild-type (WT), vts1Δ and eap1Δ cells (A) and
wild-type (WT), ccr4Δ and xrn1Δ cells (B) with galactose and then shut off with glucose and
reporter mRNA levels were assayed at the times indicated after transcriptional shutoff by
Northern blot. The results of at least two independent experiments were quantitated and
normalized using the levels of SCR1 RNA and graphed with error bars representing standard
deviation. The half-life (T1/2) of the mRNA in the different strains is as indicated.
72
Vts1. In a vts1Δ strain this pool is no longer sequestered and as such is free to act on the GFP-
SRE- mRNA, thereby accelerating its deadenylation and subsequent degradation.
3.4.3 Eap1 Does Not Function in mRNA Deadenylation
We next set out to identify at which step in the mRNA decay process Eap1 functions.
Given that Vts1 triggers mRNA decay via poly(A) tail shortening, mediated by the Ccr4-Pop2-
Not deadenylase complex, we analyzed the poly(A) tail length of GFP-SRE+ mRNA in eap1Δ
cells to determine whether Eap1 was required for deadenylation of Vts1 target mRNAs. We used
an RNase H cleavage method to measure the poly(A) tail length of the GFP-SRE+ mRNA over
an 8-min transcriptional pulse-chase time course. Total RNA samples from each time point were
treated with RNase H in the presence of a specific antisense oligonucleotide that hybridized to
the GFP open reading frame. After RNase H cleavage, the poly(A) tail of the resulting short 3’-
end fragment of the GFP reporter mRNA was measured by high resolution polyacrylamide
electrophoresis and Northern blotting. To one sample reporter-specific oligonucleotide and
oligo(dT) was added; treatment with RNase H yielded a 3’-end fragment lacking a poly(A) tail,
which served as a marker for the deadenylated 3’-end fragment.
Comparison of the RNase H cleavage data for eap1Δ cells (Figure 3-3) to our pervious
analysis in wild-type cells (Rendl et al., 2008) indicated that Eap1 is not involved in the
deadenylation of GFP-SRE+ mRNA. For example, eap1Δ cells at time zero showed two
populations of GFP-SRE+ mRNA, similar to wild-type cells. One population had tail lengths
from 30 to 50 nucleotides while the other had very short tails presumably representing transcripts
that were deadenylated during the 16-min induction with galactose, indicating the transcript was
being rapidly deadenylated. In addition, in eap1Δ cells soon after the addition of glucose, we
73
Figure 3-3 Eap1 is Not Required for Transcript Deadenylation
GFP-SRE+ reporter transcription in eap1Δ cells (A) and wild-type cells (B) was induced with
galactose and then shut off with glucose and an RNase H cleavage assay was used to measure the
poly(A) tail length of the reporter transcript at the times indicated after transcriptional shutoff.
Where indicated oligo(dT) was added to remove the poly(A) tail by RNase H treatment,
providing a marker for deadenylated mRNA (dT). The distribution of poly(A) tail lengths at the
indicated times after transcriptional shutoff are displayed on the graphs. SCR1 RNA serves as a
loading control. Note that wild-type data from Figure 2-2 is included for comparison.
74
consistently saw a decrease in the amount of longer poly(A) tail species and an increase in the
amount of the shorter tail population (compare the distribution of poly(A) tail lengths at time
zero to the 2-min time point) similar to that seem in wild-type again indicating no deadenylation
defect in eap1Δ cells (Figure 3-3). At later time points deadenylated species persisted, similar to
results we reported for mutants that are wild-type for deadenylation but defective for
downstream steps such as decapping and 5’-to-3’ exonucleolytic decay (Rendl et al., 2008).
Taken together these data suggest that Eap1 functions subsequent to deadenylation in the
degradation of GFP-SRE+ mRNA.
3.4.4 Eap1 Stimulates mRNA Decapping
After deadenylation Vts1 target transcripts are decapped and then degraded in a 5’-to-3’
direction by the exonuclease Xrn1 (Rendl et al., 2008). To determine if Eap1 is involved in the
decapping of Vts1 target transcripts we made use of an antibody that recognizes the mRNA 5’
cap structure. A transcriptional pulse-chase experiment was performed and RNA harvested 16-
min post glucose addition was immunoprecipitated with the 5’ cap antibody. The levels of GFP-
SRE+ mRNA in the immunoprecipitates and in the flow through were assayed via Northern blot
to provide a quantitative measure of the percentage of capped versus uncapped mRNA. The
results from eap1Δ cells were compared to results obtained from cells depleted of Dcp2, the
catalytic subunit of the decapping complex, and xrn1Δ cells, which are defective for 5’-to-3’
exonucleolytic decay. Depletion of Dcp2 was achieved using a strain expressing DCP2 under a
tetracycline regulatable promoter (TetO7-DCP2), where inhibition of DCP2 transcription results
from addition of the tetracycline analog doxycycline (Mnaimneh et al., 2004).
75
In these experiments we recovered on average ~50% of the GFP-SRE+ mRNA. When
cells were depleted of Dcp2 and thereby defective for decapping, GFP-SRE+ mRNA was
enriched 2.4 fold in the immunoprecipitated material, with 36.6% on average of the total GFP-
SRE+ mRNA present in the immunoprecipitated fraction and 15.2% on average present in the
supernatant (Figure 3-4). In contrast, GFP-SRE+ mRNA was not enriched in the
immunoprecipitated material from xrn1Δ cells, with similar levels of GFP-SRE+ mRNA found in
immunoprecipitated and supernatant fractions. Control experiments confirm the specificity of the
antibody for the 5’ cap structure as on average ≤3.5% of the total SCR1 RNA, which is a RNA
polymerase III transcript that lacks a cap structure (Briand et al., 2001), was precipitated in any
experiment. In eap1Δ cells, GFP-SRE+ mRNA was enriched 3 fold in immunoprecipitated
material, with 42% on average found in the immunoprecipitated fraction and 14% on average in
the supernatant. The similar enrichment of GFP-SRE+ mRNA in Dcp2 depleted cells and eap1Δ
cells indicate that Eap1 is required for efficient decapping of GFP-SRE+ mRNA.
3.4.5 Eap1 Mediates an Interaction Between Vts1 and eIF4E
Given the requirement of Eap1 for Vts1-mediated mRNA decay, we sought to determine
whether Vts1 physically interacts with Eap1 in vivo. We performed co-immunoprecipitation
experiments using whole-cell lysates from cells expressing Flag-tagged Vts1 and HA-tagged
Eap1. Vts1-Flag was immunoprecipitated using anti-Flag resin, and the immunoprecipitates were
analyzed by Western blot. Eap1-HA was present in the anti-Flag immunoprecipitate when the
Vts1-Flag protein was present (Figure 3-5A). As a control, Eap1-HA was not
immunoprecipitated from an extract lacking Vts1-Flag. The co-immunoprecipitation of Vts1
with Eap1 was RNA-independent, as it was observed when Vts1 harbored an amino acid change
(A498Q) that blocks its ability to bind RNA (Vts1RBD-) (Aviv et al., 2003).
76
Figure 3-4 Eap1 Stimulates Transcript Decapping
Total RNA was isolated from the indicated cells expressing the GFP-SRE+ reporter during
transcriptional pulse-chase experiments 16 min after transcriptional shutoff with glucose. These
samples were subjected to immunoprecipitation using an antibody that recognizes the 5’ cap and
equivalent amounts of RNA from starting material (T), immunoprecipitated material (P), and
supernatant from the immunoprecipitates (S) were assayed for GFP-SRE+ mRNA and SCR1
RNA levels via Northern blot. The results of three independent experiments were quantitated and
the average percentage of material in each sample, along with the standard deviation, are
indicated.
77
Figure 3-5 Eap1 Mediates an Indirect Interaction Between Vts1 and eIF4E
Vts1-Flag and Eap1-HA were expressed in yeast cells individually or in combination as
indicated in either vts1Δ cells (A) or eap1Δ cells (B) and crude extracts were subjected to
immunoprecipitation using anti-Flag resin. Starting crude extracts (input) and the resulting
immunoprecipitates (IP: αFLAG) were analyzed by Western blot. Input samples represent 5%
for Vts1 blots, 2% for Eap1 blots and 1% for eIF4E blots of material used in the
immunoprecipitations. The use of the Vts1 RNA-binding mutant (Vts1RBD-) and Eap1 mutant
defective for eIF4E binding (Eap1mt) are as indicated.
78
The interaction of Vts1 with the eIF4E-binding protein led us to hypothesize that Eap1
may mediate an interaction between Vts1 and eIF4E. To test whether Vts1 interacts with eIF4E
via Eap1 we repeated our co-immunoprecipitation experiments using whole cell lysates derived
from eap1Δ cells. Consistent with data presented above, Eap1-HA was present in the anti-Flag
immunoprecipitate when the Vts1-Flag protein was present (Figure 3-5B). eIF4E was also
detected in Vts1-Flag immunoprecipitates only in the presence of Eap1. eIF4E was not
significantly enriched when Vts1-Flag was immunoprecipitated from lysate expressing Eap1 that
harbored amino acid changes (Y109A;L114A) in the eIF4E-binding motif that block its ability to
bind eIF4E (Eap1mt) (Ibrahimo et al., 2006). In addition, we did not detect a significant
interaction between Eap1mt-HA and Vts1-Flag, suggesting that binding of Eap1 to eIF4E may
stabilize the Vts1/Eap1 interaction. Taken together these data indicate that Eap1 mediates an
interaction between Vts1 and eIF4E through its eIF4E-binding motif.
3.4.6 Binding to eIF4E is Required for Eap1 to Stimulate Transcript Decay
To explore the molecular mechanism that underlies the role of Eap1 in transcript decay
we asked if the Eap1/eIF4E interaction was required for mRNA degradation. This involved
assaying the stability of GFP-SRE+ mRNA by transcriptional pulse-chase in eap1Δ cells
expressing either wild-type Eap1 (Eap1-HA) or a version Eap1 harboring the Y109A;L114A
mutations that disrupt its eIF4E-binding ability (Eap1mt-HA) (Ibrahimo et al., 2006). When
Eap1-HA was expressed in the eap1Δ strain, GFP-SRE+ mRNA decayed at a rate similar to wild-
type (with a half-life ~4 min and similar levels of mRNA remaining in the last time point of the
time course) (Figure 3-6A, C and D). However when Eap1mt-HA was expressed in the eap1Δ
strain, the decay rate of GFP-SRE+ mRNA was similar to that observed in the eap1Δ strain alone
79
Figure 3-6 The Eap1/eIF4E Interaction is Required for Efficient Decay of Vts1 Target
mRNAs
eap1Δ cells were rescued with either wild-type Eap1-HA or a mutant version of Eap1 that is
defective in its ability to interact with eIF4E (Eap1mt-HA) and the stability of GFP-SRE+ mRNA
was assessed in these cells using a transcriptional pulse-chase approach and Northern blot (A) as
described in Figure 3-1. Western blot (B) was used to compare the expression of wild-type Eap1-
HA to that of Eap1mt-HA using the PSTAIR protein as a loading control. The results of four
independent experiments were quantitated and normalized using the levels of SCR1 RNA and
graphed with error bars representing standard deviation (C). Half-lives (T1/2) of the mRNAs in
the different strains are as indicated (A). The result of t tests comparing the percent of GFP-
SRE+ RNA remaining in the indicated cells at the end of the time course is indicated with error
bars representing standard deviation (D). Note that wild-type data from Figure 2-1 and eap1Δ
data from Figure 3-1 is included for comparison.
80
(with a half-life ~8 min and similar levels of mRNA remaining in the last time point of the time
course). Western blot analysis confirmed that both Eap1-HA and Eap1mt-HA were expressed at
similar levels, and therefore differences in protein expression cannot account for the observed
difference in mRNA decay kinetics (Figure 3-6B). These data demonstrate that a functional Eap1
eIF4E-binding motif is required for the rapid degradation of GFP-SRE+ mRNA.
3.4.7 Eap1 Enhances the Degradation of Another Endogenous mRNA
Our data indicate that Eap1 is not a general component of the RNA degradation
machinery and that it is required for the degradation of Vts1 target mRNAs. To further examine
the specificity of Eap1 in mRNA degradation we tested its role in the degradation of a transcript
that is regulated by another sequence-specific RNA-binding protein, Puf3. Puf3 promotes the
decay of COX17 mRNA by binding to specific sequences in the 3’UTR and enhancing the rate of
transcript deadenylation and subsequent degradation by the major mRNA decay pathway (Olivas
& Parker, 2000; Tucker et al., 2002; Jackson et al., 2004; Houshmandi & Olivas, 2005). We
measured the half-life of COX17 mRNA after treatment of cells with the transcriptional inhibitor
thiolutin. Northern blotting revealed that the transcript had a half-life of ~9 min in wild-type and
<6 min in vts1Δ cells, confirming that COX17 is not a target of Vts1-mediated mRNA decay
(Figure 3-7). The decreased stability of COX17 mRNA in vts1Δ cells compared to wild-type is
similar to that seen for GFP-SRE- mRNA (Figure 3-2) and is consistent with the model discussed
above whereby Vts1 sequestration of the Ccr4-Pop2-Not complex regulates the stability of
mRNAs that are not bound by Vts1. Interestingly we found that the half-life of COX17 mRNA
increased to >12 min in the eap1Δ strain, indicating that Eap1 is involved in the regulation of
other transcripts outside of Vts1 target mRNAs.
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Figure 3-7 The Stability of COX17 mRNA is Regulated by Eap1
Transcription was inhibited by the addition of thiolutin at time zero and endogenous COX17
mRNA levels were measured in wild-type (WT), vts1Δ, and eap1Δ cells by Northern blot. ACT1
mRNA serves as a loading control. The results of two independent experiments were quantitated
and normalized using the levels of ACT1 mRNA and graphed with error bars representing
standard deviation. The half-life (T1/2) of the mRNA in the different strains is as indicated.
82
3.5 Discussion
Here we demonstrate a role for Eap1 in Vts1-mediated mRNA decay. The ability of Eap1
to interact with the cap-binding protein eIF4E is required for this function and, consistent with
this proximity to the 5’ cap, Eap1 stimulates mRNA decapping. While Eap1 functions in the
decay of a reporter mRNA bearing wild-type SREs, it does not play a role in the degradation of a
reporter bearing mutated non-functional SREs. This suggests that Eap1 is not a component of
the basic mRNA degradation machinery but instead functions with Vts1 in the decay of target
mRNAs. In addition, we show that Vts1 interacts with Eap1 and that Eap1 is able to mediate an
indirect interaction between Vts1 and eIF4E. Taken together these data suggest a model
whereby recruitment of Eap1 to target transcripts, through its interaction with Vts1, stimulates
decapping via binding to eIF4E. The role of Eap1 in the degradation of COX17 mRNA, which is
not a Vts1 target, suggests that Eap1 stimulates decapping of additional unstable mRNAs,
perhaps via recruitment by other RNA-binding proteins.
3.5.1 The Role of Eap1 in mRNA Decapping
What is the molecular mechanism that underlies the role of Eap1 in decapping? The
mRNA cap structure stimulates translation through the ability of eIF4E to recruit eIF4G to the
mRNA and a critical step in mRNA decapping is thought to involve removal of the translation
initiation complex from the mRNA. This model is based on results arguing that the decapping
and translation machineries compete with one another for access to mRNAs (Schwartz & Parker,
1999; Vilela et al., 2000). For example, disruption of eIF4G function in yeast results in increased
rates of decapping (Schwartz & Parker, 1999). We find that the eIF4E-binding motif in Eap1 is
required for its role in Vts1-mediated degradation and thus the eIF4E/Eap1 interaction may
83
enhance decapping by destabilizing the translation initiation complex through displacement of
eIF4G from the mRNA.
Eap1 may also enhance decapping by facilitating the recruitment of decapping factors to
the 5' end of the mRNA. This is supported by systematic efforts that identified protein complexes
in S. cerevisiae, including an interaction between Eap1 and the decapping activator Dhh1 (Ho et
al., 2002). In this model the Eap1/eIF4E interaction would serve to recruit Dhh1 to the vicinity
of the cap thereby facilitating Dhh1-mediated decapping. While this model may not be
applicable to Vts1 target mRNAs, as Vts1-mediated decay of the GFP-SRE+ mRNA does not
involve Dhh1 (Rendl et al., 2008), it may apply to the degradation of COX17 mRNA, which is
Dhh1-dependent (Balagopal & Parker, 2009).
3.5.2 The Function of Eap1 in Yeast
Eap1 functions in a variety of biological processes including pseudohyphal growth
(Ibrahimo et al., 2006), the response to oxidative stress induced by exposure to diamide and
cadmium (Mascarenhas et al., 2008), the response to vesicular transport defects (Deloche et al.,
2004) and the regulation of rapamycin-induced inactivation of TOR-signaling (Cosentino et al.,
2000; Matsuo et al., 2005). In addition, polysome analysis suggests that EAP1 gene disruption
has little or no effect on bulk protein synthesis (Ibrahimo et al., 2006). Taken together with our
data, these results suggest that the role of Eap1 may be restricted to the regulation of specific
mRNAs under a variety of physiological conditions through the interaction of Eap1 with
sequence-specific RNA-binding proteins, such as Vts1 and perhaps Puf3. In addition, our data
indicate that Eap1 may exert its effect on target transcripts in these various conditions at the level
of transcript stability.
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3.5.3 The Role of eIF4E-Binding Proteins in mRNA Decay
Another eIF4E-binding protein, known as 4E-transporter (4E-T), has been shown to play
a role in transcript decay in mammalian cells (Ferraiuolo et al., 2005). Originally identified as
being responsible for transporting eIF4E into the nucleus (Dostie et al., 2000), subsequent
experiments showed that depletion of 4E-T from HeLa cells results in significant stabilization of
A+U-rich element (ARE)-containing mRNAs (Ferraiuolo et al., 2005). It is thought that 4E-T
may regulate the transition from translational repression to mRNA degradation through a
mechanism involving the localization of mRNAs to P-bodies, which are sites of mRNA storage
and decay (Andrei et al., 2005; Ferraiuolo et al., 2005). While the role of the 4E-T/eIF4E
interaction in ARE-mediated decay has not been assessed, indirect evidence suggests that 4E-T
may also stimulate decapping. As Eap1 and 4E-T are not homologous proteins, these data
suggest that eIF4E-binding proteins in other organisms might play a role in stimulating transcript
decapping through their interaction with eIF4E.
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Chapter 4
Conclusions and Future Directions
86
Chapter 4
Conclusions and Future Directions
4.1 Conclusions
In this thesis I have characterized the molecular mechanism of mRNA decay employed
by the sequence-specific RNA-binding protein Vts1 in S. cerevisiae. Using both in vivo reporter
mRNAs and a bona fide target of Vts1, I demonstrate that Vts1 stimulates mRNA degradation
through deadenylation mediated by the Ccr4-Pop2-Not deadenylase complex.
Immunoprecipitation experiments show that Vts1 interacts with Pop2, a component of the
deadenylase, in an RNA-independent manner. This suggests a model whereby Vts1 directly
recruits the Ccr4-Pop2-Not deadenylase complex to target mRNAs, resulting in mRNA decay
(Figure 4-1). Following deadenylation Vts1 target mRNAs are decapped and subsequently
degraded by the 5’-to-3’ exonuclease Xrn1. Deadenylation-dependent decapping followed by 5’-
to-3’ decay has been identified as the major mRNA decay pathway in yeast, and this work
demonstrates that Vts1 functions within this pathway.
In addition I have identified the eIF4E-binding protein Eap1 as a novel component of
Vts1-mediated decay. Eap1 influences the degradation of Vts1-target mRNAs at a step
downstream of deadenylation and consistent with this Vts1 target transcripts accumulate as
capped mRNAs in eap1Δ cells, indicating that Eap1 stimulates 5’ cap removal. Eap1 mediates an
interaction between Vts1 and eIF4E that is dependent on the Eap1 eIF4E-binding motif. This
motif is also required for Eap1-mediated destabilization of Vts1-target mRNAs. These data
suggest a model whereby Vts1 recruits Eap1 to facilitate mRNA decapping through an
interaction with the cap binding protein eIF4E (Figure 4-1). I propose that binding of Eap1 to
87
Figure 4-1 Model for Vts1-Mediated mRNA Decay
My work suggests that Vts1 physically recruits the Ccr4-Pop2-Not deadenylase complex to
induce mRNA decay by poly(A) tail shortening. After the poly(A) tail is removed, transcripts are
decapped and subsequently degraded by the 5’-to-3’ exonuclease Xrn1. In addition, Eap1 is
required for the decapping of Vts1 target transcripts. Eap1 mediates an indirect interaction
between Vts1 and eIF4E that is dependent on the Eap1 eIF4E-binding motif (YXXXXLΦ). My
data does not address whether Vts1 can simulateaously recruit both the Ccr4-Pop2-Not complex
and Eap1 to target mRNAs or whether Vts1 associates with these factors sequentially during
mRNA decay.
88
eIF4E increases the susceptibility of the mRNA to decapping following deadenylation. In
addition to regulating the stability of Vts1 target mRNAs, Eap1 also regulates the degradation of
Puf3 target transcripts. While Eap1 might play a role in the degradation of many unstable
mRNAs, it does not appear to be a component of the basic mRNA decay machinery.
4.2 Future Directions
4.2.1 Testing Direct Recruitment of the Deadenylase to Target mRNAs
As demonstrated in Chapter 2, Vts1 induces deadenylation-dependent mRNA decay and
interacts with the Ccr4-Pop2-Not deadenylase complex. Immunoprecipitation experiments
demonstrate an interaction between Vts1 and Pop2 (Figure 2-3), suggesting direct recruitment of
the deadenylase by Vts1 to target transcripts. A similar model of direct deadenylase recruitment
has been proposed for Smg-mediated mRNA decay in Drosophila (Semotok et al., 2005). As
described below, experiments aimed at confirming this model of Vts1 function would involve
understanding the molecular mechanisms that underlie the Vts1/deadenylase interaction and
generating a mutant version of Vts1 that blocks this interaction.
In vitro experiments could be used to determine whether Vts1 is in direct contact with a
component of the Ccr4-Pop2-Not deadenylase. I would generate radiolabelled versions of each
deadenylase component by in vitro translation using rabbit reticulocyte lysate and then test the
ability of each protein to bind Vts1 by glutathione S-transferase (GST) pulldown. Recombinant
Vts1 would be generated using the Baculovirus expression system since previous attempts to
generate recombinant Vts1 in bacteria have been unsuccessful. The labeled deadenylase
components would be incubated with purified recombinant GST-tagged Vts1 attached to
glutathione beads. Recombinant GST alone attached to glutathione beads would serve as a
negative control. After extensive washing, the presence of any associated proteins would be
89
assayed via SDS-PAGE. Since each component of the deadenylase is generated separately in
vitro, the association of any protein with Vts1-GST would suggest a direct interaction between
Vts1 and that subunit of the deadenylase complex. Binding assays would be repeated in the
presence of RNase A to ensure that any interactions observed were the result of direct protein-
protein interactions and not mediated by RNAs.
One concern with this method is that an observed interaction between recombinant Vts1
and a component of the deadenylase generated in rabbit reticulocyte lysate may not be direct but
instead mediated by endogenous proteins present in the rabbit reticulocyte lysate. However, one
would expect that if a rabbit reticulocyte protein were to mediate the interaction between Vts1
and the deadenylase, multiple deadenylase subunits would be immunoprecipitated. This
approach has been previously used to identify a direct interaction between recombinant Mpt5
and Pop2 generated in vitro using rabbit reticulocyte lysate (Goldstrohm et al., 2006). In this
study only Pop2 interacted with recombinant Mpt5, demonstrating that contaminant reticulocyte
proteins do not associate with the in vitro translated deadenylase subunits.
To further confirm a direct interaction between Vts1 and the Ccr4-Pop2-Not deadenylase,
I would generate an epitope tagged version of the deadenylase subunit, purified from bacteria or
Baculovirus, and test whether it binds to recombinant Vts1, generated in Baculovirus, by
immunoprecipitation. Again the immunoprecipitation would be repeated in the presence of
RNase A to ensure that the interaction was not mediated by RNA.
Having identified the component of the Ccr4-Pop2-Not deadenylase that directly interacts
with Vts1, I would next examine the stability of reporter mRNAs in cells deleted for the gene
encoding this factor. I expect that deletion of this factor would result in the stabilization of GFP-
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SRE+ mRNAs with long poly(A) tails. This would further support our model of Vts1-mediated
mRNA decay by direct deadenylase recruitment. The protein contacts between the subunits of
the Ccr4-Pop2-Not deadenylase have been mapped (Bai et al., 1999; Chen et al., 2001b; Liu et
al., 2001; Tucker et al., 2002). Some components are essential for deadenylation and integrity of
the complex (Ccr4, Pop2, and Not1), while others bind to the periphery of the complex and are
not required for the structural integrity or deadenylase activity of the core deadenylase (Not2-5,
Caf4, Caf16, Caf40 and Caf130). If it is found that Vts1 associates with a subunit of the
deadenylase that is essential to the integrity and catalytic activity of the complex, deletion of this
factor would result in the stabilization of GFP-SRE+ mRNAs with long poly(A) tails irrespective
of whether Vts1 was directly targeting the complex or not. Therefore this approach would only
support a direct recruitment model if Vts1 is found to interact with a nonessential component of
the deadenylase complex that does not normally exhibit a deadenylation defect when deleted.
After identifying the deadenylase component that directly binds Vts1, I would map the
region of Vts1 that is involved by repeating the GST-pulldown experiments using truncated
versions of Vts1-GST. Given our hypothesis that deadenylase recruitment is a conserved
mechanism of mRNA degradation for Smg family members, I would expect the Vts1
deadenylase interaction region (VDIR) to map to a region of conservation between Vts1 and
Drosophila Smg; for example, the SSR1 or SAM domain. However should the VDIR map to a
non-conserved region of Vts1, our approach would not be affected. After identifying the VDIR, I
would confirm the importance of this region in regulating mRNA stability by expressing a
version of Vts1 that carries a mutation within the VDIR that blocks deadenylase binding in vts1Δ
cells, and by examining the stability of reporter mRNAs by transcriptional pulse-chase. The goal
here would be to use a subtle VDIR mutation to reduce the likelihood that it would disrupt any
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other Vts1 interactions. If Vts1-mediated mRNA decay requires direct recruitment of the
deadenylase, mutation of the VD1R would result in a stabilization of GFP-SRE+ mRNA. As a
control I would confirm that mutation of the VDIR did not alter the general function of Vts1. As
a measure of Vts1 function I would express the Vts1VDIR- protein in yeast and test it for RNA
binding to a radiolabelled RNA by UV crosslinking. In addition I would test whether the
Vts1VDIR- protein was still capable of interacting with Eap1 by performing immunoprecipitation
experiments in yeast. The ability of Vts1VDIR- protein to bind RNA and Eap1 would indicate that
while mutation of the VDIR disrupted the interaction of Vts1 with the deadenylase complex, it
did not disrupt other Vts1 functions.
If Vts1 does not directly bind to the Ccr4-Pop2-Not deadenylase I would use structure-
function analysis to identify a region of Vts1 required for transcript deadenylation and then
perform experiments to identify proteins that interact with this region. To identify the region of
Vts1 required for mRNA decay I would generate epitope tagged Vts1 constructs that harbored
various truncations or deletions. I would then perform transcriptional pulse-chase experiments in
yeast to examine the stability and poly(A) tail length of reporter RNAs by Northern blot in vts1Δ
cells expressing these constructs. I would refine my mutational analysis until I had identified a
minimal region that was required for mRNA deadenylation, referred to as the Vts1
Deadenylation Region (VDR). This region could represent an interaction surface of an
unidentified factor required for Vts1 function that mediates an interaction between Vts1 and the
deadenylase complex. I would express an epitope tagged version of the VDR in yeast, purify it
and identify associated proteins by mass spectrometry. I would then examine the decay of
reporter RNAs in strains deleted for these VDR-binding proteins to find a component that, when
deleted, resulted in the stabilization of GFP-SRE+ mRNA and preservation of long poly(A) tails.
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Having identified a protein that interacts with the VDR and is required for the deadenylation of
GFP-SRE+ mRNA, I would perform immunoprecipitation experiments to determine whether this
protein mediates an interaction between Vts1 and the Ccr4-Pop2-Not deadenylase complex.
4.2.2 Testing Recruitment of Eap1 to Vts1 Target mRNAs
Our current model of Vts1-mediated decay suggests that Vts1 acts as a specificity
scaffold that binds to target mRNAs and then recruits other factors to induce transcript
degradation. I have shown that Eap1 is required for Vts1-mediated mRNA decay and that Eap1
mediates an interaction between Vts1 and eIF4E (Figure 3-5). These data suggest that Vts1
recruits Eap1 to stimulate the decapping of target mRNAs. As described below, experiments
aimed at confirming this model would involve understanding the Vts1/Eap1 interaction and
generating mutant proteins that block this interaction.
I would employ experiments similar to those described in the previous section to
determine whether Vts1 is in direct contact with Eap1. Briefly, I would test the ability of epitope
tagged recombinant Eap1, generated in bacteria, to bind epitope tagged recombinant Vts1,
generated in Baculovirus, by immunoprecipitation. To confirm that binding was the result of a
direct protein-protein interaction I would repeat the immunoprecipitation experiments in the
presence of RNase A.
After confirming a direct interaction between Vts1 and Eap1, I would map the region of
Eap1 that binds to Vts1. I would perform GST-pulldown experiments using Vts1-GST and
various truncated versions of Eap1 generated in rabbit reticulocyte lysate. This structure-function
analysis will allow me to identify the Eap1/Vts1 interaction region (EVIR) on Eap1. Again the
goal of this work would be to identify a subtle mutation within Vts1 that would block Eap1
93
binding while not affecting other interactions. If EVIR represents an Eap1 binding surface for
Vts1, this model predicts that mutation of the EVIR will prevent recruitment of Eap1 to Vts1
target mRNAs and result in their stabilization. Mutation of the EVIR would also disrupt the
interaction of Vts1 with eIF4E, which I have shown is mediated by Eap1. As a control I would
confirm that mutation of the EVIR did not alter the general function of Eap1. As a measure of
Eap1 function I would express the Eap1EVIR- protein in yeast and test whether it was still capable
of interacting with eIF4E by immunoprecipitation. I propose that Eap1 mediates an interaction
between Vts1 and eIF4E, therefore while mutation of the EVIR would disrupt the interaction of
Vts1 and eIF4E it should not prevent eIF4E binding to Eap1. I would also examine the
degradation of GFP-SRE+ mRNA in eap1Δ cells expressing Eap1EVIR- protein. I propose that
Eap1 is recruited by Vts1 to stimulate target mRNA decapping following deadenylation. Given
that mutation of the EVIR prevents the recruitment of Eap1 to Vts1 target mRNAs, GFP-SRE+
reporter mRNAs should accumulate as stable deadenylated transcripts that remain capped.
If the Eap1/Vts1 interaction is not direct, it is possible that the Eap1/Vts1 interaction is
being mediated by an unidentified protein. In this case I would use the structure/function analysis
described above to identify the region of Eap1 required for immunoprecipitation with Vts1. I
would express an epitope tagged version of this region in yeast, purify associated proteins and
identify them by mass spectrometry. I would then repeat the Eap1/Vts1 immunoprecipitation
experiments in strains deleted for the identified binding proteins in the hopes of discovering one
that when deleted disrupts the Eap1/Vts1 interaction. Deletion of this protein is also predicted to
stabilize GFP-SRE+ reporter mRNAs as deadenylated transcripts that remain capped.
94
4.2.3 Determining the Eap1 Mechanism of mRNA Degradation
4.2.3.1 Confirming Eap1 as an Activator of Decapping
As described in Chapter 3, Eap1 stimulates mRNA decay at a step downstream of
deadenylation. The accumulation of capped mRNAs in the eap1Δ strain indicates that Eap1
influences the rate of mRNA decapping (Figure 3-4). To further confirm the role of Eap1 in
decapping, GFP-SRE+ mRNA labeled with 32P in the cap structure would be incubated with
extracts derived from wild-type or eap1Δ cells. The resulting mRNA fragments would then be
resolved on a Thin Layer Chromatography (TLC) plate and visualized by autoradiography.
Decapping activity in wild-type extract produces radiolabelled m7GDP and is sensitive to EDTA
(Zhang et al., 1999). Failure to produce m7GDP in eap1Δ extracts would support a role for Eap1
in mRNA decapping. Extracts derived from dhh1Δ cells, a known activator of decapping
previously shown to exhibits a decapping defect in this assay (Zhang et al., 1999), would serve
as a control.
An alternative approach to confirm the role of Eap1 in decapping would test whether
mRNAs isolated from eap1Δ cells were susceptible to exonucleolytic degradation. Total RNA
would be isolated from eap1Δ cells expressing the GFP-SRE+ reporter and then treated with
recombinant purified Xrn1. Because Xrn1 has a strong preference for uncapped mRNAs,
protection against Xrn1-mediated degradation indicates the presence of an intact cap structure.
As a control equivalent samples will be treated with EDTA to inhibit Xrn1 activity (Fischer &
Weis, 2002). Total RNA from TetO7-DCP2 cells treated with DOX would serve as a control for
capped transcripts and total RNA from xrn1Δ cells would serve as a control for transcripts that
had lost the cap structure. Total RNA before and after Xrn1 treatment would be analyzed by
Northern blot and the percentage of full length capped to total full-length mRNA quantified. If
95
Eap1 has a role in stimulating decapping, most of the GFP-SRE+ mRNA isolated from eap1Δ
cells would be protected against exonucleolytic cleavage.
4.2.3.2 Does Eap1 Influence eIF4E Cap-Affinity?
A critical step in mRNA decapping is the removal of the translation initiation complex
(Schwartz & Parker, 1999; Vilela et al., 2000). We have proposed two possible models for Eap1
as a decapping activator: (1) the eIF4E-Eap1 interaction enhances decapping by destabilizing the
translation initiation complex and increasing accessibility of the cap to the decapping enzyme;
(2) Eap1 enhances decapping by recruiting the assembly of other decapping activators.
Given that the Eap1 eIF4E-binding motif is required for both mRNA decay (Figure 3-6)
and the interaction between Vts1 and eIF4E (Figure 3-5), it is possible that binding of Eap1
reduces the affinity of eIF4E for the 5’ cap structure and renders the mRNA more susceptible to
decapping. This model could be tested by measuring the affinity of eIF4E for the 5’ cap in the
presence of Eap1. UV-induced cross-linking assays have been used in previous measurements of
eIF4E cap-affinity (Haghighat & Sonenberg, 1997). This approach could be used to measure the
effect of Eap1 on eIF4E cap-affinity. This method was originally developed using recombinant
eIF4E from bacteria and recombinant eIF4G purified from Baculovirus infected insect cells
(Haghighat & Sonenberg, 1997). On its own the affinity of eIF4E for the cap is very weak,
however the addition of eIF4G dramatically enhances the cross-linking of eIF4E to capped
mRNA (Haghighat & Sonenberg, 1997). The addition of eIF4G would serve as a control
measurement for eIF4E-cap binding and allow us to measure changes in eIF4E-cap binding in
the presence of Eap1. Briefly, mRNA labeled with 32P in the cap structure would be incubated
with recombinant eIF4E and recombinant eIF4G (purified by previous methods), irradiated with
UV light, and RNase-digested. Labeled proteins are then analyzed by gel electrophoresis
96
followed by autoradiography. To test the effect of Eap1 on eIF4E cap-affinity, purified
recombinant Eap1, generated in bacteria, would be preincubated with eIF4E prior to adding
eIF4G and UV cross-linking. A decrease in eIF4E/eIF4G labeling in the presence of Eap1 would
indicate that Eap1 reduced the eIF4E cap-affinity. Prior to UV cross-linking eIF4E could be
preincubated with increasing amounts of Eap1 to illustrate a dose-dependent reduction in eIF4E
labeling. As a control recombinant Eap1 harboring mutations in the eIF4E-binding motif is not
expected to bind eIF4E and therefore not reduce the eIF4E cap-affinity.
Surface plasmon resonance (SPR) provides a more quantitative approach to the study of
eIF4E cap-affinity (Ptushkina et al., 1999). This technique measures the release kinetics for
eIF4E bound to a capped and biotinylated RNA that has been coupled to streptavidin-coated SPR
chips. By passing additional purified proteins over the chip, their effect on eIF4E cap-affinity can
be measured as a change in the eIF4E off-rate from the RNA bound to the chip. A reduction of
eIF4E cap-affinity in the presence of Eap1 would support our model of Eap1-mediated mRNA
decapping. As a control passing purified eIF4G over the chip should increase the eIF4E cap-
affinity. Since eIF4G enhances eIF4E binding to the mRNA cap (Haghighat & Sonenberg,
1997), an increase in eIF4E-cap affinity in the presence of eIF4G would validate this approach as
a method for measuring changes in eIF4E cap binding.
4.2.3.3 Does Eap1 Recruit Decapping Activators?
A number of factors that enhance mRNA decapping have been identified in S. cerevisiae,
however their mechanism of action remains unknown [reviewed in (Coller & Parker, 2004)].
Systematic identification of yeast protein complexes by mass spectrometry revealed an
interaction between Eap1 and the decapping activator Dhh1 (Ho et al., 2002). This suggests that
Eap1 may stimulate mRNA decay by recruiting Dhh1 to target transcripts. Previously we
97
demonstrated that Dhh1 was not required for Vts1-mediated mRNA decay (Rendl et al., 2008).
However this study only examined the role of Dhh1 in the decay of reporter mRNAs. It is
therefore possible that Dhh1 could be involved in the degradation of in vivo transcripts regulated
by Eap1, such as COX17 (discussed below).
Elucidating the mechanism of Eap1-mediated mRNA decay may depend on the
identification of more targets. I have shown that Eap1 is required for the regulation targets that
are both Vts1-dependent and Vts1-independent. It is unclear whether Eap1 uses a common
mechanism or may have multiple mechanisms of affecting transcript decay that are specified by
interactions with other RNA-binding proteins. As more targets of Eap1 regulation are identified
(see below) we will be able to test whether all Eap1 targets are regulated by a common
mechanism, for example Dhh1.
4.2.4 Identification and Characterization of Eap1 Target mRNAs
The identification of Eap1 mRNA targets would assist in understanding the role and
specificity of Eap1 in regulating mRNA decay. To identify putative Eap1 targets I would employ
a pulldown/microarray approach. Microarrays spotted with DNA sequences of yeast ORFs
would be hybridized with complementary DNA prepared from total RNA and RNA copurified
with epitope-tagged Eap1. I would identify Eap1 targets as RNAs enriched in the
immunoprecipitated sample relative to the total RNA control. A complimentary approach to
identify targets that were regulated by Eap1 at the level of mRNA stability would use microarray
technology to identify genes that were strongly upregulated in the eap1Δ strain compared to
wild-type. If Eap1 were involved in controlling the expression of certain genes, for example by
stimulating mRNA decay, I expect that expression of these genes to be elevated in the absence of
Eap1. However, mRNAs identified by this approach would exhibit elevated mRNA levels as a
98
result of both direct and indirect effects of Eap1 on mRNA stability. Therefore an alternative
approach would be to measure the stability of select mRNAs identified by the
pulldown/microarray approach in eap1Δ cells using transcriptional inhibitors. Transcripts with
longer half-lives in eap1Δ cells would represent candidate mRNAs regulated by Eap1 at the level
of transcript stability.
I have evidence that Eap1 regulates the stability of different groups of mRNAs, both
Vts1-dependent and Vts1-independent. After identifying a list of potential Eap1 targets, I would
search the candidates for stem-loops that match the SRE consensus. Recently ~40 yeast RNA-
binding proteins, including Vts1 and their associated mRNAs were identified by
pulldown/microarray(Hogan et al., 2008). This study identified a group of potential Vts1 targets
enriched for a motif unrelated to the SRE. I would also cross-reference my list of potential Eap1
targets to the newly identified Vts1 mRNA targets and search for common transcripts. mRNAs
in this group represent transcripts that are regulated by Vts1 and Eap1. I would then go on to
characterize the mechanism of mRNA decay for these transcripts by standard techniques.
Genes that do not contain predicted SREs represent transcripts that may be regulated by
Eap1 in a Vts1-independent manner. Eap1 does not appear to be a general regulator of mRNA
decay, suggesting that there is some element of specificity to the targeting of Eap1 to mRNAs. I
would cross-reference my list of Vts1-independent Eap1 targets to the list of newly identified
mRNA targets of ~40 yeast RNA-binding proteins (Hogan et al., 2008). Comparing these lists
may highlight a group of Eap1 targets that are common to one or more RNA-binding proteins,
Puf3 for example. I could then perform experiments similar to those outlined in Chapter 3 to
determine whether Eap1 interacts with the identified RNA-binding proteins and characterize the
mechanism of Eap1-mediated mRNA decay for these targets.
99
4.2.4.1 The Role of Eap1 in PUF Target Regulation
I propose that Eap1 is recruited to SRE-containing mRNAs through an association with
Vts1. I have shown that COX17 mRNA is stabilized in an eap1Δ strain (Figure 3-7). Given that
COX17 mRNA stability is regulated by Puf3, I hypothesize that Puf3 might recruit Eap1 to
COX17 mRNA. I would test the Puf3/Eap1 interaction in yeast by performing
immunoprecipitation experiments using epitope tagged proteins. I would also use
structure/function analysis (as described previously) to identify the region on Puf3 that is
required for Eap1 binding, the Puf3/Eap1 binding region (PEBR). After confirming that deletion
of the PEBR does not disrupt Puf3 RNA binding, I would measure the stability of COX17
mRNA in puf3Δ cells expressing a version of Puf3 in which the PEBR was removed (Puf3PEBR-).
Stabilization of COX17 mRNA when Puf3PEBR- is expressed in puf3Δ cells would suggest that
Eap1 recruitment by Puf3 is required for COX17 decay.
I have shown that the Eap1 eIF4E-binding motif is required for Vts1-mediated mRNA
decay (Figure 3-6). As a first step to test whether all Eap1 targets are regulated by a common
mechanism I would determine whether the Eap1 eIF4E-binding motif was required for COX17
mRNA degradation. I have shown that COX17 mRNA is stabilized in eap1Δ cells (Figure 3-7). I
would measure the stability of COX17 mRNA in eap1Δ cells expressing either wild-type Eap1 or
a version of Eap1 in which the eIF4E-binding motif has been mutated (Eap1mt). Failure of
Eap1mt to rescue COX17 mRNA degradation in eap1Δ cells would indicate that Eap1 employs a
common mechanism to degrade target mRNAs, both Vts1-dependent and Vts1-independent.
As mentioned previously, Eap1 may stimulate mRNA decay by recruiting the decapping
activator Dhh1 to target transcripts. In Chapter 3, I demonstrate that COX17 mRNA has an
100
increased stability in eap1Δ cells as compared to wild-type (Figure 3-7). Interestingly, the
stability of COX17 is increased in a dhh1Δ strain (Balagopal & Parker, 2009). The regulation of
COX17 stability by Eap1 and Dhh1 suggests that Eap1 and Dhh1 may act together to induce
transcript decay. I have shown that COX17 stability is not altered in a vts1Δ strain (Figure 3-7).
Sequence analysis of the transcript failed to identify any potential SREs, further confirming that
COX17 is not a target of Vts1-mediated regulation. Clearly Eap1 has a role in mRNA decay,
although its function may be limited to specific mRNAs.
To test whether Eap1 recruits Dhh1 to COX17 mRNA I would perform structure/function
analysis (similar to previously proposed experiments) to identify the region on Eap1 required for
Dhh1 binding. After identifying the Eap1/Dhh1 binding region (EDBR) I would delete this
region in Eap1 (Eap1EDBR-) and determine what effect deletion of the EDBR had on COX17
mRNA stability. If the stability of COX17 mRNA in dhh1Δ cells is similar to the stability in
eap1Δ cells expressing Eap1EDBR- it suggests that Eap1 is regulating COX17 mRNA degradation
by recruiting Dhh1.
Given our data that Eap1 regulates COX17, a Puf3 target, we may identify other Eap1
targets that are regulated by PUF proteins (discussed above). Enrichment for PUF targets would
suggest that Eap1 is recruited to these transcripts via binding to PUF proteins. In this case I
would first confirm the Eap1-PUF interactions (using previously described methods). I would
also test whether Dhh1 recruitment or the Eap1 eIF4E-binding motif was required for the
degradation of these PUF target mRNAs (using previously described methods). A stabilization of
PUF targets dependent on Dhh1 recruitment and/or the Eap1 eIF4E-binding motif would suggest
101
that Eap1 employs a conserved mechanism to induce mRNA decay and is recruited to specific
transcripts by its association with other RNA-binding proteins.
The experiments described above can be used to determine the mechanism and specificity
of Eap1-mediated mRNA decay. They can also further our understanding of mRNA regulation
by Vts1 and may highlight a mechanism of mRNA decay that is conserved within the Smg
family.
102
References
Altmann M, Schmitz N, Berset C, Trachsel H. 1997. A novel inhibitor of cap-dependent translation initiation in yeast: p20 competes with eIF4G for binding to eIF4E. EMBO J 16:1114-1121.
Amrani N, Ganesan R, Kervestin S, Mangus DA, Ghosh S, Jacobson A. 2004. A faux 3'-UTR promotes aberrant termination and triggers nonsense-mediated mRNA decay. Nature 432:112-118.
Anders KR, Grimson A, Anderson P. 2003. SMG-5, required for C.elegans nonsense-mediated mRNA decay, associates with SMG-2 and protein phosphatase 2A. EMBO J 22:641-650.
Anderson JS, Parker RP. 1998. The 3' to 5' degradation of yeast mRNAs is a general mechanism for mRNA turnover that requires the SKI2 DEVH box protein and 3' to 5' exonucleases of the exosome complex. EMBO J 17:1497-1506.
Anderson P, Kedersha N. 2006. RNA granules. J Cell Biol 172:803-808.
Andrei MA, Ingelfinger D, Heintzmann R, Achsel T, Rivera-Pomar R, Luhrmann R. 2005. A role for eIF4E and eIF4E-transporter in targeting mRNPs to mammalian processing bodies. RNA 11:717-727.
Aviv T, Lin Z, Ben-Ari G, Smibert CA, Sicheri F. 2006. Sequence-specific recognition of RNA hairpins by the SAM domain of Vts1p. Nat Struct Mol Biol 13:168-176.
Aviv T, Lin Z, Lau S, Rendl LM, Sicheri F, Smibert CA. 2003. The RNA-binding SAM domain of Smaug defines a new family of post-transcriptional regulators. Nat Struct Biol 10:614-621.
Badis G, Saveanu C, Fromont-Racine M, Jacquier A. 2004. Targeted mRNA degradation by deadenylation-independent decapping. Mol Cell 15:5-15.
Baez MV, Boccaccio GL. 2005. Mammalian Smaug is a translational repressor that forms cytoplasmic foci similar to stress granules. J Biol Chem 280:43131-43140.
Bai Y, Salvadore C, Chiang YC, Collart MA, Liu HY, Denis CL. 1999. The CCR4 and CAF1 proteins of the CCR4-NOT complex are physically and functionally separated from NOT2, NOT4, and NOT5. Mol Cell Biol 19:6642-6651.
Baker KE, Parker R. 2004. Nonsense-mediated mRNA decay: terminating erroneous gene expression. Curr Opin Cell Biol 16:293-299.
Balagopal V, Parker R. 2009. Stm1 Modulates mRNA Decay and Dhh1 Function in Saccharomyces cerevisiae. Genetics 181:93-103.
103
Barbee SA, Estes PS, Cziko AM, Hillebrand J, Luedeman RA, Coller JM, Johnson N, Howlett IC, Geng C, Ueda R, Brand AH, Newbury SF, Wilhelm JE, Levine RB, Nakamura A, Parker R, Ramaswami M. 2006. Staufen- and FMRP-containing neuronal RNPs are structurally and functionally related to somatic P bodies. Neuron 52:997-1009.
Bashirullah A, Cooperstock RL, Lipshitz HD. 2001. Spatial and temporal control of RNA stability. Proc Natl Acad Sci U S A 98:7025-7028.
Bashirullah A, Halsell SR, Cooperstock RL, Kloc M, Karaiskakis A, Fisher WW, Fu W, Hamilton JK, Etkin LD, Lipshitz HD. 1999. Joint action of two RNA degradation pathways controls the timing of maternal transcript elimination at the midblastula transition in Drosophila melanogaster. EMBO J 18:2610-2620.
Beelman CA, Parker R. 1995. Degradation of mRNA in eukaryotes. Cell 81:179-183.
Behm-Ansmant I, Rehwinkel J, Doerks T, Stark A, Bork P, Izaurralde E. 2006. mRNA degradation by miRNAs and GW182 requires both CCR4:NOT deadenylase and DCP1:DCP2 decapping complexes. Genes Dev 20:1885-1898.
Bergsten S, Gavis E. 1999. Role for mRNA localization in translational activation but not spatial restriction of nanos RNA. Development 126:659-669.
Binder R, Horowitz JA, Basilion JP, Koeller DM, Klausner RD, Harford JB. 1994. Evidence that the pathway of transferrin receptor mRNA degradation involves an endonucleolytic cleavage within the 3' UTR and does not involve poly(A) tail shortening. EMBO J 13:1969-1980.
Bonnerot C, Boeck R, Lapeyre B. 2000. The two proteins Pat1p (Mrt1p) and Spb8p interact in vivo, are required for mRNA decay, and are functionally linked to Pab1p. Mol Cell Biol 20:5939-5946.
Bouveret E, Rigaut G, Shevchenko A, Wilm M, Seraphin B. 2000. A Sm-like protein complex that participates in mRNA degradation. EMBO J 19:1661-1671.
Brachmann CB, Davies A, Cost GJ, Caputo E, Li J, Hieter P, Boeke JD. 1998. Designer deletion strains derived from Saccharomyces cerevisiae S288C: a useful set of strains and plasmids for PCR-mediated gene disruption and other applications. Yeast 14:115-132.
Brengues M, Teixeira D, Parker R. 2005. Movement of eukaryotic mRNAs between polysomes and cytoplasmic processing bodies. Science 310:486-489.
Briand JF, Navarro F, Gadal O, Thuriaux P. 2001. Cross talk between tRNA and rRNA synthesis in Saccharomyces cerevisiae. Mol Cell Biol 21:189-195.
Brock ML, Shapiro DJ. 1983. Estrogen stabilizes vitellogenin mRNA against cytoplasmic degradation. Cell 34:207-214.
104
Brown CE, Sachs AB. 1998. Poly(A) tail length control in Saccharomyces cerevisiae occurs by message-specific deadenylation. Mol Cell Biol 18:6548-6559.
Butler JS. 2002. The yin and yang of the exosome. Trends Cell Biol 12:90-96.
Cali BM, Kuchma SL, Latham J, Anderson P. 1999. smg-7 is required for mRNA surveillance in Caenorhabditis elegans. Genetics 151:605-616.
Caponigro G, Parker R. 1996a. Mechanisms and control of mRNA turnover in Saccharomyces cerevisiae. Microbiol Rev 60:233-249.
Caponigro G, Parker R. 1996b. mRNA turnover in yeast promoted by the MATalpha1 instability element. Nucleic Acids Res 24:4304-4312.
Chang TC, Yamashita A, Chen CY, Yamashita Y, Zhu W, Durdan S, Kahvejian A, Sonenberg N, Shyu AB. 2004. UNR, a new partner of poly(A)-binding protein, plays a key role in translationally coupled mRNA turnover mediated by the c-fos major coding-region determinant. Genes Dev 18:2010-2023.
Chang YF, Imam JS, Wilkinson MF. 2007. The nonsense-mediated decay RNA surveillance pathway. Annu Rev Biochem 76:51-74.
Chen CY, Gherzi R, Ong SE, Chan EL, Raijmakers R, Pruijn GJ, Stoecklin G, Moroni C, Mann M, Karin M. 2001a. AU binding proteins recruit the exosome to degrade ARE-containing mRNAs. Cell 107:451-464.
Chen J, Chiang YC, Denis CL. 2002. CCR4, a 3'-5' poly(A) RNA and ssDNA exonuclease, is the catalytic component of the cytoplasmic deadenylase. EMBO J 21:1414-1426.
Chen J, Rappsilber J, Chiang YC, Russell P, Mann M, Denis CL. 2001b. Purification and characterization of the 1.0 MDa CCR4-NOT complex identifies two novel components of the complex. J Mol Biol 314:683-694.
Chernokalskaya E, Dompenciel R, Schoenberg DR. 1997. Cleavage properties of an estrogen-regulated polysomal ribonuclease involved in the destabilization of albumin mRNA. Nucleic Acids Res 25:735-742.
Chicoine J, Benoit P, Gamberi C, Paliouras M, Simonelig M, Lasko P. 2007. Bicaudal-C recruits CCR4-NOT deadenylase to target mRNAs and regulates oogenesis, cytoskeletal organization, and its own expression. Dev Cell 13:691-704.
Chiu SY, Serin G, Ohara O, Maquat LE. 2003. Characterization of human Smg5/7a: a protein with similarities to Caenorhabditis elegans SMG5 and SMG7 that functions in the dephosphorylation of Upf1. RNA 9:77-87.
Clark IE, Wyckoff D, Gavis ER. 2000. Synthesis of the posterior determinant Nanos is spatially restricted by a novel cotranslational regulatory mechanism. Curr Biol 10:1311-1314.
105
Coller J, Parker R. 2004. Eukaryotic mRNA decapping. Annu Rev Biochem 73:861-890.
Coller J, Parker R. 2005. General translational repression by activators of mRNA decapping. Cell 122:875-886.
Coller JM, Tucker M, Sheth U, Valencia-Sanchez MA, Parker R. 2001. The DEAD box helicase, Dhh1p, functions in mRNA decapping and interacts with both the decapping and deadenylase complexes. RNA 7:1717-1727.
Cosentino GP, Schmelzle T, Haghighat A, Helliwell SB, Hall MN, Sonenberg N. 2000. Eap1p, a novel eukaryotic translation initiation factor 4E-associated protein in Saccharomyces cerevisiae. Mol Cell Biol 20:4604-4613.
Cougot N, Babajko S, Seraphin B. 2004. Cytoplasmic foci are sites of mRNA decay in human cells. J Cell Biol 165:31-40.
Crucs S, Chatterjee S, Gavis ER. 2000. Overlapping but distinct RNA elements control repression and activation of nanos translation. Molecular Cell 5:457-467.
Cui Y, Hagan KW, Zhang S, Peltz SW. 1995. Identification and characterization of genes that are required for the accelerated degradation of mRNAs containing a premature translational termination codon. Genes Dev 9:423-436.
Cunningham KS, Dodson RE, Nagel MA, Shapiro DJ, Schoenberg DR. 2000. Vigilin binding selectively inhibits cleavage of the vitellogenin mRNA 3'-untranslated region by the mRNA endonuclease polysomal ribonuclease 1. Proc Natl Acad Sci U S A 97:12498-12502.
Czaplinski K, Ruiz-Echevarria MJ, Paushkin SV, Han X, Weng Y, Perlick HA, Dietz HC, Ter-Avanesyan MD, Peltz SW. 1998. The surveillance complex interacts with the translation release factors to enhance termination and degrade aberrant mRNAs. Genes Dev 12:1665-1677.
Dahanukar A, Walker JA, Wharton RP. 1999. Smaug, a novel RNA-binding protein that operates a translational switch in Drosophila. Mol Cell 4:209-218.
Dahanukar A, Wharton RP. 1996. The nanos gradient in Drosophila embryos is generated by translational regulation. Genes & Dev 10:2610-2620.
Daugeron MC, Mauxion F, Seraphin B. 2001. The yeast POP2 gene encodes a nuclease involved in mRNA deadenylation. Nucleic Acids Res 29:2448-2455.
de la Cruz J, Iost I, Kressler D, Linder P. 1997. The p20 and Ded1 proteins have antagonistic roles in eIF4E-dependent translation in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 94:5201-5206.
Decker CJ, Parker R. 1993. A turnover pathway for both stable and unstable mRNAs in yeast: evidence for a requirement for deadenylation. Genes Dev 7:1632-1643.
106
Decker CJ, Teixeira D, Parker R. 2007. Edc3p and a glutamine/asparagine-rich domain of Lsm4p function in processing body assembly in Saccharomyces cerevisiae. J Cell Biol 179:437-449.
Dehlin E, Wormington M, Korner CG, Wahle E. 2000. Cap-dependent deadenylation of mRNA. EMBO J 19:1079-1086.
Deloche O, de la Cruz J, Kressler D, Doere M, Linder P. 2004. A membrane transport defect leads to a rapid attenuation of translation initiation in Saccharomyces cerevisiae. Mol Cell 13:357-366.
Denis CL, Chen J. 2003. The CCR4-NOT complex plays diverse roles in mRNA metabolism. Prog Nucleic Acid Re 73:221-250.
Denning G, Jamieson L, Maquat LE, Thompson EA, Fields AP. 2001. Cloning of a novel phosphatidylinositol kinase-related kinase: characterization of the human SMG-1 RNA surveillance protein. J Biol Chem 276:22709-22714.
Deshmukh MV, Jones BN, Quang-Dang DU, Flinders J, Floor SN, Kim C, Jemielity J, Kalek M, Darzynkiewicz E, Gross JD. 2008. mRNA decapping is promoted by an RNA-binding channel in Dcp2. Mol Cell 29:324-336.
Deutschbauer AM, Williams RM, Chu AM, Davis RW. 2002. Parallel phenotypic analysis of sporulation and postgermination growth in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 99:15530-15535.
Dilcher M, Kohler B, von Mollard GF. 2001. Genetic interactions with the yeast Q-SNARE VTI1 reveal novel functions for the R-SNARE YKT6. J Biol Chem 276:34537-34544.
Ding D, Parkhurst SM, Halsell SR, Lipshitz HD. 1993. Dynamic Hsp83 RNA localization during Drosophila oogenesis and embryogenesis. Mol Cell Biol 13:3773-3781.
Dlakic M. 2000. Functionally unrelated signalling proteins contain a fold similar to Mg2+-dependent endonucleases. Trends Biochem Sci 25:272-273.
Dodson RE, Shapiro DJ. 1997. Vigilin, a ubiquitous protein with 14 K homology domains, is the estrogen-inducible vitellogenin mRNA 3'-untranslated region-binding protein. J Biol Chem 272:12249-12252.
Doma MK, Parker R. 2006. Endonucleolytic cleavage of eukaryotic mRNAs with stalls in translation elongation. Nature 440:561-564.
Doma MK, Parker R. 2007. RNA quality control in eukaryotes. Cell 131:660-668.
Dostie J, Ferraiuolo M, Pause A, Adam SA, Sonenberg N. 2000. A novel shuttling protein, 4E-T, mediates the nuclear import of the mRNA 5' cap-binding protein, eIF4E. EMBO J 19:3142-3156.
107
Eulalio A, Behm-Ansmant I, Izaurralde E. 2007a. P bodies: at the crossroads of post-transcriptional pathways. Nat Rev Mol Cell Biol 8:9-22.
Eulalio A, Behm-Ansmant I, Schweizer D, Izaurralde E. 2007b. P-body formation is a consequence, not the cause, of RNA-mediated gene silencing. Mol Cell Biol 27:3970-3981.
Ferraiuolo MA, Basak S, Dostie J, Murray EL, Schoenberg DR, Sonenberg N. 2005. A role for the eIF4E-binding protein 4E-T in P-body formation and mRNA decay. J Cell Biol 170:913-924.
Fischer N, Weis K. 2002. The DEAD box protein Dhh1 stimulates the decapping enzyme Dcp1. EMBO J 21:2788-2797.
Franks TM, Lykke-Andersen J. 2007. TTP and BRF proteins nucleate processing body formation to silence mRNAs with AU-rich elements. Genes Dev 21:719-735.
Franks TM, Lykke-Andersen J. 2008. The control of mRNA decapping and P-body formation. Mol Cell 32:605-615.
Frischmeyer PA, van Hoof A, O'Donnell K, Guerrerio AL, Parker R, Dietz HC. 2002. An mRNA surveillance mechanism that eliminates transcripts lacking termination codons. Science 295:2258-2261.
Fukuhara N, Ebert J, Unterholzner L, Lindner D, Izaurralde E, Conti E. 2005. SMG7 is a 14-3-3-like adaptor in the nonsense-mediated mRNA decay pathway. Mol Cell 17:537-547.
Gao M, Fritz DT, Ford LP, Wilusz J. 2000. Interaction between a poly(A)-specific ribonuclease and the 5' cap influences mRNA deadenylation rates in vitro. Mol Cell 5:479-488.
Gao M, Wilusz CJ, Peltz SW, Wilusz J. 2001. A novel mRNA-decapping activity in HeLa cytoplasmic extracts is regulated by AU-rich elements. EMBO J 20:1134-1143.
Garneau NL, Wilusz J, Wilusz CJ. 2007. The highways and byways of mRNA decay. Nat Rev Mol Cell Biol 8:113-126.
Gatfield D, Izaurralde E. 2004. Nonsense-mediated messenger RNA decay is initiated by endonucleolytic cleavage in Drosophila. Nature 429:575-578.
Gatfield D, Unterholzner L, Ciccarelli FD, Bork P, Izaurralde E. 2003. Nonsense-mediated mRNA decay in Drosophila: at the intersection of the yeast and mammalian pathways. EMBO J 22:3960-3970.
Gavis ER, Curtis D, Lehmann R. 1996a. Identification of cis-acting sequences that control nanos RNA localization. Dev Biol 176:36-50.
Gavis ER, Lehmann R. 1994. Translational regulation of nanos by RNA localization. Nature 369:315-318.
108
Gavis ER, Lunsford L, Bergsten SE, Lehmann R. 1996b. A conserved 90 nucleotide element mediates translational repression of nanos RNA. Development 122:2791-2800.
Gerber AP, Herschlag D, Brown PO. 2004. Extensive association of functionally and cytotopically related mRNAs with Puf family RNA-binding proteins in yeast. PLoS Biol 2:E79.
Gingras A-C, Raught B, Sonenberg N. 1999. eIF4 INITIATION FACTORS: Effectors of mRNA Recruitment to Ribosomes and Regulators of Translation. Annu Rev Biochem 68:913-963.
Giraldez AJ, Mishima Y, Rihel J, Grocock RJ, Van Dongen S, Inoue K, Enright AJ, Schier AF. 2006. Zebrafish MiR-430 promotes deadenylation and clearance of maternal mRNAs. Science 312:75-79.
Goldstrohm AC, Hook BA, Seay DJ, Wickens M. 2006. PUF proteins bind Pop2p to regulate messenger RNAs. Nat Struct Mol Biol 13:533-539.
Goldstrohm AC, Seay DJ, Hook BA, Wickens M. 2007. PUF protein-mediated deadenylation is catalyzed by Ccr4p. J Biol Chem 282:109-114.
Green JB, Gardner CD, Wharton RP, Aggarwal AK. 2003. RNA recognition via the SAM domain of Smaug. Mol Cell 11:1537-1548.
Gregory RI, Yan KP, Amuthan G, Chendrimada T, Doratotaj B, Cooch N, Shiekhattar R. 2004. The Microprocessor complex mediates the genesis of microRNAs. Nature 432:235-240.
Grimson A, O'Connor S, Newman CL, Anderson P. 2004. SMG-1 is a phosphatidylinositol kinase-related protein kinase required for nonsense-mediated mRNA Decay in Caenorhabditis elegans. Mol Cell Biol 24:7483-7490.
Grosset C, Chen CY, Xu N, Sonenberg N, Jacquemin-Sablon H, Shyu AB. 2000. A mechanism for translationally coupled mRNA turnover: interaction between the poly(A) tail and a c-fos RNA coding determinant via a protein complex. Cell 103:29-40.
Haghighat A, Sonenberg N. 1997. eIF4G dramatically enhances the binding of eIF4E to the mRNA 5'-cap structure. J Biol Chem 272:21677-21680.
Hatfield L, Beelman CA, Stevens A, Parker R. 1996. Mutations in trans-acting factors affecting mRNA decapping in Saccharomyces cerevisiae. Mol Cell Biol 16:5830-5838.
He F, Brown AH, Jacobson A. 1997. Upf1p, Nmd2p, and Upf3p are interacting components of the yeast nonsense-mediated mRNA decay pathway. Mol Cell Biol 17:1580-1594.
He F, Li X, Spatrick P, Casillo R, Dong S, Jacobson A. 2003. Genome-wide analysis of mRNAs regulated by the nonsense-mediated and 5' to 3' mRNA decay pathways in yeast. Mol Cell 12:1439-1452.
109
Ho Y, Gruhler A, Heilbut A, Bader GD, Moore L, Adams SL, Millar A, Taylor P, Bennett K, Boutilier K, Yang L, Wolting C, Donaldson I, Schandorff S, Shewnarane J, Vo M, Taggart J, Goudreault M, Muskat B, Alfarano C, Dewar D, Lin Z, Michalickova K, Willems AR, Sassi H, Nielsen PA, Rasmussen KJ, Andersen JR, Johansen LE, Hansen LH, Jespersen H, Podtelejnikov A, Nielsen E, Crawford J, Poulsen V, Sorensen BD, Matthiesen J, Hendrickson RC, Gleeson F, Pawson T, Moran MF, Durocher D, Mann M, Hogue CW, Figeys D, Tyers M. 2002. Systematic identification of protein complexes in Saccharomyces cerevisiae by mass spectrometry. Nature 415:180-183.
Hodgkin J, Papp A, Pulak R, Ambros V, Anderson P. 1989. A new kind of informational suppression in the nematode Caenorhabditis elegans. Genetics 123:301-313.
Hogan DJ, Riordan DP, Gerber AP, Herschlag D, Brown PO. 2008. Diverse RNA-binding proteins interact with functionally related sets of RNAs, suggesting an extensive regulatory system. PLoS Biol 6:e255.
Hook BA, Goldstrohm AC, Seay DJ, Wickens M. 2007. Two yeast PUF proteins negatively regulate a single mRNA. J Biol Chem 282:15430-15438.
Houshmandi SS, Olivas WM. 2005. Yeast Puf3 mutants reveal the complexity of Puf-RNA binding and identify a loop required for regulation of mRNA decay. RNA 11:1655-1666.
Hsu CL, Stevens A. 1993. Yeast cells lacking 5'-->3' exoribonuclease 1 contain mRNA species that are poly(A) deficient and partially lack the 5' cap structure. Mol Cell Biol 13:4826-4835.
Ibrahimo S, Holmes LE, Ashe MP. 2006. Regulation of translation initiation by the yeast eIF4E binding proteins is required for the pseudohyphal response. Yeast 23:1075-1088.
Jackson JS, Jr., Houshmandi SS, Lopez Leban F, Olivas WM. 2004. Recruitment of the Puf3 protein to its mRNA target for regulation of mRNA decay in yeast. RNA 10:1625-1636.
Jacobson A, Peltz SW. 1996. Interrelationships of the pathways of mRNA decay and translation in eukaryotic cells. Annu Rev Biochem 65:693-739.
Jain RA, Gavis ER. 2008. The Drosophila hnRNP M homolog Rumpelstiltskin regulates nanos mRNA localization. Development 135:973-982.
Jing Q, Huang S, Guth S, Zarubin T, Motoyama A, Chen J, Di Padova F, Lin SC, Gram H, Han J. 2005. Involvement of microRNA in AU-rich element-mediated mRNA instability. Cell 120:623-634.
Johnson PE, Donaldson LW. 2006. RNA recognition by the Vts1p SAM domain. Nat Struct Mol Biol 13:177-178.
Kadyrova LY, Habara Y, Lee TH, Wharton RP. 2007. Translational control of maternal Cyclin B mRNA by Nanos in the Drosophila germline. Development 134:1519-1527.
110
Kalifa Y, Huang T, Rosen LN, Chatterjee S, Gavis ER. 2006. Glorund, a Drosophila hnRNP F/H homolog, is an ovarian repressor of nanos translation. Dev Cell 10:291-301.
Kashima I, Yamashita A, Izumi N, Kataoka N, Morishita R, Hoshino S, Ohno M, Dreyfuss G, Ohno S. 2006. Binding of a novel SMG-1-Upf1-eRF1-eRF3 complex (SURF) to the exon junction complex triggers Upf1 phosphorylation and nonsense-mediated mRNA decay. Genes Dev 20:355-367.
Klausner RD, Rouault TA, Harford JB. 1993. Regulating the fate of mRNA: the control of cellular iron metabolism. Cell 72:19-28.
Kong J, Liebhaber SA. 2007. A cell type-restricted mRNA surveillance pathway triggered by ribosome extension into the 3' untranslated region. Nat Struct Mol Biol 14:670-676.
Kshirsagar M, Parker R. 2004. Identification of Edc3p as an enhancer of mRNA decapping in Saccharomyces cerevisiae. Genetics 166:729-739.
LaGrandeur TE, Parker R. 1998. Isolation and characterization of Dcp1p, the yeast mRNA decapping enzyme. EMBO J 17:1487-1496.
Lai WS, Carballo E, Thorn JM, Kennington EA, Blackshear PJ. 2000. Interactions of CCCH zinc finger proteins with mRNA. Binding of tristetraprolin-related zinc finger proteins to Au-rich elements and destabilization of mRNA. J Biol Chem 275:17827-17837.
Lai WS, Kennington EA, Blackshear PJ. 2003. Tristetraprolin and its family members can promote the cell-free deadenylation of AU-rich element-containing mRNAs by poly(A) ribonuclease. Mol Cell Biol 23:3798-3812.
Larimer FW, Stevens A. 1990. Disruption of the gene XRN1, coding for a 5'----3' exoribonuclease, restricts yeast cell growth. Gene 95:85-90.
Leeds P, Peltz SW, Jacobson A, Culbertson MR. 1991. The product of the yeast UPF1 gene is required for rapid turnover of mRNAs containing a premature translational termination codon. Genes Dev 5:2303-2314.
Lingel A, Sattler M. 2005. Novel modes of protein-RNA recognition in the RNAi pathway. Curr Opin Struct Biol 15:107-115.
Liu HY, Chiang YC, Pan J, Chen J, Salvadore C, Audino DC, Badarinarayana V, Palaniswamy V, Anderson B, Denis CL. 2001. Characterization of CAF4 and CAF16 reveals a functional connection between the CCR4-NOT complex and a subset of SRB proteins of the RNA polymerase II holoenzyme. J Biol Chem 276:7541-7548.
Liu J, Carmell MA, Rivas FV, Marsden CG, Thomson JM, Song JJ, Hammond SM, Joshua-Tor L, Hannon GJ. 2004. Argonaute2 is the catalytic engine of mammalian RNAi. Science 305:1437-1441.
111
Lykke-Andersen J, Shu MD, Steitz JA. 2000. Human Upf proteins target an mRNA for nonsense-mediated decay when bound downstream of a termination codon. Cell 103:1121-1131.
Lykke-Andersen J, Wagner E. 2005. Recruitment and activation of mRNA decay enzymes by two ARE-mediated decay activation domains in the proteins TTP and BRF-1. Genes Dev 19:351-361.
Mader S, Lee H, Pause A, Sonenberg N. 1995. The translation initiation factor eIF-4E binds to a common motif shared by the translation factor eIF-4 gamma and the translational repressors 4E-binding proteins. Mol Cell Biol 15:4990-4997.
Mallory AC, Reinhart BJ, Jones-Rhoades MW, Tang G, Zamore PD, Barton MK, Bartel DP. 2004. MicroRNA control of PHABULOSA in leaf development: importance of pairing to the microRNA 5' region. EMBO J 23:3356-3364.
Maquat LE. 2004. Nonsense-mediated mRNA decay: splicing, translation and mRNP dynamics. Nat Rev Mol Cell Biol 5:89-99.
Mascarenhas C, Edwards-Ingram LC, Zeef L, Shenton D, Ashe MP, Grant CM. 2008. Gcn4 is required for the response to peroxide stress in the yeast Saccharomyces cerevisiae. Mol Biol Cell 19:2995-3007.
Matsuo R, Kubota H, Obata T, Kito K, Ota K, Kitazono T, Ibayashi S, Sasaki T, Iida M, Ito T. 2005. The yeast eIF4E-associated protein Eap1p attenuates GCN4 translation upon TOR-inactivation. FEBS Lett 579:2433-2438.
Mendell JT, ap Rhys CM, Dietz HC. 2002. Separable roles for rent1/hUpf1 in altered splicing and decay of nonsense transcripts. Science 298:419-422.
Mitchell P, Tollervey D. 2000. Musing on the structural organization of the exosome complex. Nat Struct Biol 7:843-846.
Mnaimneh S, Davierwala AP, Haynes J, Moffat J, Peng WT, Zhang W, Yang X, Pootoolal J, Chua G, Lopez A, Trochesset M, Morse D, Krogan NJ, Hiley SL, Li Z, Morris Q, Grigull J, Mitsakakis N, Roberts CJ, Greenblatt JF, Boone C, Kaiser CA, Andrews BJ, Hughes TR. 2004. Exploration of essential gene functions via titratable promoter alleles. Cell 118:31-44.
Moser MJ, Holley WR, Chatterjee A, Mian IS. 1997. The proofreading domain of Escherichia coli DNA polymerase I and other DNA and/or RNA exonuclease domains. Nucleic Acids Res 25:5110-5118.
Muhlrad D, Decker CJ, Parker R. 1994. Deadenylation of the unstable mRNA encoded by the yeast MFA2 gene leads to decapping followed by 5'-->3' digestion of the transcript. Genes Dev 8:855-866.
112
Muhlrad D, Decker CJ, Parker R. 1995. Turnover mechanisms of the stable yeast PGK1 mRNA. Mol Cell Biol 15:2145-2156.
Muhlrad D, Parker R. 1994. Premature translational termination triggers mRNA decapping. Nature 370:578-581.
Nelson MR, Leidal AM, Smibert CA. 2004. Drosophila Cup is an eIF4E-binding protein that functions in Smaug-mediated translational repression. EMBO J 23:150-159.
Nielsen DA, Shapiro DJ. 1990. Estradiol and estrogen receptor-dependent stabilization of a minivitellogenin mRNA lacking 5,100 nucleotides of coding sequence. Mol Cell Biol 10:371-376.
Oberstrass FC, Lee A, Stefl R, Janis M, Chanfreau G, Allain FH. 2006. Shape-specific recognition in the structure of the Vts1p SAM domain with RNA. Nat Struct Mol Biol 13:160-167.
Ohnishi T, Yamashita A, Kashima I, Schell T, Anders KR, Grimson A, Hachiya T, Hentze MW, Anderson P, Ohno S. 2003. Phosphorylation of hUPF1 induces formation of mRNA surveillance complexes containing hSMG-5 and hSMG-7. Mol Cell 12:1187-1200.
Okamura K, Ishizuka A, Siomi H, Siomi MC. 2004. Distinct roles for Argonaute proteins in small RNA-directed RNA cleavage pathways. Genes Dev 18:1655-1666.
Olivas W, Parker R. 2000. The Puf3 protein is a transcript-specific regulator of mRNA degradation in yeast. EMBO J 19:6602-6611.
Page MF, Carr B, Anders KR, Grimson A, Anderson P. 1999. SMG-2 is a phosphorylated protein required for mRNA surveillance in Caenorhabditis elegans and related to Upf1p of yeast. Mol Cell Biol 19:5943-5951.
Pal M, Ishigaki Y, Nagy E, Maquat LE. 2001. Evidence that phosphorylation of human Upfl protein varies with intracellular location and is mediated by a wortmannin-sensitive and rapamycin-sensitive PI 3-kinase-related kinase signaling pathway. RNA 7:5-15.
Pan X, Ye P, Yuan DS, Wang X, Bader JS, Boeke JD. 2006. A DNA integrity network in the yeast Saccharomyces cerevisiae. Cell 124:1069-1081.
Parker R, Sheth U. 2007. P bodies and the control of mRNA translation and degradation. Mol Cell 25:635-646.
Parker R, Song H. 2004. The enzymes and control of eukaryotic mRNA turnover. Nat Struct Mol Biol 11:121-127.
Pastori RL, Moskaitis JE, Buzek SW, Schoenberg DR. 1991. Coordinate estrogen-regulated instability of serum protein-coding messenger RNAs in Xenopus laevis. Mol Endocrinol 5:461-468.
113
Peng SS, Chen CY, Xu N, Shyu AB. 1998. RNA stabilization by the AU-rich element binding protein, HuR, an ELAV protein. EMBO J 17:3461-3470.
Pilkington GR, Parker R. 2008. Pat1 contains distinct functional domains that promote P-body assembly and activation of decapping. Mol Cell Biol 28:1298-1312.
Poole TL, Stevens A. 1997. Structural modifications of RNA influence the 5' exoribonucleolytic hydrolysis by XRN1 and HKE1 of Saccharomyces cerevisiae. Biochem Biophys Res Commun 235:799-805.
Ptushkina M, von der Haar T, Karim MM, Hughes JM, McCarthy JE. 1999. Repressor binding to a dorsal regulatory site traps human eIF4E in a high cap-affinity state. EMBO J 18:4068-4075.
Qiao F, Bowie JU. 2005. The many faces of SAM. Sci STKE 2005:re7.
Ren YG, Martinez J, Virtanen A. 2002. Identification of the active site of poly(A)-specific ribonuclease by site-directed mutagenesis and Fe(2+)-mediated cleavage. J Biol Chem 277:5982-5987.
Rendl LM, Bieman MA, Smibert CA. 2008. S. cerevisiae Vts1p induces deadenylation-dependent transcript degradation and interacts with the Ccr4p-Pop2p-Not deadenylase complex. RNA 14:1328-1336.
Scheller N, Resa-Infante P, de la Luna S, Galao RP, Albrecht M, Kaestner L, Lipp P, Lengauer T, Meyerhans A, Diez J. 2007. Identification of PatL1, a human homolog to yeast P body component Pat1. Biochimica et biophysica acta 1773:1786-1792.
Schwartz D, Decker CJ, Parker R. 2003. The enhancer of decapping proteins, Edc1p and Edc2p, bind RNA and stimulate the activity of the decapping enzyme. RNA 9:239-251.
Schwartz DC, Parker R. 1999. Mutations in translation initiation factors lead to increased rates of deadenylation and decapping of mRNAs in Saccharomyces cerevisiae. Mol Cell Biol 19:5247-5256.
Semotok JL, Cooperstock RL, Pinder BD, Vari HK, Lipshitz HD, Smibert CA. 2005. Smaug Recruits the CCR4/POP2/NOT Deadenylase Complex to Trigger Maternal Transcript Localization in the Early Drosophila Embryo. Curr Biol 15:284-294.
Semotok JL, Luo H, Cooperstock RL, Karaiskakis A, Vari HK, Smibert CA, Lipshitz HD. 2008. Drosophila maternal Hsp83 mRNA destabilization is directed by multiple SMAUG recognition elements in the open reading frame. Mol Cell Biol 28:6757-6772.
Serin G, Gersappe A, Black JD, Aronoff R, Maquat LE. 2001. Identification and characterization of human orthologues to Saccharomyces cerevisiae Upf2 protein and Upf3 protein (Caenorhabditis elegans SMG-4). Mol Cell Biol 21:209-223.
114
She M, Decker CJ, Svergun DI, Round A, Chen N, Muhlrad D, Parker R, Song H. 2008. Structural basis of dcp2 recognition and activation by dcp1. Mol Cell 29:337-349.
Sheth U, Parker R. 2003. Decapping and decay of messenger RNA occur in cytoplasmic processing bodies. Science 300:805-808.
Smibert CA, Lie YS, Shillinglaw W, Henzel WJ, Macdonald PM. 1999. Smaug, a novel and conserved protein, contributes to repression of nanos mRNA translation in vitro. RNA 5:1535-1547.
Smibert CA, Wilson JE, Kerr K, Macdonald PM. 1996. smaug protein represses translation of unlocalized nanos mRNA in the Drosophila embryo. Genes & Dev 10:2600-2609.
Steiger M, Carr-Schmid A, Schwartz DC, Kiledjian M, Parker R. 2003. Analysis of recombinant yeast decapping enzyme. RNA 9:231-238.
Stevens A. 1980. Purification and characterization of a Saccharomyces cerevisiae exoribonuclease which yields 5'-mononucleotides by a 5' leads to 3' mode of hydrolysis. J Biol Chem 255:3080-3085.
Stevens A, Maupin MK. 1987. A 5'----3' exoribonuclease of Saccharomyces cerevisiae: size and novel substrate specificity. Arch Biochem Biophys 252:339-347.
Tadauchi T, Matsumoto K, Herskowitz I, Irie K. 2001. Post-transcriptional regulation through the HO 3'-UTR by Mpt5, a yeast homolog of Pumilio and FBF. EMBO J 20:552-561.
Tadros W, Goldman AL, Babak T, Menzies F, Vardy L, Orr-Weaver T, Hughes TR, Westwood JT, Smibert CA, Lipshitz HD. 2007. SMAUG is a major regulator of maternal mRNA destabilization in Drosophila and its translation is activated by the PAN GU kinase. Dev Cell 12:143-155.
Tadros W, Lipshitz HD. 2005. Setting the stage for development: mRNA translation and stability during oocyte maturation and egg activation in Drosophila. Dev Dyn 232:593-608.
Teixeira D, Parker R. 2007. Analysis of P-body assembly in Saccharomyces cerevisiae. Mol Biol Cell 18:2274-2287.
Teixeira D, Sheth U, Valencia-Sanchez MA, Brengues M, Parker R. 2005. Processing bodies require RNA for assembly and contain nontranslating mRNAs. RNA 11:371-382.
Tharun S, He W, Mayes AE, Lennertz P, Beggs JD, Parker R. 2000. Yeast Sm-like proteins function in mRNA decapping and decay. Nature 404:515-518.
Tharun S, Parker R. 2001. Targeting an mRNA for decapping: displacement of translation factors and association of the Lsm1p-7p complex on deadenylated yeast mRNAs. Mol Cell 8:1075-1083.
115
Tritschler F, Eulalio A, Truffault V, Hartmann MD, Helms S, Schmidt S, Coles M, Izaurralde E, Weichenrieder O. 2007. A divergent Sm fold in EDC3 proteins mediates DCP1 binding and P-body targeting. Mol Cell Biol 27:8600-8611.
Tucker M, Staples RR, Valencia-Sanchez MA, Muhlrad D, Parker R. 2002. Ccr4p is the catalytic subunit of a Ccr4p/Pop2p/Notp mRNA deadenylase complex in Saccharomyces cerevisiae. EMBO J 21:1427-1436.
Tucker M, Valencia-Sanchez MA, Staples RR, Chen J, Denis CL, Parker R. 2001. The transcription factor associated Ccr4 and Caf1 proteins are components of the major cytoplasmic mRNA deadenylase in Saccharomyces cerevisiae. Cell 104:377-386.
Unterholzner L, Izaurralde E. 2004. SMG7 acts as a molecular link between mRNA surveillance and mRNA decay. Mol Cell 16:587-596.
Valencia-Sanchez MA, Liu J, Hannon GJ, Parker R. 2006. Control of translation and mRNA degradation by miRNAs and siRNAs. Genes Dev 20:515-524.
van Hoof A, Frischmeyer PA, Dietz HC, Parker R. 2002. Exosome-mediated recognition and degradation of mRNAs lacking a termination codon. Science 295:2262-2264.
van Hoof A, Parker R. 1999. The exosome: a proteasome for RNA? Cell 99:347-350.
Vilela C, Velasco C, Ptushkina M, McCarthy JE. 2000. The eukaryotic mRNA decapping protein Dcp1 interacts physically and functionally with the eIF4F translation initiation complex. EMBO J 19:4372-4382.
Wang C, Dickinson LK, Lehmann R. 1994. Genetics of nanos localization in Drosophila. Dev Dynam 199:103-115.
Wang C, Lehmann R. 1991. Nanos is the localized posterior determinant in Drosophila. Cell 66:637-647.
Weiss IM, Liebhaber SA. 1994. Erythroid cell-specific determinants of alpha-globin mRNA stability. Mol Cell Biol 14:8123-8132.
Wharton RP, Struhl G. 1991. RNA regulatory elements mediate control of Drosophila body pattern by the posterior morphogen nanos. Cell 67:955-967.
Wickens M, Bernstein DS, Kimble J, Parker R. 2002. A PUF family portrait: 3'UTR regulation as a way of life. Trends Genet 18:150-157.
Wilusz CJ, Gao M, Jones CL, Wilusz J, Peltz SW. 2001a. Poly(A)-binding proteins regulate both mRNA deadenylation and decapping in yeast cytoplasmic extracts. RNA 7:1416-1424.
Wilusz CJ, Wormington M, Peltz SW. 2001b. The Cap-to-Tail guide to mRNA turnover. Nat Rev Mol Cell Biol 2:237 - 246.
116
Yamashita A, Chang TC, Yamashita Y, Zhu W, Zhong Z, Chen CY, Shyu AB. 2005. Concerted action of poly(A) nucleases and decapping enzyme in mammalian mRNA turnover. Nat Struct Mol Biol 12:1054-1063.
Yamashita A, Ohnishi T, Kashima I, Taya Y, Ohno S. 2001. Human SMG-1, a novel phosphatidylinositol 3-kinase-related protein kinase, associates with components of the mRNA surveillance complex and is involved in the regulation of nonsense-mediated mRNA decay. Genes Dev 15:2215-2228.
Yekta S, Shih IH, Bartel DP. 2004. MicroRNA-directed cleavage of HOXB8 mRNA. Science 304:594-596.
Zaessinger S, Busseau I, Simonelig M. 2006. Oskar allows nanos mRNA translation in Drosophila embryos by preventing its deadenylation by Smaug/CCR4. Development 133:4573-4583.
Zhang S, Williams CJ, Wormington M, Stevens A, Peltz SW. 1999. Monitoring mRNA decapping activity. Methods 17:46-51.