Characterization of the N-terminal Region of the RNA-
Binding Protein Smaug in Post-transcriptional Regulation
During Drosophila Embryogenesis
by
Matthew Hong Kei Cheng
A thesis submitted in conformity with the requirements
for the degree of Master of Science
Graduate Department of Biochemistry
University of Toronto
© Copyright by Matthew Hong Kei Cheng 2013
ii
Characterization of the N-terminal Region of the RNA-Binding Protein
Smaug in Post-transcriptional Regulation During Drosophila Embryogenesis
Matthew Hong Kei Cheng
Master of Science
Graduate Department of Biochemistry
University of Toronto
2013
Abstract
The Drosophila sequence-specific RNA-binding protein Smaug (Smg) regulates the
expression of mRNAs in the early fly embryo. It is the founding member of a conserved family
of post-transcriptional regulators defined by an RNA-binding sterile alpha motif (SAM) domain.
Smg regulates gene expression through its ability to repress the translation, and/or induce
degradation of target mRNAs. Through a structure-function analysis using smg truncation
mutants, I show that sequences N- and C-terminal to the Smg SAM domain are involved, and
have partially redundant roles in mRNA decay. Moreover, another conserved region of Smg
modulates the mRNA decay function of the N-terminal sequences in a transcript-specific
manner. Finally, sequences within the Smg N- and C-terminal regions are also required for the
degradation of Smg protein.
iii
Acknowledgments
I am sincerely grateful for everyone who has helped me, and made this research possible.
I am extremely thankful to my supervisor, Dr. Craig Smibert, whose guidance and patience were
invaluable for my development as a researcher and scientist. I am truly privileged to have gotten
my first research experience with him.
I would like to thank my committee members, Dr. Henry Krause and Dr. David Williams, whose
insights strengthened my research and knowledge.
A warm thanks to all members of the Smibert and Lipshitz labs, for their discussions, advice and
training. I would like to acknowledge Najeeb Siddiqui, who generated the FLsmg and NTsmg
transgenic constructs used in this work.
I am eternally indebted to my parents, John Cheng and Wandy Lam, for their love,
encouragement and support throughout my life and studies. Thanks to my sister, Gloria, for her
love and encouragement. I could not have done this without them.
I am grateful for my girlfriend, Marion Weberruβ, for her love and support, as well as the many
wonderful experiences we shared.
Last but not least, I would like to thank my friends for the great times we had together, the
delightful conversations and their interest in my research.
iv
Table of Contents
Acknowledgments ........................................................................................................................ iii
List of Figures .............................................................................................................................. vii
List of Tables .............................................................................................................................. viii
List of Abbreviations ................................................................................................................... ix
1 Introduction .............................................................................................................................. 1
1.1 Post-transcriptional regulation .......................................................................................... 1
1.2 The action of RNA-binding proteins in post-transcriptional regulation ........................... 2
1.2.1 Subcellular mRNA localization .............................................................................. 2
1.2.2 Translational control ............................................................................................... 3
1.2.3 Control of mRNA stability ...................................................................................... 5
1.2.4 ASH1 mRNA, an example of the combinatory effects of RBPs ............................. 7
1.3 Post-transcriptional regulation in Drosophila embryogenesis ......................................... 8
1.3.1 Bicoid, the anterior determinant ............................................................................. 9
1.3.2 Oskar, a posterior determinant and component of pole plasm formation ............... 9
1.3.3 Nanos, the posterior determinant .......................................................................... 10
1.4 Smaug, an RNA-binding protein .................................................................................... 11
1.4.1 Mechanisms of Smg-mediated translation repression .......................................... 13
1.4.2 Smg protein in the embryo .................................................................................... 15
1.4.3 Targets of Smg-mediated regulation ..................................................................... 17
1.4.4 Spatial regulation of Smg function ....................................................................... 18
1.5 Thesis rationale ............................................................................................................... 18
2 Materials and Methods .......................................................................................................... 19
2.1 Fly stocks and crosses ..................................................................................................... 19
2.2 P-element excision .......................................................................................................... 19
v
2.3 Genomic DNA extraction and PCR ................................................................................ 19
2.4 Hatch rate analysis .......................................................................................................... 20
2.5 DAPI staining ................................................................................................................. 20
2.6 Cuticle preparations ........................................................................................................ 22
2.7 Transgene construction ................................................................................................... 22
2.8 Extract preparation and Western blotting ....................................................................... 23
2.9 RNA methods and RT-qPCR .......................................................................................... 24
2.10 SRE prediction ................................................................................................................ 26
3 Results ..................................................................................................................................... 26
3.1 Generation of smg protein null alleles ............................................................................ 26
3.2 Characterization of the smg30
and smg47
alleles ............................................................. 28
3.3 Generation of smg constructs and transgenic smg flies .................................................. 35
3.4 NTsmg and NTdSSR2 proteins are expressed at wild-type levels ................................... 37
3.5 NTsmg and NTdSSR2 proteins are expressed in later stage embryos ............................. 37
3.6 NTsmg and NTdSSR2 do not rescue hatching defects .................................................... 39
3.7 NTsmg and NTdSSR2 attenuates nuclear division defects .............................................. 39
3.8 NTsmg and NTdSSR2 proteins partially rescue cuticle formation .................................. 42
3.9 NTsmg and NTdSSR2 can mediate mRNA decay ........................................................... 45
3.10 Two copies of the NTsmg and NTdSSR2 transgenes enhance rescue of the smg
mutant phenotype ............................................................................................................ 53
3.11 Co-expression of NTsmg and CTsmg offer modest improvement over a single copy
of NTsmg alone ............................................................................................................... 57
4 Discussion ................................................................................................................................ 60
4.1 Summary and Conclusions ............................................................................................. 60
4.1.1 Generation of smg protein null flies ...................................................................... 60
4.1.2 Both the Smg N- and C-termini are important for Smg function ......................... 60
vi
4.1.3 Smg employs multiple mechanisms to induce transcript decay ........................... 61
4.1.4 The SSR2 domain plays a transcript-specific role in Smg function ..................... 62
4.1.5 The Smg C-terminus functions in Smg protein degradation ................................ 62
4.2 Future Directions ............................................................................................................ 63
4.2.1 The role of NTsmg in mRNA decay ..................................................................... 63
4.2.2 The mechanism of NTsmg function in mRNA decay ........................................... 64
4.2.3 The mechanism of CTsmg function in mRNA decay ........................................... 68
4.2.4 The role of the SSR2 in Smg function .................................................................. 69
4.2.5 Mechanisms of Smg-mediated translation repression .......................................... 69
4.2.6 Mechanism of Smg protein degradation ............................................................... 71
4.2.7 Assessing additional smg mutant proteins ............................................................ 72
References .................................................................................................................................... 73
vii
List of Figures
Figure 1 Smg is the founding member of a family of conserved post-transcriptional regulators.12
Figure 2 Known mechanisms of Smg-mediated regulation. ........................................................ 16
Figure 3 Generation of smg mutant alleles by imprecise P-element excision mutagenesis ........ 21
Figure 4 Significant portions of the smg gene region is deleted in the smg30
and smg47
mutant
alleles. ........................................................................................................................................... 29
Figure 5 Progression of syncytial nuclear divisions is defective in smg30
and smg47
embryos. .. 33
Figure 6 A schematic of the proteins expressed by the transgenic constructs employed in this
study. ............................................................................................................................................. 36
Figure 7 The FLsmg, NTsmg, and NTdSSR2 proteins are expressed at similar levels as wild-type
Smg ............................................................................................................................................... 38
Figure 8 Stabilization of the NTsmg and NTdSSR2 proteins. ...................................................... 40
Figure 9 NTsmg and NTdSSR2 proteins attenuate nuclear division defects found in smg mutant
embryos. ........................................................................................................................................ 43
Figure 10 NTsmg or NTdSSR2 proteins partially rescue cuticle formation ................................ 46
Figure 11 NTsmg and NTdSSR2 proteins can mediate Hsp83 mRNA decay .............................. 49
Figure 12 NTsmg and NTdSSR2 proteins can mediate arrest mRNA decay ............................... 50
Figure 13 NTsmg and NTdSSR2 proteins can mediate BicC mRNA decay ................................. 52
Figure 14 Enhanced rescue of the cuticle phenotype of smg47
mutant embryos by increased copy
number of the NTsmg transgene, or co-expression of NTsmg and CTsmg ................................... 55
Figure 15 Improved rescue of the nuclear division defects of smg47
mutant embryos by increased
copy number of the NTsmg transgene, or co-expression of NTsmg and CTsmg. ......................... 58
Figure 16 Six additional motifs in the Smg protein are conserved among Drosophilids and some
insects. ........................................................................................................................................... 66
viii
List of Tables
Table 1 Hatch rate analysis of smg30
and smg47
mutant embryos ................................................ 31
Table 2 Hatch rate analysis of smg47
mutant embryos rescued with a single copy of the FLsmg,
NTsmg, or NTdSSR2 transgenes .................................................................................................... 41
Table 3 Hatch rate analysis of smg47
mutant embryos rescued with two copies of the NTsmg
transgene, or co-expression of the NTsmg and CTsmg transgenes ............................................... 54
ix
List of Abbreviations
4E-BP – eIF4E-binding protein
AEL – after egglaying
Ago – Argonaute
ARE – A/U-rich element
ASH1 – asymmetric synthesis of HO 1
Aub – Aubergine
bcd - bicoid
BicC – Bicaudal C
BicD – Bicaudal D
Bru – Bruno
CPE – cytoplasmic polyadenylation element
CPEB – cytoplasmic polyadenylation element-binding protein
CTsmg – C-terminus Smg
Dcp – decapping enzyme
DNA – deoxyribonucleaic acid
DP1 – Dodeca-satellite-binding protein 1
Egl - Egalitarian
eIF – eukaryotic initiation factor
EJC – exon junction complex
EMS – ethyl methanesulfonate
ESCRT- II - endosomal sorting complexes required for transport II
Exu - Exuperentia
FLsmg – full length Smg
FMRP – fragile X mental retardation protein
GLD-2 – defective in germ line development 2
Glo – Glorund
GM-CSF – granulocyte macrophage colony-stimulating factor
gt – giant
hb – hunchback
hnRNP – heterogeneous ribonucleoprotein particle
x
HRP – horseradish peroxidase
Hrp48 – Heterogeneous nuclear ribonucleoprotein at 27C
Hsp83 – Heat shock protein 83
IRE – iron-responsive element
IRP – iron regulatory protein
Khd1p – KH domain protein 1
kni – knirps
miRNA – microRNA
mRNA – messenger RNA
mRNP – messenger ribonucleoprotein particle
mTOR – mammalian target of rapamycin
MZT – maternal-to-zygotic transition
nos - nanos
NTdSSR2 – N-terminus Smg delta SSR2
NTsmg – N-terminus Smg
ORF – open reading frame
osk - oskar
PABP – poly(A)-binding protein
PAP – poly(A) polymerase
PARN – poly(A) ribonuclease
P-bodies – processing bodies
PBS – phosphate buffered saline
PCR – polymerase chain reaction
PIC – pre-initiation complex
piRNA – Piwi-interacting RNA
PTW – 0.1% Tween in 1X PBS
Puf6p – Pumilio/FBP protein family 6 protein
RBD – RNA-binding domain
RBP – RNA-binding protein
RISC – RNA-induced silencing complex
RNA – ribonucleic acid
xi
RT-qPCR – reverse transcription quantitative polymerase chain reaction
Rump – Rumpelstiltskin
SAM – sterile alpha motif
SDS – sodium dodecyl sulfate
Smg – Smaug
Sqd – Squid
SRE – Smg recognition element
SSR – Smg similarity region
Stau - Staufen
Swa - Swallow
TfR – transferrin receptor
TNF-α – tumor necrosis factor alpha
TTP - Tristetraproline
UTR – untranslated region
Vts1p – VTII-2 suppressor
XRN1 – exoribonuclease 1
ZBP – zipcode binding protein
1
1 INTRODUCTION
1.1 Post-transcriptional regulation
Multiple levels of regulation exist to ensure that specific subsets of an organism’s genes
are expressed in distinct cell types, at different times, and/or under certain circumstances
(Alberts et al., 2002). Aside from transcriptional control, mechanisms acting post-
transcriptionally are also major contributors in regulating gene expression (Sonenberg and
Hinnebusch, 2009). These mechanisms – falling under the term post-transcriptional regulation –
include splicing, transport, localization, translation activation/repression, and mRNA
stabilization/destabilization (Alberts et al., 2002). Together they greatly influence protein
expression, and over 90% of mRNAs are subject to post-transcriptional regulation
(Schwanhäusser et al., 2011; Pichon et al., 2012).
The ability to manipulate the translation and stability of mRNAs in the cytoplasm after
they have been transcribed allows rapid changes in protein levels in response to stimuli and/or
developmental cues (Lipshitz and Smibert, 2000; Sonenberg and Hinnebusch, 2009). Specific
regulatory events can act on subsets of mRNAs to stabilize or destabilize them, and/or actively
repress their translation, thus controlling the amounts of the corresponding proteins (Sonenberg
and Hinnebusch, 2009). Moreover, subcellular mRNA localization, as well as spatial regulation
of translation or mRNA stability can be important mechanisms to control protein localization
(Lipshitz and Smibert, 2000; Macdonald, 2011). Finally, with the exception of mRNA
degradation, these mechanisms of regulation are reversible (Lipshitz and Smibert, 2000; Nelson
et al., 2004; Baez et al., 2011).
Post-transcriptional controls that function in the cytoplasm are important in a variety of
cell types. For example in dendritic cells, translational activation of specific mRNAs at
individual synapses upon neuronal stimulation is important for synaptic plasticity, learning and
memory (Sonenberg and Hinnebusch, 2009; Baez et al., 2011). Cytoplasmic post-transcriptional
controls are particularly important in cells where transcriptional regulation is not an option
(Lipshitz and Smibert, 2000; Lasko 2009). For example, platelets are anucleated, and post-
transcriptional mechanisms modulate cellular pathways in response to inflammatory signals
(Weyrich et al., 2004). Similarly, during early embryogenesis in animals, nuclei are
transcriptionally silent, and thus this stage of development is driven by maternal mRNAs that are
2
deposited into the cytoplasm of the oocyte (Tadros and Lipshitz, 2005). During this phase, post-
transcriptional regulatory events are in place to ensure that these maternal mRNAs are expressed
in the correct spatial and temporal context (Lasko, 2009).
The importance of post-transcriptional regulation is highlighted by the numerous disease
states associated with dysregulation of this form of regulation (Sonenberg and Hinnesbusch,
2009; Lasko, 2009). For example, overexpression of eIF4E in human and mouse cells in tissue
culture have been linked to tumorigenesis by causing misregulated translation of mRNAs
involved in regulating cell proliferation and apoptosis (Larsson et al., 2007; Mamane et al.,
2007). Moreover, mice deficient in the downstream mTOR effectors, 4E-BP1 and 4E-BP2,
showed sensitivity to obesity and insulin resistance resulting from increased translation of
mRNAs associated with adipogenesis (Le Bacquer et al., 2007). Finally, reduced expression of
the translation repressor fragile X mental retardation protein (FMRP) is associated with the
neuropsychiatric disease fragile X syndrome (Sonenberg and Hinnesbusch, 2009). This results
from increased translation of mRNAs whose products are involved in synaptic plasticity and
brain development (Napoli et al., 2008). The mechanisms of post-transcriptional control are
largely mediated by RNA-binding proteins (RBPs) which can influence the localization,
translation, and/or stability of bound mRNAs (Pichon et al., 2012).
1.2 The action of RNA-binding proteins in post-transcriptional regulation
Post-transcriptional regulation can be achieved through the binding of cis-elements in
mRNAs by trans-acting factors, including RBPs, whose actions influence the fate of a bound
mRNA (Lipshitz and Smibert, 2000; Tadros and Lipshitz, 2005). In the cytoplasm, there are
three ways in which RBPs can affect an mRNA’s expression. They are the control of the
transcript’s subcellular localization, translational status, and/or its stability (Lipshitz and Smibert
2000; Tadros and Lipshitz, 2005; Macdonald, 2011).
1.2.1 Subcellular mRNA localization
Directed transport of mRNAs is one mechanism of mRNA localization which involves
motor-driven movement of transcripts along cytoskeletal elements (Lipshitz and Smibert, 2000).
Over long distances – in large cells such as dendrites – transport utilizes the microtubule network
(Pokrywka and Stephenson, 1991; Mach and Lehmann, 1997). In contrast, transport over short
3
distances can utilize microfilament networks (Erdélyi et al., 1995; Beach, et al., 1999; Lipshitz
and Smibert, 2000; Blower, 2013). mRNAs directly transported along cytoskeletal networks are
packaged into messenger ribonucleoprotein particles (mRNPs) (Mcdonald, 2011). Formation of
these mRNPs involves recognition and binding of cis-element(s) within transcripts – referred to
as zipcodes – by zipcode-binding proteins (ZBPs). These RBPs typically facilitate mRNA
localization through their interaction with adaptor proteins which in turn interact with molecular
motors (Blower, 2013). For example, during Drosophila oogenesis, a number of maternal
mRNAs are bound by the RBP Egalitarian (Egl) which interacts with Bicaudal D (BicD), which
in turn binds the motor protein dynein. The formation of the mRNA/Egl/BicD/dynein mRNP
serves to transport the mRNA from the nurse cells to the oocyte (Mach and Lehmann, 1997;
Clark et al., 2007; Dienstbier et al., 2009). Similarly in S. cerevisiae, ASH1 mRNAs are localized
to daughter cells by the RBP She2p, the myosin cargo adaptor She3p, and the myosin Myo4p
(Long et al., 1997; Münchow et al., 1999).
1.2.2 Translational control
In addition to directed transport, RBPs can also alter the translational status of a bound
mRNA (Besse and Ephrussi, 2008). Translation can be divided into the steps of initiation,
elongation, and termination (Besse and Ephrussi, 2008; Pichon et al., 2012). A key part of
initiation is the step-wise assembly of the eIF4F complex, which consists of the cap-binding
factor eIF4E, the RNA helicase eIF4A, and the scaffold protein eIF4G at the mRNA’s 5’ m7G
cap (Sonenberg and Hinnebusch, 2009; Aitken and Lorsch, 2012). eIF4E recognizes and binds
the cap of an mRNA, and recruits eIF4G through a direct interaction. eIF4A is then anchored
through eIF4G to form the eIF4F complex. This complex is responsible for recruitment of the
40S ribosomal subunit to the 5’ end of the mRNA through the interaction of eIF4G with eIF3,
which in turn binds to the 40S ribosomal subunit. Once recruited, the 40S subunit scans the
mRNA in the 5’ to 3’ direction, a process that is facilitated by disruption of mRNA secondary
structure by the helicase activity of eIF4A. Scanning proceeds until the 40S subunit recognizes
the translation start codon at which point the 60S ribosomal subunit joins the 40S subunit,
creating the full 80S ribosome that then translates the transcript’s open reading frame (ORF)
(Aitken and Lorsch, 2012).
4
Translation initiation also involves an mRNA’s poly(A) tail (Sonenberg and Hinnebusch,
2009). The poly(A) tail is bound by the poly(A)-binding proteins (PABP), which have also been
shown to interact with eIF4G (Aitken and Lorsch, 2012). Thus, a poly(A) tail can facilitate the
recruitment of eIF4G to the mRNA via PABP binding (Besse and Ephrussi, 2008). Moreover,
the poly(A) tail/PABP/eIF4G/eIF4E/5’ cap interaction leads to a circularization of the mRNA
into a “closed loop”, which is thought to enhance translation by efficiently promoting pre-
initiation complex (PIC) joining and facilitating reinitation of ribosomes after termination
(Sonenberg and Hinnebusch, 2009). Therefore, the presence of both a cap and a poly(A) tail
work synergistically to increase initiation efficiency (Besse and Ephrussi, 2008; Sonenberg and
Hinnebusch, 2009).
As the initiation step is important for translation, it is not surprising that RBPs target this
step to regulate translation (Besse and Ephrussi, 2008; Sonenberg and Hinnebusch, 2009).
Indeed, several translational repressors are eIF4E-binding proteins (4E-BPs), whose interaction
with eIF4E blocks eIF4G binding, thereby blocking 40S subunit recruitment (Besse and
Ephrussi, 2008). In addition, RBPs can also function to control translation by recruiting poly(A)
polymerases (PAPs) or deadenylases to modulate poly(A) tail length, thereby altering efficiency
of initiation (Parker and Song, 2004; Besse and Ephrussi, 2008; Eckmann et al., 2011).
During Xenopus oocyte maturation, c-Mos and cyclin B1 mRNAs are among a number of
targets of translational regulation (Tadros and Lipshitz, 2005). Their regulation is mediated by
cis-elements in their 3’UTRs termed cytoplasmic polyadenylation elements (CPE). These
elements represent binding sites for the cytoplasmic polyadenylation element-binding protein
(CPEB), which facilitates the recruitment of a number of translational regulators to an mRNA
(Besse and Ephrussi, 2008). In immature oocytes, CPEB represses translation by recruiting both
Maskin and poly(A) ribonuclease (PARN) to a bound mRNA. Maskin is a 4E-BP, and as such
blocks the eIF4E/eIF4G interaction (Stebbins-Boaz et al., 1999). In parallel, PARN facilitates
translational repression by shortening the transcript’s poly(A) tail (Kim and Richter, 2006).
Interestingly, in immature oocytes the CPEB complex also contains the poly(A) polymerase
GLD-2. However, the greater activity of the PARN enzymes ensures that CPEB target mRNAs
have short poly(A) tails (Kim and Richter, 2006; Radford et al., 2008).
5
Upon progesterone stimulation, the oocyte undergoes maturation and CPEB becomes
phosphorylated by a kinase thought to be Aurora A, altering the regulatory effects of CPEB on
bound mRNAs (Mendez et al., 2000a,b). The phosphorylation of CPEB ejects PARN, allowing
GLD-2 to lengthen the transcript’s poly(A) tail, which in turn leads to PABP recruitment. PABP
is then able to recruit eIF4G, stimulating translation in part through the ability of newly recruited
eIF4G to disrupt the eIF4E/Maskin interaction (Mendez et al., 2000a,b; Radford et al., 2008).
1.2.3 Control of mRNA stability
The cap and poly(A) tail are also important for the stability of an mRNA as they prevent
5’ to 3’ and 3’ to 5’ exonucleolytic decay (Parker and Song, 2004; Eckmann et al., 2011; Jones et
al., 2012). The circularization of mRNAs is thought also to preclude the engagement of the
degradation machineries (Jones et al., 2012). Therefore, mRNA destabilization often begins with
deadenylation (Parker and Song, 2004). There are two deadenylases in eukaryotes which are
thought to deadenylate transcripts in the cytoplasm: the CCR4/POP2/NOT complex and PARN
(Copeland and Wormington, 2001; Chen et al., 2002; Tucker et al., 2002).
Following shortening of an mRNA’s poly(A) tail beyond a threshold level, the body of
the mRNA can be degraded by one of two pathways (Parker and Song, 2004). One of these
involves 3’ to 5’ exonucleolytic decay, mediated by a large protein complex called the exosome.
The exosome is composed of nine 3’ to 5’ nuclease subunits arranged in a ring-like manner (van
Hoof and Parker, 1999; Mitchell and Tollervey, 2000; Symmons et al., 2000). To mediate
mRNA turnover in the cytoplasm, the exosome interacts with a heterotrimeric complex
composed of the Ski2p, Ski3p and Ski8p subunits (Brown et al., 2000). The Ski2p subunit is a
DEAD box RNA helicase, thought to unwind RNA secondary structures using ATP hydrolysis
(Anderson and Parker, 1998), while the Ski3p-Ski8p complex recruits the exosome via a
bridging interaction mediated by the cytoplasm specific Ski7p subunit (Araki et al., 2001).
In macrophages, the Tristetraproline (TTP) protein mediates rapid decay of many
inflammatory and cancer associated mRNAs (Sanduja et al., 2011). Specifically, TTP can
recognize and bind A/U-rich elements (AREs) present in target mRNAs – including the tumor
necrosis factor-alpha (TNF-α) mRNA (Lykke-Andersen and Wagner, 2005; Cao et al., 2007).
During inflammatory responses, synthesis of TTP and its subsequent binding to TNF-α mRNAs
6
leads to deadenylation by the CCR4/POP2/NOT complex (Carballo et al., 1998; Lykke-
Andersen and Wagner, 2005). Following poly(A) tail removal, the exosome is recruited to the 3’
end of deadenylated TNF-α mRNAs via direct interaction with TTP, which results in the 3’ to 5’
decay of the mRNA body (Chen et al., 2002).
Transcript deadenylation can also trigger the removal of the 5’ m7G cap (decapping) by a
decapping enzyme, which leaves the mRNA susceptible to decay by 5’ to 3’ exonucleases
(Parker and Song, 2004). There are two types of decapping enzymes in eukaryotes, the scavenger
decapping enzyme (DcpS) and the Dcp1-Dcp2 complex (Beelman et al., 1996; Dunckley and
Parker, 1999; Liu et al., 2002). The scavenger DcpS is only able to decap short RNA substrates,
and is thought to release m7GDP from mRNAs which have been degraded in the 3’ to 5’
direction (Liu et al., 2002; Parker and Song, 2004). In contrast, the Dcp1-Dcp2 protein is the
major decapping enzyme in eukaryotes (Beelman et al., 1996). The catalytic activity lies in the
Dcp2 subunit, whose activity is thought to be stimulated by Dcp1 (Dunckley and Parker, 1999;
Parker and Song, 2004; She et al., 2004). Decapping of mRNAs can be stimulated by the
recruitment of the Dcp1-Dcp2 complex via direct interaction with RBPs (Parker and Song,
2004).
After decapping, the 5’ monophosphate becomes available for decay by the 5’ to 3’
exonuclease XRN1 (Stevens and Maupin, 1987; Parker and Song, 2004; Jones et al., 2012). For
example in mammalian cells, the GM-CSF and c-fos mRNAs contain AREs (Li and Kiledjian,
2010). These mRNAs are bound by TTP, which can recruit the Dcp1-Dcp2 complex through
direct interaction. Moreover, this interaction results in the sequestering of GM-CSF and c-fos
mRNAs to mRNA decay foci called P-bodies (Carballo et al., 2001; Stoecklin et al., 2006;
Franks and Lykke-Anderson, 2007). Here, the mRNAs are decapped by Dcp1-Dcp2, leaving the
vulnerable 5’ end for XRN1-mediated decay (Parker and Song, 2004; Stoecklin et al., 2006; Li
and Kiledjian, 2010). Interestingly, TTP can mediate both decapping/5’ to 3’ decay, as well as 3’
to 5’ decay of target mRNAs (Sanduja et al., 2011). The mechanism for TTP-mediated mRNA
decay appears to depend on whether the mRNA is being degraded in or outside of P-bodies
(Sanduja et al., 2011; Jones et al., 2012).
In addition to the two decay pathways outlined above, mRNA degradation can also occur
through endonucleolytic cleavage (Schoenberg, 2011; Jones et al., 2012). Endonuclease activity
7
cleaves the mRNA body, and does not require decapping or poly(A) tail shortening (Schoenberg,
2011). The endonucleolytic cleavage leaves a 5’ and 3’ RNA fragments which can then be
targeted for exonucleolytic decay by the exosome and XRN1, respectively (Parker and Song,
2004). The recruitment of endonucleases to mRNAs is mediated through their binding to cis-
elements (Schoenberg, 2011). However, in the absence of the proper environmental conditions or
stimuli, RBPs which bind to the same cis-elements can block endonuclease recruitment.
In mammalian cells, the stability of the transferrin receptor (TfR) mRNA is regulated in
response to intracellular levels of iron (Schoenberg, 2011). The TfR mRNA contains five copies
of a stem-loop cis-element called iron-responsive elements (IREs) in its 3’UTR (Casey et al.,
1988). Under low iron conditions these IREs are bound by iron regulatory proteins IRP1 and
IRP2, which stabilizes the mRNA (half-life ~3hrs) (Hirling et al., 1994; Philpott et al., 1994;
DeRusso et al., 1995). This results in an increase in the levels of TfR on the cell surface, allowing
the cell to internalize extracellular iron complexed to Transferrin. The stabilization of TfR
mRNA is thought to be due to the occupation of IREs by IRP1 and IRP2, thereby blocking
endonuclease cleavage. Upon elevation of intracellular iron levels, IRP1 and IRP2 are targeted
for degradation, leaving IREs accessible to an as yet unidentified endonuclease (Binder et al.,
1994; Schoenberg, 2011; Anderson et al., 2012). This iron-dependent regulatory mechanism
results in the rapid decay of TfR mRNAs (half-life ~45min) (Binder et al., 1994). Thus, RBPs
can have both stabilizing and destabilizing effects on bound mRNAs.
1.2.4 ASH1 mRNA, an example of the combinatory effects of RBPs
Several examples have been identified whereby the mechanisms of post-transcriptional
regulation outlined above can function together to regulate the expression of the same mRNA
(Hogan et al., 2008). For example in S. cerevisiae, the expression of ASH1 mRNA, which
encodes a transcriptional repressor that inhibits mating-type switching in daughter cells, is
regulated by a combination of mechanisms controlling ASH1 mRNA translation and localization
(Beach and Bloom, 2001). As described above, the directed transport of ASH1 mRNA is
mediated by the action of the microfilament-motor complex comprised of the RBP She2p, the
adaptor She3p, and the myosin Myo4p. During its transit ASH1 mRNA is translationally
repressed to prevent ectopic expression in the mother cell (Beach and Bloom, 2001; Besse and
Ephrussi, 2008). One component of the ASH1 mRNP which represses translation is the 4E-BP
8
Khd1p. Its interaction with eIF4E blocks eIF4G binding, thereby inhibiting recruitment of the
40S ribosomal subunit and translation initiation (Paquin et al., 2007). Another component
mediating ASH1 translation repression is the RBP Puf6p, which also prevents translation
initiation. Interestingly, Puf6p functions by blocking recruitment of the 60S ribosomal subunit
through interaction with the general translation factor eIF5B (Gu et al., 2004).
1.3 Post-transcriptional regulation in Drosophila embryogenesis
Development during the first 2.5 hours of Drosophila embryogenesis occurs in a
syncytium – a multinucleated cell – where a fertilized pronucleus undergoes 13 rounds of rapid
divisions without cytokinesis (Tadros and Lipshitz, 2009). During this time, transcription is
quiescent and development is driven by maternally contributed mRNAs and proteins deposited
into the egg during oogenesis. Over time, these maternal factors are replaced by zygotically
transcribed factors in a process termed the maternal-to-zygotic transition (MZT). Post-
transcriptional regulation is essential in regulating proper temporal and spatial expression of
maternal mRNAs, including their timely degradation during the embryo’s transition to
zygotically controlled development (Tadros and Lipshitz, 2005, 2009).
Approximately 55% of the Drosophila genome – or roughly 7,000 genes – are loaded
into the oocyte and are present in the early embryo (Tadros et al., 2007). Several of these
mRNAs encode spatial determinants whose expression in a particular region of the embryo
directs the development of particular structures. Post-transcriptional regulatory events help
ensure that spatial determinants are expressed at the right time and in the right place, and these
controls are often essential as ectopic expression of spatial determinants can result in lethal body
patterning defects (Gavis and Lehmann, 1992; Lipshitz and Smibert, 2000; Tadros and Lipshitz,
2005; Lasko, 2009). In the embryo, the antero-posterior axis is established by mRNAs that are
localized at opposite poles of the syncytium. Major determinants include the anteriorly localized
bicoid (bcd) mRNA, and the posteriorly localized oskar (osk) and nanos (nos) mRNAs (Berleth
et al., 1988; Strul et al., 1989; Ephrussi et al., 1991; Gavis and Lehmann, 1992). In the following
sections, I will discuss the post-transcriptional regulation of these three critical spatial
determinants.
9
1.3.1 Bicoid, the anterior determinant
Bicoid (Bcd) protein is a transcriptional activator and its expression in a gradient
emanating from the anterior of the embryo drives development of anterior body structures. This
protein gradient is established by the regulated translation of bcd mRNAs localized to the
anterior pole (Strul et al., 1989). The proper localization of bcd mRNA is dependent on cis-
elements in its 3’UTR and a number of trans-acting factors mediating microtubule-associated
transport, translation repression during transport, and translation activation at the anterior.
Transport of bcd is mediated by RBPs including the BicD/Egl complex, Exuperantia (Exu),
Swallow (Swa), and the double-stranded RNA-binding protein Staufen (Stau) (Berleth et al.,
1988; Cha et al., 2001; Arn et al., 2003; Clark et al., 2007; Weil et al., 2010). bcd mRNA
reaching the anterior pole is anchored in place through separate processes involving the ESCRT-
II complex, Stau, and Swa (Ferrandon et al., 1994; Irion and St. Johnston, 2007; Weil et al.,
2010). To prevent ectopic expression of bcd while it is in transit and during oogenesis, the
mRNA is translationally repressed by poorly understood mechanism(s) (Kugler and Lasko,
2009). In the embryo, translation is thought to be stimulated in part by poly(A) tail lengthening
via the poly(A) polymerase Wisp (Juge et al., 2002; Benoit et al., 2008). These regulatory
mechanisms result in an anterior gradient of Bcd protein, which regulates the transcription of
many target genes (Strul et al., 1989).
1.3.2 Oskar, a posterior determinant and component of pole plasm formation
Formation of posterior body pattern is dependent on localized expression of osk and nos
mRNAs at the posterior pole of the embryo (Ephrussi et al., 1991; Wang and Lehmann, 1991).
Oskar (Osk) protein exists in two isoforms – long and short – with both being essential
components for specification of posterior structures (Markussen et al., 1995). In addition, both
isoforms of the Osk protein are required for proper localization of osk mRNA (Ephrussi et al.,
1991; Kugler and Lasko, 2009).
Localized expression of osk mRNA also involves cis-elements in the mRNA’s 3’UTR
and the functions of a variety of trans-acting factors (Lasko, 2009). Similar to bcd, osk mRNA is
transported along microtubules through the functions of BicD/Egl, Exu, Stau, and hnRNP
proteins such as Hrp48, Squid (Sqd), and Glorund (Glo) (Martin et al., 2003; Huynh et al., 2004;
Yano et al., 2004; Norvell et al., 2005; Steinhauer and Kalderon, 2005; Kalifa et al., 2008).
10
Interestingly, components of the exon junction complex (EJC) have also been found to be
required for osk localization, suggesting a role for splicing in this process (Hachet and Ephrussi,
2004). Enrichment of osk mRNA at the posterior pole is thought to involve Par-1 and the long
isoform of Osk, acting in a positive feedback loop (Riechmann et al., 2002; Doerflinger et al.,
2006; Zimyanin et al., 2007). During transport, osk is translationally repressed by separate
mechanisms involving two RBPs, Bruno (Bru) and Bicaudal-C (BicC). Bru binds to sequences in
the osk mRNA’s 3’UTR and recruits the 4E-BP Cup (see below for description of Cup). BicC is
also required to repress osk translation. It has been speculated that it could recruit the
CCR4/POP2/NOT deadenylase to osk mRNA, although direct interaction between the BicC
protein and osk mRNA has not been documented (Lasko, 2009; Kugler and Lasko, 2009).
Derepression and activation of osk translation at the posterior occurs through poly(A) tail
lengthening by Orb, and the poly(A) polymerases, PAP and Wisp (Chang et al., 1999; Juge et al.,
2002; Castagnetti and Ephrussi, 2003; Benoit et al., 2008).
1.3.3 Nanos, the posterior determinant
Nanos (Nos) is a translation repressor, and directs the development of abdominal
structures in the developing embryo (Wang and Lehmann, 1991). Its expression is restricted to a
gradient emanating from the posterior, established through regulatory mechanisms involving cis-
elements in the nos 3’UTR (Gavis and Lehmann, 1992; Smibert et al., 1996). However, unlike
bcd and osk, nos mRNA is localized by a combination of cytoplasmic diffusion and anchoring
(Lipshitz and Smibert, 2000; Forrest and Gavis, 2003). This is an inefficient method such that
only about 4% of mRNAs are localized at the posterior pole, and unlocalized nos in the bulk
cytoplasm is translationally repressed (Bergsten and Gavis, 1999; Lipshitz and Smibert, 2000;
Kugler and Lasko, 2009). In late oocytes, this repression is achieved through a mechanism at the
level of initiation dependent on both the cap and poly(A) tail mediated by Glo, and an additional
separate mechanism acting post-initiation (Bergsten and Gavis, 1999; Andrews et al., 2011). nos
regulation in early embryogenesis is primarily mediated by Smaug (Smg), through translation
repression, and to an extent mRNA degradation (see below for description of Smg) (Smibert et
al., 1999; Zaessinger et al., 2006). nos mRNA at the posterior pole is anchored there through the
function of Rumpelstiltskin (Rump) which binds directly to the nos mRNA’s 3’UTR (Jain and
Gavis, 2008). nos mRNA localized to the posterior escapes the translational repression that
affects the unlocalized nos mRNA, resulting in accumulation of Nos protein at the posterior of
11
the embryo. While the molecular mechanisms that permit translation of nos mRNA at the
posterior are unclear, one model proposes that binding of the mRNA localization machinery to
nos mRNA prevents binding of translation repressors, thereby specifically activating nos
translation at the posterior (Bergsten and Gavis, 1999).
Translation of localized nos mRNA leads to a posterior gradient of Nos, which
antagonizes bcd and hb mRNAs (Wang and Lehmann, 1991). Nos-mediated repression of bcd
and hb allows the expression of the gap genes knirps (kni) and giant (gt), and downstream
development of posterior structures (Gavis and Lehmann, 1992).
1.4 Smaug, an RNA-binding protein
A major post-transcriptional regulator in Drosophila embryogenesis is the RBP Smaug
(Smg) (Smibert et al., 1996, 1999; Tadros et al., 2007; Benoit et al., 2009), which is the focus of
my thesis. The Smg protein was first identified as a translation repressor of unlocalized nos
mRNA in early Drosophila embryos (Smibert et al., 1996). Subsequently, Smg was found to
destabilize a large fraction of maternal mRNAs in the early embryo (Semotok et al., 2005;
Tadros et al., 2007). In the embryo, Smg is essential to the normal development during the
syncytial blastoderm stage and in directing the MZT (Dahanukar et al., 1999; Benoit et al.,
2009). Defects in embryos laid by homozygous smg mutant mothers (from here referred to as
smg mutant embryos) are first observed starting at division cycle 11 of the syncytial nuclear
divisions, when cortical nuclei fall out of a surface array and form aggregates (Dahanukar et al.,
1999). These defects reflect a failure to activate DNA replication checkpoints, which slows
nuclear division cycles (Benoit et al., 2009). Independent of its role in DNA replication
checkpoint activation, Smg is also necessary at a later stage to facilitate the MZT by triggering
transcription of the zygotic genome and the degradation of maternal mRNAs (Tadros et al.,
2007; Benoit et al., 2009; Siddiqui et al., 2012). Given Smg’s major role in embryogenesis, smg
mutant embryos fail to hatch (Dahanukar et al., 1999).
The Smg protein is a 999 amino acid protein and contain three domains conserved in the
human and mouse homologs (Figure 1) (Smibert et al., 1999). Two of these domains are called
12
Figure 1 – Smg is the founding member of a family of conserved post-transcriptional
regulators. The Drosophila Smg is the founding member of the family of post-transcriptional
regulators, conserved from yeast to humans, displayed in the cladogram. This family of proteins
is defined by the RNA-binding SAM domain. Moreover, several homologs contain additional
conserved domains termed Smg similarity regions – SSR1 in the case of yeast, and SSR1 and
SSR2 in the case of mice and humans. The species of the indicated Smg family member is
indicated as follows: hs (Homo sapiens), mm (Mus musculus), ce (Caenorhabditis elegans), dm
(Drosophila melanogaster), ag (Anopheles gambiae), sp (Saccharomyces pombe), sc
(Saccharomyces cerevisiae), and ca (Candida albicans).This figure has been reproduced from
Aviv et al., 2003.
13
Smg Similarity Regions (SSR1 and SSR2). SSR1 functions as a dimerization domain (Tang et
al., 2007), while the function of SSR2 is unknown. The SSRs lie in the N-terminal region of the
protein, with SSR1 spanning amino acids 69-120 and the SSR2 spanning amino acids 199-287.
C-terminal to the SSRs and spanning amino acids 583-763, is Smg’s RNA-binding domain
(RBD), which contains a sterile alpha motif (SAM) domain (Dahanukar et al., 1999; Smibert et
al., 1999; Aviv et al., 2003; Green et al., 2003). Interestingly, while SAM domains have
traditionally been annotated to mediate protein-protein interactions, the Smg SAM domain
represents a novel RBD (Dahanukar et al., 1999; Smibert et al., 1999). A number of conserved
basic residues on the surface of the Smg SAM domain interact directly with RNA (Aviv et al.,
2003; Green et al., 2003). The Smg SAM domain is the defining feature for a family of post-
transcriptional regulators conserved from yeast to humans, of which Drosophila Smg was the
founding member (Figure 1) (Aviv et al., 2003).
Smg and its homologs exert their regulatory effects on target transcripts by binding to cis-
acting RNA elements termed Smg Recognition Elements (SREs) (Smibert et al., 1996, 1999).
SREs are stem-loop structures, with a loop sequence of CNGGN0-3 on a non-specific stem of at
least four base pairs (Aviv et al., 2003, 2006; Semotok et al., 2008). The crystal and NMR
structures of the SAM domain of the yeast Smg homolog, Vts1p, bound to RNA identified the
molecular interactions which underlie RNA binding of this protein family (Aviv et al., 2006;
Johnson and Donaldson, 2006; Oberstrass et al., 2006). First, a Watson-Crick base pair between
nucleotides 1 and 4 of the loop sequence (with a preference of nucleotide 1 for pyrimidines) is
necessary for the SRE to adopt the conformation required for binding by the SAM domain. Side
chains in the SAM domain form hydrogen bonds with the phosphate groups of four nucleotides
in the 5’ stem, as well as nucleotide 2 of the loop sequence (note that the interaction with
nucleotide 2 of the loop is independent of base-specificity). Finally, in a base-specific
interaction, the guanine of nucleotide 3 in the loop sequence hydrogen bonds to side chains in the
SAM domain (Aviv et al., 2006).
1.4.1 Mechanisms of Smg-mediated translation repression
When bound to target mRNAs, Smg can regulate different transcripts through different
means. One form of Smg-mediated regulation is translation repression, and work from various
groups have shown that this is accomplished by a number of mechanisms (Nelson et al., 2004;
14
Semotok et al., 2005; Jeske et al., 2006,2010; Zaessinger et al., 2006; Rouget et al., 2010; Pinder
and Smibert, 2013). One mechanism is through the recruitment of the eIF4E-binding protein,
Cup, to the mRNA (Nelson et al., 2004; Jeske et al., 2011). Once recruited, Cup interacts with
the cap-binding protein eIF4E. The mRNA/Smg/Cup/eIF4E interaction blocks the interaction of
eIF4E with eIF4G, which as described above, is a scaffold protein that is involved in recruitment
of the 40S ribosomal subunit to an mRNA, thereby inhibiting translation at the level of initiation
(Figure 2a) (Nelson et al., 2004; Macdonald, 2004; Andrews et al., 2011). The Cup model
however, is inconsistent with work showing repressed nos mRNAs associated with polysomes,
and argues for addition mechanisms for Smg-mediated translation repression (Clark et al., 2000).
Another mechanism of Smg-mediated translation repression involves Argonaute 1
(Ago1), a member of the Argonaute family of proteins (Pinder and Smibert, 2013). Argonaute
proteins are components of the RNA-induced silencing complex (RISC) and are typically
recruited to target mRNAs via association with small non-coding RNAs. In Drosophila,
recruitment of Ago1 to a target mRNA is mediated by miRNAs, and induces translational
repression and/or transcript degradation (Hutvagner and Simard, 2008). Interestingly Pinder and
Smibert (2013) showed Ago1 is required to repress nos translation and that Smg recruits Ago1 to
nos mRNA in a miRNA-independent fashion (Figure 2b).
The mechanisms of Smg-mediated translation repression have also been investigated
using in vitro translation extracts derived from Drosophila embryos (Jeske et al., 2006, 2011). In
this system, repression involves the formation of a stable repressor complex on a target mRNA
(Jeske et al., 2011). The formation of this complex is ATP-dependent and partially helped by the
presence of a poly(A) tail on the target transcript. Components of the repressor complex include
Smg, Cup, eIF4E, and subunits of the CCR4/POP2/NOT deadenylase complex (see below for
further description of the CCR4/POP2/NOT deadenylase complex), but exclude eIF4G.
Moreover, it blocks the assembly of the 48S pre-initiation complex in a cap-independent manner
(Jeske et al., 2011). Additionally, this complex also represses translation at a step post-initiation
through an as yet unknown mechanism, which may in part explain the association of repressed
nos mRNA with polysomes (Clark et al., 2000; Jeske et al., 2011).
Smg-binding can also regulate target mRNAs through its ability to trigger transcript
deadenylation (Semotok et al., 2005, 2008; Zaessinger et al., 2006; Jeske et al., 2006). Given the
15
importance of the poly(A) tail in translation and transcript stability, deadenylation can result in
translational inhibition and/or transcript decay. Smg-mediated deadenylation functions through
Smg’s ability to bind to and recruit the CCR4/POP2/NOT deadenylase to an mRNA in a
complex that is distinct from the Smg/Cup complex (Figure 2c) (Semotok et al., 2005;
Zaessinger et al., 2006). The CCR4/POP2/NOT deadenylase is highly conserved from yeast to
humans, is the major deadenylase in yeast, and has been shown to function as a Drosophila
deadenylase (Tucker et al., 2001; Temme et al., 2004; Semotok et al., 2005). Indeed, the
CCR4/POP2/NOT complex was shown in yeast to mediate deadenylation and degradation of
mRNAs targeted by the Smg homolog, Vts1p (Aviv et al., 2003) suggesting that this is a
conserved mechanism through which the Smg family regulates target mRNAs.
Additional work showed that Smg and the CCR4/POP2/NOT deadenylase interact with
Aubergine (Aub) and Ago3, two Argonaute family members involved in the piRNA pathway
(Rouget et al., 2010). This work also indicated that the degradation of Smg target mRNA
involves elements in the 3’UTR which represent binding sites for piRNAs bound by Aub and/or
Ago3. Based on these results, the authors suggested that efficient deadenylase recruitment
involves a Smg/Aub/Ago3 complex which makes contact with target mRNAs through SREs and
piRNA complementary sites within the mRNA’s 3’UTR.
1.4.2 Smg protein in the embryo
Smg is a maternally contributed component, deposited as mRNA into the oocyte, which
is translated beginning in the early embryo (Smibert et al., 1999; Tadros et al., 2007). This
translation activation of smg mRNA occurs in a process requiring the PAN GU kinase, and leads
to Smg protein accumulation (Tadros et al., 2007). During this time, Smg is ubiquitously
distributed throughout the embryo at high levels, where it acts on target mRNAs in the bulk
cytoplasm (Smibert et al., 1999). After the third hour of embryogenesis, Smg protein is degraded
from the bulk embryo (Smibert et al., 1996, 1999). This degradation may be important to allow
for the accumulation of zygotic proteins whose mRNAs contain SREs (Benoit et al., 2009). As
Smg protein is removed from the bulk embryo, it becomes localized to the posterior pole cells,
where it becomes enclosed in primordial germ cells after budding (Smibert et al., 1999; Siddiqui
et al., 2012).
16
Figure 2 – Known mechanisms of Smg-mediated regulation. Smg can regulate target mRNAs
by repressing their translation, and/or inducing their decay. A) In one mechanism, Smg recruits
the 4E-BP Cup to bound mRNAs, which in turn interacts with eIF4E. This complex blocks the
eIF4E/eIF4G interaction, which is required for recruitment of the 40S ribosomal subunit and
translation initiation. B) Smg can also recruit Ago1 to bound mRNAs in a miRNA-independent
manner, resulting in translational repression of the target mRNA. C) Smg has also been shown to
recruit the CCR4/POP2/NOT deadenylase complex to bound mRNAs. This results in poly(A)
tail shortening and leads to translational repression and/or transcript destabilization.
17
Distribution of Smg protein is generally diffuse, with a fraction forming foci in the bulk
of the embryo (Smibert et al., 1999; Zaessinger et al., 2006). In later stage embryos, Smg is
enriched in large foci at the posterior of the embryo, and have been suggested to be associated
with polar granules, large ribonucleoprotein structures involved in germ cell specification
(Smibert et al., 1999). Interestingly, the mammalian homolog, Smaug1, has also been shown to
form mRNA-silencing foci in neuron post-synapses (Baez et al., 2011). Despite these
observations, the exact nature of their formation and function are not well understood.
1.4.3 Targets of Smg-mediated regulation
There are two well-studied targets of Smg-mediated regulation: nos and Hsp83 mRNAs
(Smibert et al., 1996, 1999; Nelson et al., 2004; Semotok et al., 2005, 2008; Zaessinger et al.,
2006; Jeske et al., 2006, 2011; Rouget et al., 2010; Pinder and Smibert, 2013). Smg represses the
translation of nos mRNA in the bulk of the embryo through two SREs located in the transcript’s
3’UTR, but only plays a small role in regulating nos mRNA stability (Smibert et al., 1996, 1999;
Semotok et al., 2005; Semotok and Lipshitz, 2007). Unlike nos, Smg is only involved in
destabilization of Hsp83 mRNA, and not in repressing its translation (Semotok et al., 2005). The
Hsp83 mRNA contains eight predicted SREs distributed over the ORF, six of which are required
for its degradation in the early embryo (Semotok et al., 2008). Hsp83 is deposited maternally into
the late oocyte in a ubiquitous manner and its loading is independent of active transport (Ding et
al., 1993). In the first 3 hours of embryogenesis, Hsp83 undergoes Smg-mediated decay in the
bulk cytoplasm but is protected at the posterior by pole plasm components (Ding et al., 1993;
Semotok et al., 2005). This destabilization/protection leads to a localized pool of Hsp83 mRNA
at the posterior end of the embryo which is taken up by pole cells upon budding (Ding et al.,
1993).
Additional to nos and Hsp83, Smg also plays a major role in the destabilization of
maternal mRNAs in Drosophila embryos (Tadros et al., 2007). Approximately 35% of all
maternal mRNAs present in mature oocytes are destabilized following egg activation and during
the MZT (Tadros et al., 2007; Walser and Lipshitz, 2011). Two thirds of these unstable maternal
mRNAs are dependent on Smg for degradation. Thus, Smg-mediated decay of maternal mRNAs
is thought to facilitate the handover of developmental cues to the zygotic genome (Tadros and
Lipshitz, 2009). Moreover, these mRNAs dependent on Smg for decay are enriched in GO terms
18
related to cell cycle categories, protein or macromolecule catabolism (Tadros et al., 2007).
Specifically under the cell cycle categories are transcripts involved in DNA damage response
such as arrest, deadhead, loki, grapes, cyclins A and C, suggesting that Smg-mediated
degradation of maternal cell cycle mRNAs is essential for proper progression through the final
syncytial nuclear divisions during late stage MZT (Tadros et al., 2007).
1.4.4 Spatial regulation of Smg function
The translation of nos mRNA and the stabilization of Hsp83 mRNA at the posterior of
the embryo (Ding et al., 1993; Bergsten and Gavis, 1999) argues that Smg function must be
blocked at the posterior of the embryo. It has been suggested that the spatial regulation of Smg
activity involves its interaction with posteriorly localized Osk protein, as Osk blocks Smg’s
ability to bind mRNA in vitro (Dahanukar et al., 1999; Zaessinger et al., 2006). In this model,
any Smg target mRNAs found at the posterior of the embryo would escape Smg-mediated
repression. However, this model is inconsistent with the apparent requirement for additional cis-
elements in the nos mRNA which are necessary for nos translational activation (Dahanukar and
Wharton, 1996; Smibert et al., 1996).
1.5 Thesis rationale
Smg can employ several mechanisms to translationally repress and/or degrade target
mRNAs (Nelson et al., 2004; Tadros et al., 2007; Pinder and Smibert, 2013). To explore these
mechanisms further, I set out to perform a structure-function analysis of Smg. This approach
would allow me to assess the roles of the Smg N- and C-terminal sequences and the various
mechanisms in Smg function. It could also potentially allow me to uncover new regulatory
mechanisms that Smg can utilize. Moreover, it could shed light on why different Smg targets,
such as nos and Hsp83, are regulated via different mechanisms. Finally, a structure-function
analysis may elucidate the functional significance of Smg foci in the bulk of the embryo as well
as at the posterior.
To begin these experiments, I generated two smg protein null alleles, into which
transgenic smg proteins representing the N- and C-terminal regions of Smg were introduced.
Assessment of the function of these transgenic proteins suggests that Smg contains multiple
19
regions involved in repressing the expression of target mRNAs. Moreover, the function of these
Smg regions may be somewhat redundant with one another.
2 MATERIALS AND METHODS
2.1 Fly stocks and crosses
Drosophila melanogaster stocks used in this work included w1118
, smg1, and a deficiency
covering the smg gene Df(3L)ScfR6
(Dahanukar et al., 1999). GE21229 (GenExel) and w-;[Δ2,3
Sb ry506
]/TM6,Ubx stocks were used to generate the two smg excision alleles – smg30
and smg47
(see below). The smg30
and smg47
alleles were carried as smg30
/TM3,Sb and smg47
/TM3,Sb
stocks. smg30
and smg47
alleles were also crossed into Sp/CyO;Ly/TM3,Sb to generate
Sp/CyO;smg30
/TM3,Sb and Sp/CyO;smg47
/TM3,Sb stocks. Flies were maintained at 25°C for the
duration of all experiments unless otherwise specified.
2.2 P-element excision
Imprecise P-element excision was carried out using GE21229 (GenExel) and w-;[Δ2,3 Sb
ry506
]/TM6,Ubx stocks and published methods (Figure 3a, Hummel and Klämbt, 2008). Briefly,
GE21229 females were mated to w-;[Δ2-3 Sb ry
506]/ TM6,Ubx males (which carry the
transposase). Mobilization of the P-element (here on called P[excision]) in the F1 progeny was
screened based on mosaic eye colour, and F1 males with mosaic eyes were mated to Ly/TM3,Sb
virgin females. In the F2 progeny, 105 males showing white eyes (P[excision]/Ly or
P[excision]/TM3,Sb) were mated individually to Ly/TM3,Sb virgin females. Of the 105 crosses,
93 produced F3 progeny showing white eyes and stubble hairs (P[excision]/TM3,Sb) from which
fly lines were established. These lines were screened for maternal effect lethality, a characteristic
of the smg1 allele (Dahanukar et al., 1999). Homozygotes could not be established for 2 of the 93
lines, suggesting that they were homozygous lethal excisions. In contrast, 6 of the remaining 91
lines showed maternal effect lethality, suggesting they could carry excisions affecting the smg
gene.
2.3 Genomic DNA extraction and PCR
Genomic DNA was extracted from individual homozygous male flies (Gloor et al., 1993)
from each of the six lines showing maternal effect lethality. In separate 0.5ml tubes, whole flies
were homogenized in 50μl of squishing buffer (10mM Tris-Cl pH 8.2, 1mM EDTA, 25mM
20
NaCl, and 200μg/ml Proteinase K). The homogenate was then incubated at 25-37ºC for 20-30
minutes. After incubation, Proteinase K was inactivated by heating to 95°C for 1-2 minutes.
DNA samples were stored in -20°C until use.
To assay whether the excisions removed parts of the smg ORF, 3’UTR, the upstream
and/or the downstream genes, fragments in these regions were amplified using Pfx and a number
of primer sets. This initial round of PCR showed that only three of the six excisions did not
remove portions of either the genes upstream or downstream of the smg gene.
The general size of excision in the smg gene was assayed by a second round of PCR.
Here, the smg gene region was amplified using a single forward primer (annealing to the gene
upstream of the smg gene) in combination with a series of reverse primers annealing to
sequences in the smg gene. These primer combinations yield amplification products of ~4.5kb,
~5.5kb, and ~7kb using the wild-type genomic DNA as template (Figure 3b). The approximate
size of sequences deleted in the four smg excision alleles were identified based on size
differences between the amplification products of the wild-type genomic DNA and those of the
smg excision alleles.
2.4 Hatch rate analysis
A selection of 50 embryos laid overnight on apple juice agar plates were arranged and
aged for at least an additional 24 hours in 25°C. Embryos were then observed for hatching under
a Leica MZ6 modular stereomicroscope illuminated by a Volpi NCL 150 light source.
2.5 DAPI staining
Embryos collected 0-3 or 1-4 hours after egg-lay (AEL) were dechlorionated in 100%
bleach, washed well with water, and fixed for 20 minutes in equal volumes heptane and fixative
(4% formaldehyde in 1x PBS) with agitation. The embryos were devitellinized with methanol
and vortexing. Devitellinized embryos were transferred to eppendorfs, washed three times with
methanol, and rehydrated with PTW (0.1% Tween in 1x PBS). Embryos were mounted in
Vectashield Mounting Medium for Fluorescence with DAPI (Vector Laboratories, Inc.) and the
slides were stored at 4°C until viewing. Slides were observed at 10x objective on a Zeiss Axio
Imager.Z1 microscope using a DAPI reflector (440nm) and X-Cite Series 120 lamp source
21
Figure 3 – Generation of smg mutant alleles by imprecise P-element excision mutagenesis.
smg mutant alleles were generated using imprecise P-element excision mutagenesis. A) A brief
schematic of the technique. A source of transposase is introduced into female flies carrying a
non-autonomous transposable P-element (grey) in, or proximal to the gene of interest (black).
The transposase typically catalyzes the precise excision of the P-element (left), but in ~1% of
excision events, portions of genetic materials flanking the P-element are also excised. B) Shows
a brief schematic of the strategy used to map the approximate size of the excisions in the fly lines
where imprecise P-element excision occurred and was associated with maternal effect lethality.
The GE21229 (GenExel) P-element insertion (teal triangle) is shown relative to the smg gene
region (blue bar highlighted in pink, adapted from http://flybase.org). Four recovered alleles
carried deletions of the smg gene region, but did not affect the upstream or downstream genes.
The size of excisions were estimated by PCR of genomic DNA using a single forward primer
(red arrows) annealing to the gene upstream of the smg gene, in combination with three different
reverse primers (green) annealing to various sequences in the smg gene. These primer
combinations yield amplification products of ~4.5kb, ~5.5kb, and ~7kb with the wild-type smg
gene as DNA template (grey bars). A detailed description of this strategy is found in the body of
the results section.
22
(Lumen Dynamics). Photos were taken with a mounted Hamamatsu ORCA-ER C4742-80
camera running Volocity Imaging software v4.3.1 (Improvision).
2.6 Cuticle preparations
Embryo cuticles were visualized without the removal of the vitalline membranes, such
that smg mutant embryos – which do not develop cuticle structures – could be detected. Briefly,
flies were allowed to lay eggs for 3 hours on apple juice agar plates. The plates were set aside
and aged for an additional 24-36 hours at 25ºC before processing. Aged embryos were
dechlorionated in 100% bleach, washed well and collected into 0.1% Triton X-100 on ice.
Embryos were mounted in 5:3 Hoyers media: Lactic acid, and the slides warmed at 65°C
overnight. Slides were observed on a Nikon Eclipse E400 light microscope at 10x objective.
Photographs were taken with a mounted Hamamatsu ORCA-ER C4742-80 camera on a Zeiss
Axio Imager.Z1 microscope using dark-field illumination, and Volocity Imaging software v4.3.1
(Improvision).
2.7 Transgene construction
The FLsmg and NTsmg transgenes were generated by NU Siddiqui and HD Lipshitz and
served as template for my generation of the NTdSSR2 and CTsmg proteins. The base vector for
the construction of these transgenes was the smg5’UTR-BsiWI-smg3’UTR (SBS) plasmid (Tadros
et al., 2007). A linker carrying a start codon, the FLAG/p53 epitope tags and AscI and PmeI
restriction sites was inserted into the BsiWI site of SBS, between the smg UTRs, to generate a
modified SBS plasmid (SB’S). Genomic sequences of corresponding transgenic smg proteins
were inserted between the AscI and PmeI sites on the linker.
The FLsmg genomic transgene (encompassing the coding sequence for amino acids 1-
999) was amplified from a smg genomic rescue construct (Dahanukar et al., 1999) using a
5’primer with an AscI linker and a 3’primer with PmeI. The NTsmg genomic transgene
(encompassing the coding sequence for amino acids 1-766) was amplified using a 5’primer with
an AscI linker and a 3’primer with PmeI. The NTdSSR2 genomic transgene was generated by
excising an AvaI and XbaI genomic smg fragment – where the SSR2 had been deleted via quick-
change PCR – from a previously generated smg ΔSSR2 construct (plasmid A17, AL Orlowicz
and CA Smibert). The ΔSSR2 fragment was cloned into the corresponding position in the NTsmg
23
transgene. The CTsmg genomic transgene (encompassing the coding sequence for amino acids
583-999) was amplified from a smg genomic rescue construct, using a 5’primer with an AscI
linker and a 3’primer with PmeI. The ORF was inserted between the AscI and PmeI sites in the
linker of the SB’S plasmid.
Primers used for the generation of CTsmg genomic transgene are:
Sequence
Forward 5’-TAGGCGCGCCGAATTCAAGCCCAATTATATTAAGTTC -3’
Reverse 5’-TAGTTTAAACTTAGAATAGCGTAAAATGTTGATCAAATTTGGCC-3’
All genomic smg transgenes were then inserted into a pCaSpeR-4 cloning vector with an
attB site (Markstein et al., 2008; Tadros et al., 2007). Transgenic smg constructs were injected
into an attP40 landing site on the second chromosome (2L:25C7) (Markstein et al., 2008) by
Genetic Services (Cambridge, MA) using PhiC31, a site-specific integrase (Groth et al., 2004).
The inserted transgenes were then crossed into a smg47
mutant background to generate
transgene/CyO;smg47
/TM3,Sb stocks. Using the FLsmg transgene as an example, the flies used
in all experiments – unless otherwise specified – are of the genotype FLsmg;smg47
/smg47
and
were generated by mating FLsmg/FLsmg;smg47
/smg47
males to smg47
/TM3,Sb virgins, and
selected from the progeny.
2.8 Extract preparation and Western blotting
Embryos collected at various times AEL were dechlorionated in 100% bleach, washed
well with water and homogenized in a minimal volume of lysis buffer (150mM KCl, 20mM
HEPES-KOH pH 7.4, 1mM MgCl2, 1mM DTT, 1mM AEBSF, 2mM Benzamidine, 2μg/ml
Leupeptin and 2μg/ml Pepstatin A) with plastic pestle on ice. The lysate was centrifuged in 4°C
for 10 minutes at 8,000rpm and the supernatant stored at -80°C until use.
After thawing on ice, the equivalent of 5μg of total protein from each sample extract was
loaded into each lane on 8% or 10% SDS polyacrylamide gels. After electrophoresis, gels were
equilibrated in transfer buffer for 10 minutes on rocker and then transferred onto 0.2μm Protran
Nitrocellulose transfer membrane (PerkinElmer) at 100V for 45 minutes. Membranes were
incubated for 10 minutes in 1x PBS, then 20 minutes in 0.5% w/v milk (milk powder in PTW),
followed by incubation with primary antibody overnight at 4°C.
24
Membranes incubated with primary antibody were washed four times for 5 minutes with
PTW and incubated with secondary antibody for 2 hours at room temperature. Membranes were
then washed four times for 5 minutes with PTW and incubated with Amersham ECL Prime
Western Blotting Detection Reagent (GE Healthcare Life Sciences) as per manufacturer’s
protocol. Membranes were exposed for 10-30 seconds in a Bio-Rad Versadoc Imager and
analyzed on Quantity One software v4.6.6 (Bio-Rad).
Primary antibodies used were: guinea pig anti-Smg (1:10,000) (Tadros et al., 2007),
mouse anti-FLAG (1:5,000) (Sigma-Alrich), rabbit anti-BicC (1:5,000) (gift from Paul
MacDonald), and guinea pig anti-DP1 (1:5,000) (Tadros et al., 2007) diluted in 0.5% w/v milk.
Secondary antibodies anti-guinea pig-HRP, anti-mouse-HRP, and anti-rabbit-HRP (Jackson
ImmunoResearch) were used at a 1:5,000 dilution in 0.5% w/v milk.
2.9 RNA methods and RT-qPCR
RNA was extracted from embryos collected at 0-1, 1-2, 2-3, and 3-4 hours AEL.
Collected embryos were dechlorionated in 100% bleach, washed well with water and cold 0.1%
Triton X-100 on ice. Embryos were homogenized in 600μl TRI reagent (Sigma-Aldrich) with
plastic pestle. The lysate was centrifuged in 4°C for 10 minutes at 13,000rpm and the supernatant
was extracted with 1/5 the volume of RNase-free chloroform at room temperature for 5 minutes.
Aqueous layer was separated by centrifugation in 4°C for 15 minutes at 13,000rpm then
extracted a second time with equal volume of RNase-free chloroform. 4μl of glycogen
(Fermentas) was added to the aqueous phase and the RNA was precipitated with equal volume of
RNase-free isopropyl alcohol at room temperature for 10 minutes. The RNA was pelleted by
centrifugation in 4°C for 15 minutes at 13,000rpm and resuspended in 200μl of DEP-C treated
water. Remaining contaminants such as guanidinium chloride was further removed from the
RNA with 1/10 the volume of 3M NaAc pH 5.2 and 3 volumes of cold ethanol. The RNA was
pelleted again by centrifugation in 4°C for 12 minutes at 13,000rpm, resuspended in 20μl DEP-C
treated water, and then stored at -80°C until use. Concentration and purity of the RNA samples
were assessed using a NanoDrop 1000 spectrophotometer (Thermo Scientific).
After thawing on ice, the equivalent of 25ng of total RNA from each sample was used for
reverse transcription with SuperScript II reverse transcriptase (Invitrogen) and gene specific
25
primers (1pmol/gene/reaction). All reverse transcription reactions contained equi-molar amounts
of primer specific to the gene of interest and the loading control rp49. The primers used are as
follows:
mRNA targeted Sequence
Hsp83 5’-CATCGGAAGCGTTCGAGATCAA-3’
arrest 5’-CTTTAATGGCCGAAATGGCAGC-3’
BicC 5’-CGCAATACTCTCACAGGCGAAG-3’
rp49 5’-CGTTGTGCACCAGGAACTTCT-3’
A 10 fold dilution series (1:100 to 1:1,000,000) was generated for RT-qPCR using pooled
cDNAs from all time points of the same samples. Each cDNA sample was used at a dilution of
1:500 for the RT-qPCR. Reactions were carried out using Power SYBR Green PCR Master Mix
(Life Technologies) and 230nM each of the forward and reverse primers. The primer sets used
for each of the genes are as follows:
Gene targeted Sequence
Hsp83 (forward) 5’-ACAACAAGCAGCGTCTGAAAAG-3’
(reverse) 5’-CCTGGAATGCAAAGGTCTCTG-3’
arrest (forward) 5’- TGAACGCAAACTCTTTGTGG-3’
(reverse) 5’- GGCTCCGTGGACTTCAAATA-3’
BicC (forward) 5’-TCTCCACACCGCTGCTCATCT-3’
(reverse) 5’-GAGGTATGCAATTTTGGACGCG-3’
rp49 (forward) 5’-AGTCGGATCGATATGCTAAGCTG-3’
(reverse) 5’-AGTCGGATCGATATGCTAAGCTG-3’
RT-qPCRs were carried out in Hard-Shell Thin-Wall 384-Well Skirted PCR Plates (Bio-
Rad; Catalog# HSP3805) using a CFX384 Real-Time PCR Detection System (Bio-Rad) running
the following PCR program:
Step Temperature (°C) Time
Denaturation 95 10:00 minutes
Amplification
(40 cycles)
95 0:15 minutes
60 1:00 minute
95 0:15 minutes
Melting Curve Generation 60-95 in 0.5°C increments 1:00 minute/increment
RT-qPCR for each target mRNA was performed in triplicates for each of three biological
replicates. Data were analyzed using CFX Manager Software v3.0 (Bio-Rad).
26
2.10 SRE prediction
SREs are defined as stem-loop structures containing a loop sequence of CNGGN0-3 (N =
any nucleotide) on a non-specific stem of at least four base pairs (Aviv et al., 2006). SREs in the
BicC-RA sequence (defined at http://flybase.org) were identified by a prediction algorithm
(http://www.pathetique.com/craig/test2.html).
3 RESULTS
3.1 Generation of smg protein null alleles
The goal of my Smg structure-function analysis was to identify regions of Smg which are
critical to its function. As such, my analysis involved assaying the function of various mutant
smg transgenic constructs in a smg mutant background. At the time I began this work, the only
smg mutant allele available was smg1, generated by EMS mutagenesis (Dahanukar et al., 1999).
The mutation introduced a premature stop codon in the smg ORF, resulting in the translation of a
C-terminally truncated smg1 protein (Benoit et al., 2008). The smg
1 protein retains the SSR1 and
SSR2 domains, but this truncation removes the protein’s RNA-binding SAM domain and thus it
behaves as a loss-of-function allele (Dahanukar et al., 1999). The fact that the smg1 allele
expresses a truncated protein suggests that it is not ideal for use in a structure-function analysis
as this protein could affect the function of transgenic smg proteins. Indeed, the ability of the
SSR1 domain to function as a dimerization domain could result in the formation of dimers
between the protein encoded by the smg1 allele and transgenic smg proteins that carry the SSR1.
To avoid the possibility of such complications, I began this work by generating a smg protein
null allele by imprecise P-element excision.
P-elements are transposable DNA elements inserted into the chromosome, and have been
widely used to manipulate the Drosophila genome (Ryder and Russell, 2004). Typically, a P-
element is 2.9kb in size, harbouring 31 base pair inverse terminal repeats, and sequences
encoding a transposase necessary for its mobilization. The coding sequence of the transposase
spread over four exons (numbered 0, 1, 2, and 3), and complete splicing of the pre-RNA in the
germ line results in the expression of an 87kDa functional transposase protein (Hummel and
Klämbt, 2008). In somatic cells, the splicing of the intron between exons 2 and 3 is repressed,
resulting in the translation of a non-functional, truncated polypeptide (Robertson et al., 1988;
Hummel and Klämbt, 2008). Thus, transposition of P-elements is restricted to the germ line
27
(Hummel and Klämbt, 2008). Parts of the P-element may be lost through natural mutations or
technical manipulation, and in instances where sequences encoding the transposase are affected,
the element becomes non-autonomous (e.g. unable to mobilize itself) (Ryder and Russell, 2004).
However, non-autonomous elements can be mobilized with the introduction of an exogenous
source of transposase (Spradling and Rubin, 1982; Ryder and Russell, 2004).
Imprecise P-element excision is a mutagenesis technique that exploits the mechanism of
the transposase activity (Hummel and Klämbt, 2008). When a P-element is excised from the
genome through endonuclease activity, a double stranded break is generated in the DNA (Ryder
and Russell, 2004). The double stranded break leaves 3’ sticky ends that can be repaired such
that the wild-type sequence of the DNA is restored, resulting in a so-called “precise excision”
(Ryder and Russell, 2004; Hummel and Klämbt, 2008). In ~1% of cases, the sticky ends of the
double stranded break are degraded before repair occurs, leading to deletions that can remove
several kbps of the sequences flanking the P-element’s site of insertion, resulting in an imprecise
excision (Figure 3a) (Ryder and Russell, 2004; Hummel and Klämbt, 2008).
To generate a smg null allele, the non-autonomous P-element GE21229 (GenExel), which
is inserted 2,499 base pairs 5’ of the smg start codon (Figure 3b, 4 top panel), was excised and 93
independent excision lines were established. The smg1 allele shows maternal effect lethality (e.g.
flies which are homozygous for the smg1 allele are viable, but females lay eggs that do not hatch)
(Dahanukar et al., 1999). I was unable to isolate flies which were homozygous for the excised
chromosome for 2 of the 93 excision stocks, suggesting that the excisions that occurred in these
lines are lethal. Of the remaining 91 stocks, six gave homozygous females that showed maternal
effect lethality similar to the smg1 allele.
To assess if these six stocks carry deletions in the smg gene, genomic DNA was isolated
from individual homozygous males and analyzed by PCR (see Materials and Methods). Briefly,
an initial round of PCR showed the excision in three of the six maternal effect lethal lines did not
remove either of the genes upstream or downstream of the smg gene. A subsequent round of
PCR (Figure 3b) showed a decrease in expected size of amplification products in only two of the
excision lines, suggesting loss of large fragments of the smg gene. These two alleles were
designated smg30
and smg47
, and their deletions were precisely mapped by sequencing (Figure 4).
28
There are five isoforms of smg transcripts – designated RA through RD – defined by the
Flybase database (http://flybase.org/). Isoforms RA, RB, RC, and RE uses different transcriptional
start sites, but encode the same protein. The RD isoform uses the same start codon for translation
as the other four isoforms but is alternatively spliced, resulting in a longer ORF.
Sequencing revealed that the smg30
allele is a 4,514 base pair deletion of the smg gene
beginning 2,480 base pairs 5’ of, and ending 2034 base pairs 3’ of the smg start codon. Note that
the excision event left behind a 933 base pair fragment of the P-element insertion. The smg30
deletion removes 2,034 of 2,997 base pairs of the ORF in four of five smg isoforms (RA, RB, RC,
and RE) and 2,034 of 3,327 base pairs in the smg-RD isoform.
The smg47
allele is a 5,542 base pair deletion beginning 2,483 base pairs 5’ of, and ending
3,059 base pairs 3’ of the smg start codon. This deletion leaves 39 base pairs of the ORF in four
of five smg isoforms (RA, RB, RC, and RE) and 325 base pairs of the ORF in the smg-RD
isoform.
3.2 Characterization of the smg30
and smg47
alleles
The procedure used to isolate the smg30
and smg47
alleles suggested that embryos laid by
homozygous smg30
or smg47
mothers (hereafter referred to as smg30
and smg47
mutant embryos)
failed to hatch. To confirm this, smg30
and smg47
mutant embryos were assessed for hatching
(Table 1). As expected, smg30
and smg47
mutant embryos do not hatch (n=300 for each mutant
allele). Additionally, embryos laid by mothers heterozygous for smg30
or smg47
and either smg1
or a large deletion that removes a number of genes including smg (Df(3L)ScfR6
) also do not hatch
(n=300 for each genotype). In comparison, 82% of observed wild-type embryos (n=300)
hatched. Taken together, these data indicate that the maternal effect lethal phenotype associated
with the smg30
and smg47
alleles result from deletion of the smg gene.
Smg function is also required for embryos to progress normally through the syncytial
blastoderm stage of embryogenesis – when nuclei divide in the absence of cytokinesis (Benoit et
al., 2008; Dahanukar et al., 1999). After fertilization, the pronuclei undergo several rapid
divisions in the centre of the syncytium. Nuclei begin to migrate at cycle 8, all arriving in a
patterned array at the cortex of the embryo by cycle 10 (Sullivan and Theurkauf, 1995). Between
cycles 11 and 13 the length of interphase increases, reflecting checkpoint activation in a Smg-
29
Figure 4 – Significant portions of the smg gene is deleted in the smg30
and smg47
mutant
alleles. Two smg mutant alleles were generated using imprecise P-element excision, and termed
smg30
and smg47
. A) The GE21229 (GenExel) P-element is diagramed on a schematic of the smg
gene region (blue bar, adapted from http://flybase.org). B-C) Sequencing revealed the boundaries
of the deletions in the smg30
and smg47
alleles caused by imprecise P-element excision. B) The
smg30
mutant allele carries a 4,582 base pair deletion of the smg gene region, and retains a 933
base pair fragment of the original P-element (not diagramed) C) The smg47
mutant allele carries a
5,542 base pair deletion of the smg gene region. The deletions are diagramed in square brackets
over top of a schematic of the smg gene (blue bar), smg transcripts (isoforms RA through RE,
orange bars), and encoded proteins (isoforms RA through RE, purple bars) adapted from
http://flybase.org. The exact spans of the smg30
and smg47
deletions are described in the body of
the results section.
30
31
Genotype Number Hatched Percent Hatched n
wild-type 247 82% 300
smg30
/smg30
0 0% 300
smg30
/smg1 0 0% 300
smg30
/Df(scf) 0 0% 300
smg47
/smg47
0 0% 300
smg47
/smg1 0 0% 300
smg47
/Df(scf) 0 0% 300
Table 1 – Hatch rate analysis of smg30
and smg47
mutant embryos
32
dependent manner (Benoit et al., 2008).
In embryos laid by homozygous smg1 mothers (hereafter referred to as smg
1 mutant
embryos) syncytial nuclear divisions progress normally until cycle 11, when defects begin to
manifest and replication checkpoint activation fails (Dahanukar et al., 1999; Benoit et al., 2008).
Nuclear division defects include the appearance of gaps and disorganization in the cortical
nuclear array. To assess the progression of smg30
and smg47
mutant embryos through the
syncytial blastoderm stage, embryos were stained with the DNA dye DAPI. The division cycle of
wild-type, smg1, smg
30 and smg
47 embryos was estimated based on nuclear density, then
categorized into those at division cycles 10 and prior, division cycle 11, or division cycles 12 and
after (Figure 5). Embryos were then assigned a score of 1 through 5 based on the severity of
nuclear division defects where a wild-type pattern is given a score of 1. Embryos showing any
one of gap(s) or disorganization in the cortical nuclear array, or asynchronous nuclear division
were given a score of 2, while those showing a combination were given a score of 3. Embryos
which displayed aggregated nuclei but still retain some cortical nuclear array were given a score
of 4 and those whose nuclei have completely aggregated and collapsed – resembling the end
point of smg1 mutant embryos – were given a score of 5. It should be noted that embryos given a
score of 5 – where cortical nuclear density is not apparent – were considered to be at division
cycle 12 and after, based on published data (Dahanukar et al., 1999; Benoit et al., 2009).
No defects were detected in smg1 mutant embryos at or before division cycle 10 while all
cycle 11 embryos demonstrated some degree of division defect. At cycle 12 and beyond,
approximately three quarters of embryos showed defects considered to be level 4 or 5 in severity,
characterized by aggregated and collapsed nuclei (Figure 5b). The syncytial nuclear division
phenotypes for smg1 mutant embryos reported here are in agreement with those previously
published (Dahanukar et al., 1999; Benoit et al., 2009). DAPI analysis of smg30
and smg47
mutant
embryos (Figure 5c,d) showed that they had very similar defects to those seen in smg1 mutant
embryos, consistent with smg1, smg
30 and smg
47 all being strong loss-of-function alleles.
Accordingly, none of the smg30
and smg47
mutant embryos observed showed formation of
any cuticle structures or body segments, indicating early embryonic lethality. These smg mutant
embryos display as empty egg shells, same as that seen for smg1 mutant embryos (data not
shown). These results, taken together with sequencing results (see above) strongly shows that the
33
Figure 5 – Progression of syncytial nuclear divisions is defective in smg30
and smg47
embryos. Syncytial nuclear division was assayed in smg30
and smg47
mutant embryos using the
DNA dye DAPI. Embryos were divided into three division cycle categories: division cycle 10 or
prior, division cycle 11, and division cycles 12 and after. Scores of one to five were assigned to
embryos based on the severity of observable defects in syncytial nuclear division. The five levels
of defects are as follows: level 1 – wild-type; level 2 – any one of gap(s) or disorganization in the
cortical nuclear array, or asynchronous nuclear division; level 3 – showing a combination of
gap(s) or disorganization in the cortical nuclear array, or asynchronous nuclear division; level 4 –
aggregated nuclei but retaining some cortical nuclear array; and level 5 – complete nuclear
aggregation and collapse resembling a smg mutant phenotype. A) Wild-type embryos progress
through the syncytial nuclear divisions without any defect. B) Consistent with published data,
smg1 mutant embryos first show syncytial nuclear division defects at division cycle 11. All
embryos observed at this stage harbour some level of defect (levels two and three). At division
cycles 12 and beyond, all smg1 mutant embryos observed displayed syncytial nuclear division
defects, with a majority showing defects in the two most severe levels. C, D) Similar to the smg1
allele, smg30
and smg47
mutant embryos first show syncytial nuclear division defects at division
cycle 11. At division cycles 12 and beyond, all of the smg30
and smg47
mutant embryos observed
displayed syncytial nuclear division defects, with a majority showing defects in the two most
severe levels. The number of embryos observed for each division cycle category is shown on the
corresponding bar in each graph. E-K) Embryos at division cycles 12 and after, demonstrating
various levels of syncytial nuclear division defects. E) Embryo showing no defects (level 1). F)
Embryo showing disorganization of the cortical nuclear array (arrow, level 2). G) A close up of a
section of the embryo in F). H) Embryo showing asynchronous nuclear division (arrow, level 2).
I) A close up of a section of the embryo in H). J) Embryo showing aggregated nuclei but
retaining some cortical nuclei (level 4). K) Embryo showing complete nuclear aggregation and
collapse resembling a smg mutant phenotype (level 5). All embryos are orientated with the
anterior to the left and posterior to the right.
34
35
smg30
and smg47
alleles are smg protein-null mutants, which are ideal for use in a structure-
function analysis. As the sequencing results show that the smg47
allele contains the larger
deletion, it was used in the remainder of this work.
3.3 Generation of smg constructs and transgenic smg flies
The aim of my thesis work is to identify the regions in the Smg protein which are
important for its function. The structure-function approach I have taken to address this question
involves reconstitution of transgenic smg mutant proteins (Figure 6) into the smg47
mutant
background. In order to assess the functions mediated by the transgenic smg proteins without
introducing exogenous mRNAs targeting/binding systems, all of the smg proteins retained the
SAM containing Smg RBD.
Three smg proteins, FLsmg, NTsmg and CTsmg, were generated which represented the
Smg N- and C-terminal sequences (Figure 6). The FLsmg protein contained the full Smg protein
sequence (amino acids 1-999) and served as a control in my analyses. The NTsmg protein
contained the Smg protein sequences up to the end of the Smg RBD (amino acids 1-766), while
the CTsmg protein contained the Smg protein sequences between the start of the Smg RBD until
the end of the Smg protein (amino acids 583-999). Limited proteolysis and protein crystallization
experiments have shown that the Smg RBD is a fully folded domain (Green et al., 2003).
Therefore, defining the carboxy and amino boundaries of the NTsmg and CTsmg proteins,
respectively, based on the Smg RBD increases the chance that they will be expressed as stably
folded proteins.
To assess the role of the SSR2 domain, a third smg protein NTdSSR2 was generated in
which the SSR2 was removed from the NTsmg protein (amino acids 1-766Δ199-287). Work in
our lab (AL Orlowicz) has shown that an SSR2 deletion in the full length Smg protein does not
show a phenotype. This suggested that the SSR2 is either not essential for Smg function, or that
other sequences in the full length Smg protein are able to compensate for its loss. To ask whether
a role can be detected for the SSR2 in a sensitized smg mutant protein, the SSR2 was deleted
from the C-terminally truncated NTsmg protein. All of these smg proteins were N-terminally
tagged with a FLAG/p53 epitope to allow antibody detection and affinity purification. The
36
Figure 6 – A schematic of the proteins expressed by the transgenic constructs employed in
this study. Diagrams of wild-type Smg protein, the C-terminally truncated protein encoded by
the smg1 allele, and the transgenic smg truncation mutant proteins used in my thesis work. The
N-terminal FLAG/p53 epitope tags are diagramed as ovals while the SSR1, SSR2, and SAM
domains are shown as labeled rectangles. The residue numbers corresponding to the wild-type
Smg protein, which are present in the smg mutant proteins, are shown in parentheses.
37
FLsmg and NTsmg proteins were generated by NU Siddiqui in HD Lipshitz’s lab, while I
generated the CTsmg and NTdSSR2 proteins.
The smg mutant transgenes were generated based on a genomic smg rescue fragment
which contained the endogenous smg 5’ and 3’ regulatory elements (Dahanukar et al., 1999;
Semotok et al., 2005). Thus, the expression of these smg mutant proteins should be comparable
to endogenous Smg. To avoid variation in transgene expression associated with random insertion
of transgenes into the Drosophila genome, the PhiC31 integrase-mediated transgenesis system
(Bischof et al., 2007) was used to insert all transgenes into the attP40 landing site on the second
chromosome (Markstein et al., 2008). Expression of transgenes at the attP40 insertion site has
been shown to be free from positional effects (Markstein et al., 2008).
3.4 NTsmg and NTdSSR2 proteins are expressed at wild-type levels
Expression of the various smg proteins was assayed in embryos collected 0-3 hours AEL
– when Smg protein levels are at their peak (Smibert et al., 1996; Benoit et al., 2008) – by
Western blot using anti-FLAG or anti-Smg antibodies. As seen in Figure 7a and b, the FLsmg,
NTsmg and NTdSSR2 proteins are highly expressed at similar levels to each other, as well as
endogenous Smg protein in 0-3 hour embryos. Conversely, the CTsmg protein is barely
detectable in these extracts (Figure 7c). The CTsmg protein carries a deletion that removes both
intronic and exonic sequences, and as such, the low level of expression could result from
removal of a transcriptional enhancer or an element that stabilizes smg mRNA. Alternatively the
CTsmg protein might be unstable. In light of these expression data, any differences in the
function of the FLsmg, NTsmg or NTdSSR2 proteins is not likely due to a difference in the level
of protein that is expressed. Also, due to the low level expression of the CTsmg protein, it was
not included in my analysis unless otherwise stated.
3.5 NTsmg and NTdSSR2 proteins are expressed in later stage embryos
I next assayed the expression profiles of the smg proteins over the course of early
embryogenesis. Smg protein expression is regulated during the course of embryogenesis, with its
expression repressed in the oocyte until fertilization, when protein levels accumulate to a peak in
the second and third hour of embryogenesis (Smibert et al., 1996; Dahanukar et al., 1999; Tadros
et al., 2007; Benoit et al., 2009). In the fourth hour of embryogenesis, Smg protein is
38
Figure 7 – The FLsmg, NTsmg, and NTdSSR2 proteins are expressed at similar levels as
wild-type Smg. The transgenic smg mutant proteins were detected in extract prepared from
embryos collected 0-3 hours post egglaying. A) Detection of the FLsmg, NTsmg, and NTdSSR2
proteins by Western blot against the FLAG epitope show that they are expressed at similar
levels. In contrast, anti-FLAG antibodies did not detect any proteins in extract prepared from
wild-type embryos or smg47
mutant embryos. B) Detection of the wild-type Smg, FLsmg,
NTsmg, and NTdSSR2 proteins by Western blot using guinea pig anti-Smg antibody show that
the transgenic smg mutant proteins are expressed at wild-type levels. C) The CTsmg protein is
expressed at low levels compared to the NTsmg and NTdSSR2 proteins. Detection of the DP1
protein acted as a loading control for all Western blots.
39
degraded (Smibert et al., 1996; Tadros et al., 2007). To ask whether the smg proteins are also
expressed with the same profile as wild-type Smg, accumulation of the constructs was observed
over the first four hours of embryogenesis (Figure 8). Both wild-type Smg and the FLsmg
proteins were expressed in the first hour with peak levels in the second and third hours.
Subsequently both proteins were degraded and absent in the fourth hour of embryogenesis.
Strikingly, while expression of both the NTsmg and NTdSSR2 proteins in the first three
hours of embryogenesis resembles that of wild-type Smg, both proteins are detectable in the
fourth hour. The CTsmg protein – despite its low level expression – show an expression profile
similar to that of wild-type Smg; with amounts of protein increasing over the first three hours of
embryogenesis and gone by the fourth.
The expression profiles of the NTsmg and NTdSSR2 proteins show that they are not
properly regulated during embryogenesis. The persistence of these two proteins could be a result
of protein stabilization, or translation of stabilized mRNA. However, observations in our lab (see
Discussion) suggest that these N-terminal Smg proteins are stabilized, while the corresponding
mRNAs are not, suggesting that sequences in the Smg C-terminus are required to degrade Smg
protein.
3.6 NTsmg and NTdSSR2 do not rescue hatching defects
As described above, Smg protein is required for embryos to hatch, and thus I have
assayed the ability of a single copy of the smg transgenes to rescue the hatching defect of smg47
mutant embryos. As summarized in Table 2, hatching was restored in smg47
mutant embryos
rescued with a single copy of the FLsmg transgene to 81% (n=600), compared to 95% (n=300)
hatching for wild-type embryos. In contrast, 0% of the smg47
mutant embryos rescued with a
single copy of the NTsmg (n=600) or NTdSSR2 (n=600) transgenes hatched. Similarly, the
CTsmg transgene in a single copy also does not rescue the hatching in smg47
mutant embryos
(CA Smibert, personal communication). Thus, sequences in the C-terminus of the Smg protein
are required for wild-type Smg function.
3.7 NTsmg and NTdSSR2 attenuates nuclear division defects
Next, I assayed the ability of the FLsmg, NTsmg and NTdSSR2 proteins to rescue the
nuclear division defects of smg47
mutant embryos via DAPI staining (Figure 9). As expected,
40
Figure 8 – Stabilization of the NTsmg and NTdSSR2 proteins. The expression profiles of
wild-type Smg, FLsmg, NTsmg, NTdSSR2 and CTsmg proteins were observed in otherwise wild-
type embryos over the first four hours of embryogenesis. Wild-type Smg protein is expressed in
embryos collected 0-1 hours post egglaying and levels peak during the second and third hours of
embryogenesis. In the fourth hour, Smg protein is degraded and undetectable. Similar expression
profiles are also observed for the FLsmg and CTsmg proteins. Due to the low level of CTsmg
expression, the Western blot showing its expression profile has been over exposed to better
visualize the protein. Expression of the NTsmg and NTdSSR2 proteins in the first three hours of
embryogenesis resembles that of wild-type Smg. However, both of these proteins are detectable
in the fourth hour. Wild-type Smg was detected using an anti-Smg antibody while the smg
mutant proteins were detected using an anti-FLAG antibody. Detection of the DP1 protein acted
as loading control for all Western blots.
41
Genotype Number Hatched Percent Hatched n
wild-type 284 95% 300
smg47
/smg47
0 0% 300
FLsmg;smg47
/smg47
482 80% 600
NTsmg;smg47
/smg47
0 0% 600
NTdSSR2;smg47
/smg47
0 0% 600
Table 2 – Hatch rate analysis of smg47
mutant embryos rescued with a single copy of the
FLsmg, NTsmg, or NTdSSR2 transgenes
42
smg47
mutant embryos rescued with the FLsmg protein showed no syncytial nuclear division
defects through cycle 11, and 90% of those at cycles 12 and beyond showed no defects,
indicating rescue to near wild-type levels (Figure 9c).
In smg47
mutant embryos rescued with NTsmg protein (Figure 9d), 82% of those at
division cycle 11 showed no defects in syncytial nuclear division while 18% showed mild
defects (level 2). At division cycle 12 and beyond, 11% showed no defects while only 21%
showed defects considered levels 4 or 5 in severity. This is in contrast to smg47
mutant embryos,
of which 100% of those at cycle 11 had some level of defect, and 98% of those at cycle 12 and
beyond demonstrated severity levels 4 or 5 defects. Similarly for smg47
embryos rescued with the
NTdSSR2 protein (Figure 9e), 69% of those at division cycle 11 showed no defects in syncytial
nuclear division while 31% showed mild defects (level 2). For those embryos at division cycle
12 and beyond, 11% showed no defects and only 20% showed levels 4 and 5 defects. Taken
together, these data suggest that both the NTsmg and NTdSSR2 proteins are partially functional,
as they are able to attenuate the nuclear division defects associated with loss of Smg protein.
Moreover, these results also suggest that the Smg C-terminal sequences are required for wild-
type Smg function. Finally, the similarity in rescue provided by the NTsmg and NTdSSR2
proteins argues that the role played by the Smg N-terminal sequences in proper progression
through the syncytial nuclear divisions are largely independent of the SSR2.
3.8 NTsmg and NTdSSR2 proteins partially rescue cuticle formation
The smg proteins were next assessed for their ability to rescue the cuticle and body
segment formation defects that are found in smg47
mutant embryos. Following the syncytial
blastoderm stage, individual cortical nuclei become enclosed in plasma membrane to form a
single cell layer in a process called cellularization (Payre 2004; Loncar and Singer, 1995). In
concert with gradients of morphogens established in oogenesis and early embryogenesis, this
monolayer of epithelium organizes into segments along the antero-posterior body axis which
ultimately defines the body plan of the larvae (Nüsslein-Volhard and Wieschaus, 1980; Payre
2004). Eventually the cells comprising these segments secrete cuticle – a protective substance
containing proteins, chitin and lipids – which can be visualized (DiNardo et al., 1994; Payre
2004; Alexandre 2008; see Materials and Methods). The easily visible features of a wild-type
cuticle – on which my cuticle preparations are scored – include from the anterior to posterior: the
43
Figure 9 – NTsmg and NTdSSR2 proteins attenuate syncytial nuclear division defects found
in smg mutant embryos. Syncytial nuclear division was assayed in smg47
mutant embryos
rescued with the FLsmg, NTsmg, or NTdSSR2 proteins using the DNA dye DAPI. Embryos were
divided into division cycle categories, and assigned a severity score as previously described and
shown in figure 5. A, C) Wild-type embryos and smg47
mutant embryos rescued with the FLsmg
protein progress through the syncytial nuclear division defects with minimal defects. D) The
NTsmg protein reduced the syncytial nuclear division defect of smg47
mutant embryos. At
division cycle 11, there is a reduction in the percentage of smg47
mutant embryos which show
defects when rescued with the NTsmg protein. At division cycles 12 and beyond, smg47
mutant
embryos rescued with the NTsmg protein, a small fraction displayed no defects, and only one
fifth showed defects in the two most severe levels. E) The NTdSSR2 protein provided a level of
rescue similar to that observed for the NTsmg protein. Together, these results suggest that the
NTsmg and NTdSSR2 proteins are partially functional, that their roles in syncytial nuclear
division is independent of SSR2, and that the Smg C-terminal sequences are important for full
Smg function. The number of observed embryos (n) for each nuclear division cycle is indicated.
44
45
head skeletal structure, three thoracic segments, eight abdominal segments and the posterior
terminal segment called the telson (Lohs-Schardin et al., 1979; Jürgens et al., 1986; Nüsslein-
Volhard and Wieschaus 1980).
smg mutant embryos – whose development stops prior to cellularization – do not form
any cuticle structures or body segments (Dahanukar et al., 1999; Benoit et al., 2009; and Figure
10). For each genotype, 100 embryos were observed and scored into three groups: 1) those that
have formed all cuticle structures and body segments normally (wild-type cuticle), 2) those that
die during late stage embryogenesis, showing abnormalities in any of the cuticle structure and
body segment patterns (late stage embryonic lethal), and 3) those that die during early stage
embryogenesis, displaying a smg mutant phenotype where no cuticle structures form (early stage
embryonic lethal). As seen in Figure 10, 85% of wild-type embryos formed wild-type cuticle
while the remaining 15% did not form any cuticle structures (which are likely unfertilized eggs).
Similarly, 82% of smg47
mutant embryos rescued with the FLsmg protein developed wild-type
cuticle structures and body segments and the remaining 18% did not form any cuticle structures.
In contrast, 73% of smg47
mutant embryos rescued with the NTsmg protein were late
stage embryonic lethals while 27% embryos were early stage embryonic lethals. In comparison,
64% of smg47
mutant embryos rescued with the NTdSSR2 protein were late stage embryonic
lethals while 36% were early stage embryonic lethals. None of the observed smg47
mutant
embryos rescued with the NTsmg or NTdSSR2 proteins developed wild-type cuticles. Thus,
consistent with the nuclear division defect rescue assay, these data indicate that the NTsmg and
NTdSSR2 proteins are partially functional, and that the C-terminus of the Smg protein is also
required for wild-type protein function.
3.9 NTsmg and NTdSSR2 can mediate mRNA decay
One of Smg’s roles is to mediate the decay of maternal mRNAs in the early embryo
(Bashirullah et al., 1999; Tadros et al., 2007; Siddiqui et al., 2012), and as such I assayed the
ability of the NTsmg and NTdSSR2 proteins to induce the decay of three targets of Smg-mediated
mRNA degradation: Hsp83, arrest, and BicC, over the first four hours of embryogenesis. The
level of Hsp83, arrest, and BicC mRNAs were quantified by RT-qPCR in total RNAs harvested
at various times after egglaying from wild-type and smg47
mutant embryos, as well as smg47
46
Figure 10 – NTsmg or NTdSSR2 proteins partially rescue cuticle formation. A) Cuticle
preparations show that 85% of wild-type embryos develop normal cuticle structures and body
segments (B, wild-type cuticle), while 100% of smg47
mutant embryos showed no cuticle
formation (C, early embryonic lethal). For smg47
mutant embryos rescued with the FLsmg
protein, 82% displayed wild-type cuticle. NTsmg and NTdSSR2 proteins provided some rescue of
the cuticle defects associated with the smg47
mutant. 73% of the smg47
mutant embryos rescued
with the NTsmg protein, and 64% of those rescued with the NTdSSR2 protein developed
abnormal cuticle structures and body segments (D-G, late embryonic lethal). The remaining 27%
of the smg47
mutant embryos rescued with the NTsmg protein, and 36% of those rescued with the
NTdSSR2 protein showed early embryonic lethal. Abnormalities observed in late embryonic
lethals include D) abnormal head skeletal structure (indicated by the arrowhead), E) failure of
head involution and head closure (indicated by the *), compression of anterior structures, F)
absence of one or more body segments, and/or G) hole(s) in the body structure (indicated by the
*). All cuticles are orientated with the anterior to the left and posterior to the left. The number of
observed embryos is 100 for all genotypes.
47
48
mutant embryos rescued with the FLsmg, NTsmg or NTdSSR2 proteins. The stable rp49 mRNA
served as a loading control.
Consistent with published results (Bashirullah et al., 1999; Semotok et al., 2005), Hsp83
mRNA is degraded over this time course in wild-type embryos, while the mRNA is stable in
smg47
mutant embryos (Figure 11). When smg47
mutant embryos were rescued with the FLsmg
protein, Hsp83 mRNA decay was restored to wild-type kinetics. In smg47
mutant embryos
rescued with the NTsmg protein, Hsp83 mRNA decay was delayed during the first two hour of
embryogenesis after which degradation of the mRNA occurs. Interestingly, Hsp83 mRNA decay
in smg47
mutant embryos rescued with the NTdSSR2 protein did not show a delay, and the
decrease in the levels of Hsp83 mRNA between 0-1 and 1-2 hours AEL occurred at a rate similar
to that observed in wild-type embryos. At subsequent time points, Hsp83 mRNA degradation
continued, albeit at a reduced rate. Taken together, these data suggest that both the Smg N- and
C-terminal sequences are required to achieve wild-type degradation of Hsp83 mRNA. The C-
terminal sequences are required early in the process while the N-terminal sequences function
later. In addition, the SSR2 domain blocks the ability of the N-terminal sequences to induce
decay in early embryos.
I next assayed the decay of arrest mRNA. arrest is a maternally deposited mRNA
encoding the ovarian protein Bru, a translational regulator of osk mRNA (Kim-Ha et al., 1995;
Webster et al., 1997). The prediction of five SREs residing in the arrest ORF made it a likely
target of Smg-mediated regulation; its binding by Smg was confirmed by co-
immunoprecipitation and RT-qPCR (Siddiqui et al., in preparation). Smg regulates arrest mRNA
by repressing its translation in the posterior of the embryo, while inducing its degradation in the
bulk cytoplasm (Siddiqui et al., 2012; Siddiqui et al., in preparation).
Consistent with previous results, arrest mRNA is degraded in early embryos in a Smg-
dependent manner (Siddiqui et al., in preparation). In smg47
mutant embryos rescued with the
FLsmg protein, arrest mRNA degradation is similar to wild-type, with the exception of a very
modest delay in the degradation at the 1-2 hour time point (Figure 12). In smg47
mutant embryos
rescued with the NTsmg protein, decay of arrest mRNA was delayed over the first two hours of
embryogenesis, after which arrest mRNA decay initiates. A similar pattern of arrest mRNA
decay was also seen in smg47
mutant embryos rescued with the NTdSSR2 protein. This data is
49
Figure 11 – NTsmg and NTdSSR2 proteins can mediate Hsp83 mRNA decay. Hsp83 mRNA
decay mediated by the FLsmg, NTsmg, and NTdSSR2 proteins was assayed over the first four
hours of embryogenesis. Hsp83 mRNA levels were quantified by RT-qPCR from total RNA
harvested at 0-1, 1-2, 2-3, and 3-4 hour AEL from wild-type and smg47
mutant embryos, as well
as smg47
mutant embryos rescued with the FLsmg, NTsmg, and NTdSSR2 proteins. Consistent
with published data, Hsp83 mRNA is degraded in wild-type embryos, but stable in smg47
mutant
embryos. When smg47
mutant embryos were rescued with the FLsmg protein, Hsp83 mRNA
decay was restored to wild-type kinetics. In contrast, in smg47
mutant embryos rescued with the
NTsmg protein, Hsp83 mRNA was stabilized in the first two hours of embryogenesis, after which
decay initiated. In smg47
mutant embryos rescued with NTdSSR2, Hsp83 mRNA decay begins at
the start of embryogenesis without any delay, but the rate at later time points is slower than in
wild-type embryos. The stable rp49 mRNA served as an internal standard for data normalization.
50
Figure 12 – NTsmg and NTdSSR2 proteins can mediate arrest mRNA decay. arrest mRNA
decay mediated by the FLsmg, NTsmg, and NTdSSR2 proteins was assayed over the first four
hours of embryogenesis. arrest mRNA is degraded in wild-type embryos, but stabilized in smg47
mutant embryos. In smg47
mutant embryos rescued with the FLsmg protein, arrest mRNA decay
is similar to wild-type, with the exception of a very modest delay in degradation at the 1-2 hour
time point. In contrast, in smg47
mutant embryos rescued with either the NTsmg or NTdSSR2
proteins, the rate of arrest mRNA decay was slowed, such that more arrest mRNA is present at
the 2-3 hour time point. The stable rp49 mRNA served as an internal standard for data
normalization.
51
consistent with the argument that sequences in both the Smg N- and C-termini are required for
the timely degradation of mRNA. However, unlike for Hsp83, the SSR2 does not appear to play
any role in the degradation of arrest mRNA.
Finally, I assayed the decay of BicC mRNA. BicC mRNA encodes a 905 amino acid post-
transcriptional regulator involved in a variety of processes during oogenesis, including
cytoskeletal organization and polarity establishment (Mahone et al., 1995; Chicoine et al., 2007;
Gamberi and Lasko, 2012). BicC protein binds to target mRNAs via three noncanonical KH
domains and triggers deadenylation by recruiting the CCR4/NOT complex (Chicoine et al.,
2007; Gamberi and Lasko, 2012). Unpublished genome-wide work from the Smibert and
Lipshitz labs indicate that BicC mRNA is bound by Smg and is a likely target of Smg-mediated
regulation (LE Chen, J Dumelie, HD Lipshitz and CA Smibert, unpublished data). Moreover, a
search of the BicC mRNA identified four SREs in its ORF, and one more in its 3’ UTR (see
Materials and Methods). My analysis shows that the pattern of BicC mRNA decay is very similar
to that of arrest mRNA decay (Figure 13). As with Hsp83 and arrest mRNAs, BicC mRNA is
degraded over the first four hours of embryogenesis in wild-type embryos, but is stabilized in
smg47
mutant embryos. In smg47
mutant embryos rescued with the FLsmg protein, BicC decay is
largely restored, although with a minor delay at the 1-2 hour time point. In smg47
mutant
embryos rescued with either the NTsmg or NTdSSR2 proteins, BicC mRNA decay was delayed in
the first two hours of embryogenesis, after which decay takes place. Although levels of BicC
mRNA during the 1-2 hour time point in smg47
mutant embryos rescued with NTsmg was higher
than compared to those rescued with the NTdSSR2 protein, BicC mRNA was degraded to the
same level by the fourth hour of embryogenesis for both. Similar to the arrest decay, these data
suggest that both the Smg N- and C-termini, but not the SSR2, play roles in the degradation of
BicC mRNA.
While these data suggest that the Smg C-terminus is required for early decay of Hsp83,
arrest, and BicC mRNAs, the failure of FLsmg to fully rescue decay of arrest and BicC mRNAs
in the first two hours of embryogenesis makes it difficult to draw a clear conclusion.
Nonetheless, these results strongly argue that both the Smg N- and C-termini contribute to Smg’s
ability to fully degrade Hsp83, arrest, and BicC mRNAs. It is unclear whether both the Smg N-
and C-termini induce mRNA decay through deadenylation, or if another mechanism of mRNA
52
Figure 13 – NTsmg and NTdSSR2 proteins can mediate BicC mRNA decay. BicC mRNA
decay mediated by the FLsmg, NTsmg, and NTdSSR2 proteins was assayed over the first four
hours of embryogenesis. BicC mRNA decay mediated by the transgenic smg mutant proteins is
similar to that observed for arrest mRNA. BicC mRNA is degraded in wild-type embryos, but
stabilized in smg47
mutant embryos. In smg47
mutant embryos rescued with the FLsmg protein,
BicC mRNA decay is largely restored, but with a minor delay at the 1-2 hour time point. In smg47
mutant embryos rescued with either the NTsmg or NTdSSR2 proteins, the rate of BicC mRNA
decay was slowed, such that more BicC mRNA is present at the 2-3 hour time point. The stable
rp49 mRNA served as an internal standard for data normalization.
53
degradation such as endonucleolytic or exonucleolytic degradation is involved. Interestingly, in
the absence of the Smg C-terminus, the SSR2 appears to play a role in stabilizing only Hsp83
mRNA in early embryogenesis, but not arrest and BicC mRNAs. Note that any differences
related to the decay of these three mRNAs do not relate to variables such as embryo staging, as
the levels of all three mRNAs were assayed using the same RNA samples. Thus, I propose that
this difference might reflect different cis-elements in Hsp83, arrest, and/or BicC mRNAs which
influence the mechanisms which underlie their degradation (see Discussion for further details).
3.10 Two copies of the NTsmg and NTdSSR2 transgenes enhance rescue of the smg
mutant phenotype
Although a single copy of the NTsmg or NTdSSR2 transgenes were not able to rescue the
hatching defects in smg47
mutant embryos, subsequent analyses showed that they were able to
provide some Smg function. To investigate whether two copies of the N-terminal smg transgenes
might provide a greater level of rescue to the smg47
mutant phenotype, I analyzed embryos laid
by smg47
mutant mothers carrying two copies of the various smg transgenes.
The ability of two copies of the smg transgenes to rescue hatching of smg47
mutant
embryos was assayed first (Table 3). As expected, smg47
mutant embryos rescued with two
copies of the FLsmg transgene hatched (88%, n=1,150). Strikingly, smg47
mutant embryos
rescued with two copies of either the NTsmg or NTdSSR2 transgenes showed hatch rates of 80%
(n=1,200) and 75% (n=1,150), respectively. These hatch rates provide further evidence that the
NTsmg and NTdSSR2 proteins are to some extent functional.
I next assessed the cuticle phenotypes of smg47
mutant embryos rescued with two copies
of the NTsmg transgene (Figure 14). Consistent with the results of the hatch rate experiments, an
increase in the dose of the NTsmg transgene decreases the severity of the cuticle phenotypes in
smg47
mutant embryos compared to a single copy of the transgene. Strikingly, 44% of these
embryos displayed a new category of cuticle phenotype, in which they have hatched as L1
larvae, and display largely wild-type cuticles with the exception of a missing third abdominal
denticle belt (L1 larvae w/ abdominal segment defects). This result is consistent with the
increased hatching of smg47
mutant embryos rescued with two copies of the NTsmg transgene.
Finally, I assessed the syncytial nuclear division defects in smg47
mutant embryos rescued
54
Genotype Number Hatched Percent Hatched n
wild-type 1064 89% 1200
FLsmg/FLsmg;smg47
/smg47
1009 88% 1150
NTsmg/NTsmg;smg47
/smg47
955 80% 1200
NTdSSR2/NTdSSR2;smg47
/smg47
863 75% 1150
NTsmg/CTsmg;smg47
/smg47
5 0.42% 1200
Table 3 – Hatch rate analysis of smg47
mutant embryos rescued with two copies of the
NTsmg transgene, or co-expression of the NTsmg and CTsmg transgenes
55
Figure 14 – Enhanced rescue of the cuticle phenotype of smg47
mutant embryos by
increased copy number of the NTsmg transgene, or co-expression of NTsmg and CTsmg. A)
Two copies of the NTsmg transgene enhanced rescue of the cuticle phenotype of smg47
mutant
embryos. Notably, 27% of smg47
mutant embryos rescued with two copies of the NTsmg
transgenes observed developed wild-type cuticles, versus 0% in smg47
mutant embryos rescued
with only a single copy of the NTsmg transgene. Moreover, 44% of smg47
mutant embryos
rescued with two copies of the NTsmg transgenes displayed a new phenotype (C, L1 larvae with
abdominal segment defects), in which they hatch as L1 larvae and displayed largely wild-type
cuticles with the exception of a missing third abdominal denticle belt. In contrast, expression of
the CTsmg transgene in addition to the NTsmg transgene provided a modest improvement to the
rescue of the cuticle phenotype of smg47
mutant embryos, over just a single copy of the NTsmg
transgene. None of the smg47
mutant embryos rescued with co-expression of the NTsmg and
CTsmg transgenes formed wild-type cuticle, and 89% developed deformed cuticle structures and
body segments (late embryonic lethal). This is in contrast to the 73% of smg47
mutant embryos
when rescued with only a single copy of the NTsmg transgene. Additionally, 2% of smg47
mutant
embryos rescued with co-expression of the NTsmg and CTsmg transgenes demonstrated the L1
larvae w/ abdominal segment defects phenotype. This modest improvement to the rescue of the
cuticle phenotype of smg47
mutant embryos provided by co-expression of the NTsmg and CTsmg
transgenes over just a single copy of the NTsmg transgene may be an underestimation given the
low level expression of the CTsmg protein. B) Wild-type cuticle showing a head skeleton, eight
denticle belts, the telson and filzkörper. C) The L1 larvae w/ abdominal segment defects
phenotype shown by 44% smg47
mutant embryos rescued with two copies of the NTsmg
transgene. The white arrowhead denotes the missing third denticle belt. The cuticles are
orientated with the anterior to the left and posterior to the right.
56
57
with two copies of the NTsmg transgene via DAPI staining and scoring the phenotypes as
categorized above (Figure 15d). Interestingly, while two copies of the NTsmg transgene gives a
similar rescue to syncytial division defects at division cycle 11 as a single copy, at subsequent
stages the increased dose of the NTsmg transgene provides a greater rescue.
Taken together, these phenotypic assays indicate that two copies of the NTsmg transgene
provide greater rescue to the smg mutant phenotype than does a single copy. Moreover, the hatch
rate and cuticle data strongly suggests that an increased dose of the NTsmg protein is largely
capable of mediating the Smg functions required to complete embryogenesis. This argues that
there may be some degree of redundancy in the functions of the Smg N- and C-termini, and that
the loss of the Smg C-terminus may be compensated for by increased expression of the N-
terminus.
3.11 Co-expression of NTsmg and CTsmg offer modest improvement over a single copy of
NTsmg alone
The data described above suggest that the Smg N- and C-termini might have somewhat
redundant functions. To further explore this issue, I assessed the ability of the CTsmg protein to
enhance the ability of NTsmg to rescue the smg mutant phenotype. I first assayed the hatch rate
of smg47
mutant embryos rescued with a single copy each of the NTsmg and CTsmg transgenes
and found a very small number of these embryos (5 of 1,200 assayed) hatched (Table 3).
Whether this represents a significant difference from smg47
mutant embryos rescued with a
single copy of the NTsmg transgene, where 0 of 600 embryos assayed hatched, is unclear.
I next asked whether the CTsmg protein could enhance the ability of the NTsmg protein to
rescue the syncytial nuclear division defects associated with the smg mutant phenotype via DAPI
staining. Interestingly, the co-expression of both transgene showed a modest improvement in the
syncytial nuclear division defects over smg47
mutant embryos rescued with a single copy of the
NTsmg transgene (Figure 15e). This improvement was noted in embryos at nuclear division cycle
12 and beyond. Consistent with this modest effect, I also detected a modest improvement in the
rescue of the cuticle defects in smg47
mutant embryos carrying both the NTsmg and CTsmg
transgenes compared to a single copy of the NTsmg transgene by itself (Figure 14).
Taken together, these data suggest that despite the low level expression of the CTsmg
58
Figure 15 – Improved rescue of the syncytial nuclear division defects of smg47
mutant
embryos by increased copy number of the NTsmg transgene, or co-expression of NTsmg
and CTsmg. Analyses of syncytial nuclear division by DAPI show that two copies of the NTsmg
transgene further attenuates the nuclear division defect of smg47
mutant embryos compared to
those with just a single copy of the transgene. D) At division cycle 12 and beyond, two copies of
the NTsmg transgenes provided a greater rescue to the syncytial nuclear division defects of smg47
mutant embryos rescued with a single copy of the NTsmg transgene, such that a majority of them
displayed no defects. Additionally, none of these embryos showed defects in the two most severe
levels. E) Expression of the CTsmg transgene in addition to the NTsmg transgene also modestly
improved the rescue of the nuclear division defects of smg47
mutant embryos over just a single
copy of the NTsmg transgene. At division cycle 12 and beyond, an increased percentage of
embryos – two fifths – of these embryos displayed no defects, while none showed defects in the
most severe level.
59
60
protein, it is somewhat functional. Additionally, the level of rescue in smg47
mutant embryos
expressing both the NTsmg and CTsmg transgenes is likely under represented and reflect the low
level expression of the CTsmg protein.
4 DISCUSSION
4.1 Summary and Conclusions
4.1.1 Generation of smg protein null flies
Here I have reported the generation of two protein null smg alleles – smg30
and smg47
–
using imprecise P-element excision. Significant portions of the smg gene were excised in both
alleles, and my analyses indicate that both show very similar phenotypes to the published smg1
mutant allele (Dahanukar et al., 1999). The smg30
and smg47
mutant alleles are considered to be
protein null alleles given the extent of their excisions. This makes both alleles ideal for use in my
structure-function analysis, as it reduces the potential for interactions of the protein expressed by
the smg1 allele with my transgenic smg mutant proteins. In addition, the similarity in the
phenotypes displayed by the smg1, smg
30 and smg
47 alleles suggest that the protein expressed by
the smg1 allele is not functional, consistent with the fact that this protein does not have the Smg
RBD.
4.1.2 Both the Smg N- and C-termini are important for Smg function
My analyses demonstrate that both the N- and C-termini of Smg are important for the
protein’s function. For example, while a single copy of the NTsmg transgene does not rescue the
hatching of smg47
mutant embryos, the transgenic protein is partially functional, based on the
results of the nuclear division and cuticle phenotype assays. The failure of the NTsmg protein to
fully rescue the smg mutant phenotype indicates that the C-terminal region also plays an
important role. Consistent with this, the CTsmg transgene is able to enhance the rescue mediated
by a single copy of the NTsmg transgene. In addition, the robust rescue of smg47
mutant embryos
provided by two copies of the NTsmg transgene suggests that the N- and C-termini are partially
redundant with one another. As such, the increased expression of the Smg N-terminus can
compensate for the loss of the C-terminus. This model is supported by the fact that both the N-
and C-terminal regions of Smg function in transcript decay, as discussed below.
61
4.1.3 Smg employs multiple mechanisms to induce transcript decay
Assaying the stability of Smg target mRNAs in smg47
mutant embryos rescued with the
NTsmg protein shows that both the N- and C-terminus of Smg participate in the degradation of
target mRNAs. The results suggest that Smg employs at least two mechanisms to induce
transcript decay. Additionally, the Hsp83 mRNA decay data further suggest that the mechanism
employed by the C-terminus functions at early time points while the mechanisms employed by
the N-terminus functions at later time points.
As described above, I propose that the roles that both the Smg N- and C-terminal regions
play in transcript decay underlies the ability of two copies of the NTsmg transgene to provide
robust rescue of the smg mutant phenotype. When the NTsmg transgene is only present in a
single copy, the resulting defect in mRNA decay leads to embryonic lethality. In contrast, when
two copies of the NTsmg transgene are present, the resulting increase in the levels of the NTsmg
protein enhances the extent of mRNA decay, such that the embryonic lethality of smg47
mutant
embryos is reduced. Similarly, CTsmg could also enhance mRNA decay, albeit to a lesser extent
due to its low level expression. Therefore, co-expression of the CTsmg transgene with the NTsmg
transgene enhances the rescue of smg47
mutant embryos more than just a single copy of the
NTsmg transgene alone.
What is the nature of the N- and C-terminal sequences of the Smg protein in mediating
transcript decay? Previous work proposed a model where Smg induces the degradation of Hsp83
mRNA through Smg’s ability to bind and recruit the CCR4/POP2/NOT deadenylase to the
bound mRNA. My work here on mRNA decay expands upon this model with two possible
modes of CCR4/POP2/NOT deadenylase recruitment. In one possibility, both the Smg N- and C-
terminal sequences can interact with the CCR4/POP2/NOT deadenylase. Thus, the loss of the C-
terminal region could reduce the stability of the Smg/deadenylase complex, leading to reduced
efficiency of deadenylase binding, resulting in a delay in the decay of Smg target mRNAs. In
another possibility, one of the Smg N- or C-terminal regions could induce decay via recruitment
of the CCR4/POP2/NOT deadenylase while the other region recruits another factor able to
induce transcript decay. Candidates for this other factor include the other deadenylases, or
factors which promote transcript decapping.
62
4.1.4 The SSR2 domain plays a transcript-specific role in Smg function
The results of my mRNA decay assays show that deletion of the SSR2 domain alleviates
the early delay in Hsp83 mRNA decay associated with the removal of the Smg C-terminus.
Conversely, the SSR2 does not appear to play a role in the decay of arrest and BicC mRNAs.
Thus, Smg protein appears to be subject to SSR2-mediated auto-regulatory control, at least with
regards to the decay of one of its target mRNAs. The fact that the auto-regulatory function of the
SSR2 is transcript-specific suggests that its function is regulated by other factors which interact
with the target mRNA. For example, Hsp83 mRNA could be bound by a factor which interacts
with SSR2 to block transcript decay mediated by the Smg N-terminus. Alternatively, arrest and
BicC mRNAs could be bound by a factor which interacts with SSR2 to block the inhibitory
activity of the SSR2.
Interestingly, auto-regulatory effects have been demonstrated for two conserved motifs in
the post-transcriptional regulator protein Pum, in Drosophila (Weidmann and Goldstrohm,
2012). Together, the two motifs – termed Pumilio Conserved Motifs a and b (PCMa and PCMb)
– work to modulate the translation repression function of the Pum N-terminus. PCMb negatively
regulates the translation repression activity of the Pum N-terminus, but is itself antagonized by
PCMa through auto-inhibitory interactions (Weidmann and Goldstrohm, 2012).
4.1.5 The Smg C-terminus functions in Smg protein degradation
Previous work has shown that sequences in the C-terminal half of Smg are required for
the rapid degradation of Smg protein three hours AEL (W Tadros and HD Lipshitz, personal
communication). This conclusion is based on the persistence of the C-terminally truncated Smg
protein encoded by the smg1 allele. This truncated protein results from a premature stop codon at
codon 479 within the smg ORF, such that all of the wild-type smg mRNA sequences – including
regulatory elements – remain intact. Thus, the persistence of the smg1 protein likely reflects
stabilization of the protein, rather than the mRNA. My work has shown that the NTsmg protein,
whose C-terminal boundary is at amino acid 766, also persists after the third hour of
embryogenesis. Moreover, Northern blot analysis of NTsmg mRNA demonstrates that the
mRNA is degraded with wild-type kinetics (A Marsolais, personal communication). Together,
these data suggest that the protein sequences downstream of amino acid 766 are required for the
degradation of the Smg protein. In addition, since the stabilization of the smg1 protein is more
63
robust than that observed for the NTsmg protein, this suggests that sequences between amino
acids 479 and 766 also contribute to the destabilization of the Smg protein.
While the mechanism for Smg protein degradation is not currently known, the
Drosophila homolog of the MAST kinase Drop Out (Dop) has been implicated in this process
(BD Pinder, CA Smibert, and HA Muller, unpublished data). In addition, Dop co-
immunoprecipitates with the full length Smg protein, while I have preliminary data which
indicate that Dop does not co-immunoprecipitate with the smg1 protein. This maps the Dop
binding site between amino acids 479-999 (e.g. the sequence which is missing from the smg1
protein) and establishes a correlation between the Dop/Smg interaction and degradation of the
Smg protein. Furthermore, a phosphoproteome study of the Drosophila embryo (Zhai et al.,
2008) identified five potential sites for phosphorylation present in Smg. They include three
serines just N-terminal to the SAM domain, and two serines at the C-terminus of the protein.
Although it is unknown whether phosphorylation of Smg is indeed necessary for its degradation,
it is interesting to note that none of the phospho-serines identified by Zhai et al. (2008) are
present in the smg1 protein. In addition, stabilization of the NTsmg protein – which retains the
three phospho-serines N-terminal to the SAM domain – is not as robust as the smg1 protein.
4.2 Future Directions
4.2.1 The role of NTsmg in mRNA decay
My hatching assays show a robust rescue of smg47
mutant embryos provided by two
copies of the NTsmg transgene over a single copy. Given that both the Smg N- and C-termini
participate in mediating the decay of Smg target mRNAs, it is possible that the increase in
NTsmg protein from two copies of the transgene might enhance the extent of mRNA decay
mediated by NTsmg. Such an enhancement could in turn compensate for the impaired mRNA
decay associated with a loss of the Smg C-terminus, and reduce embryonic lethality in smg47
mutant embryos. To ask whether increased expression of the NTsmg protein can restore mRNA
decay to near wild-type levels, I would assay the decay of Hsp83, arrest, and BicC mRNAs in
smg47
mutant embryos rescued with two copies of the NTsmg transgene.
If these assays show that Hsp83, arrest, and BicC mRNAs are destabilized to a greater
extent than in smg47
mutant embryos rescued with a single copy of the NTsmg transgene, it
64
would support the hypothesis in which two copies of the NTsmg transgene can reduce embryonic
lethality by compensating for the loss of mRNA decay associated with the Smg C-terminus.
Moreover, such a result would further strengthen the notion that the roles of the Smg N- and C-
termini in mRNA decay are at least partially redundant and overlap with one another.
In contrast, negligible difference in the decay of Hsp83, arrest, and BicC mRNAs by two
copies of the NTsmg transgene over just a single copy would contradict the proposed hypothesis.
Furthermore, this would suggest that increased NTsmg protein may be reducing embryonic
lethality in smg47
mutant embryos via other mechanisms and/or functions. For example, an
enhancement in translation repression by increased levels of the NTsmg protein may compensate
for the loss in mRNA decay associated with the Smg C-terminus, reducing embryonic lethality
of smg47
mutant embryos.
4.2.2 The mechanism of NTsmg function in mRNA decay
Although my analyses have shown that the Smg N-terminus is able to mediate mRNA
decay, the mechanism through which it functions is unclear. As previously mentioned, both the
Smg N- and C-termini may bind and recruit the CCR4/POP2/NOT deadenylase to a target
mRNA to induce deadenylation. An alternative model could be that only one of the Smg N- or
C-termini binds and recruits the CCR4/POP2/NOT deadenylase, while the other mediates
transcript decay through another mechanism. To test whether the Smg N-terminus mediates
transcript decay via deadenylation, I would first assess deadenylation of Hsp83, arrest, and BicC
mRNAs in smg47
mutant embryos rescued with NTsmg using an RNase H cleavage assay. If
these mRNAs are not deadenylated, it would suggest that NTsmg mediates transcript decay via a
mechanism separate from deadenylation (see below for further details). Alternatively, if the
mRNAs are deadenylated, it would suggest that the Smg N-terminus induces transcript decay
through deadenylation. This result would allow me to proceed to identify the deadenylase
through which the Smg N-terminus functions.
There are two known deadenylases in Drosophila, the CCR4/POP2/NOT and Pan2-Pan3
deadenylases. If NTsmg induces transcript deadenylation, which of these deadenylases does the
NTsmg protein function through? To answer this question, I will test whether either of the
CCR4/POP2/NOT or Pan2-Pan3 deadenylases co-immunoprecipitates with NTsmg by
65
purification of the NTsmg protein followed by Western blots using antibodies against the
deadenylases. If the CCR4/POP2/NOT deadenylase co-immunoprecipitates with NTsmg, then it
would argue that the Smg N-terminus induces transcript decay through a deadenylation
mechanism involving the CCR4/POP2/NOT deadenylase. Alternatively, if the CCR4/POP2/NOT
deadenylase does not co-immunoprecipitate with NTsmg, it would imply that it is the Smg C-
terminus which mediates deadenylation through the CCR4/POP2/NOT deadenylase. Ultimately,
I will attempt to precisely map the region of the Smg N-terminus which bridges the
Smg/deadenylase interaction by assessing which domains and motifs within the NTsmg protein
are required for the interaction by generating additional smg mutant proteins.
There are three domains in the Drosophila Smg protein which are also conserved in the
mouse and human homologs: SSR1 and SSR2, as well as the SAM domain (Smibert et al.,
1999). Additionally, there are six motifs which are identified by a PSI-BLAST search to be
conserved among all Drosophilids, and several species of mosquitoes and insects (Figure 16).
Four of these motifs are N-terminal of the Smg SAM domain, while two are C-terminal of the
Smg SAM domain. To investigate whether the SSRs or the four N-terminal conserved motifs are
involved in mRNA decay and/or deadenylase recruitment by NTsmg, I will first generate
transgenic smg mutant proteins where the SSRs and the four conserved motifs are deleted
individually or in combination from the NTsmg protein. These mutant proteins will allow me to
assay the roles of these domains and motifs in mRNA decay and deadenylation.
Deletions of the SSRs and/or conserved motifs from NTsmg may not show any additional
effects on mRNA decay over NTsmg. While this would suggest that the SSRs or conserved
motifs are not involved in Smg-mediated mRNA decay, it could indicate their roles in other Smg
functions such as translation repression (see below for further details). However, if deletions of
one or more of the SSRs and/or conserved motifs in NTsmg further affect mRNA decay, I would
assay whether deadenylase binding with the NTsmg protein is abolished in these mutants.
Defects in mRNA decay, deadenylation and deadenylase binding upon mutation of a specific
motif or domain would strongly argue that NTsmg induces transcript decay through deadenylase
recruitment.
An alternative result from the RNase H cleavage assay proposed earlier is that the Hsp83,
arrest, and BicC mRNAs are not deadenylated by the NTsmg protein. This result would suggest
66
Figure 16 – Six additional motifs in the Smg protein are conserved among Drosophilids and
some insects. A PSI-BLAST search of the Smg protein sequences excluding the SSRs and SAM
domain revealed six motifs conserved among Drosophilids and some insects. Additionally, a
phosphoproteome study of Drosophila melanogaster embryos identified five potential phospho-
serines in the Smg protein. A) Sequences of interest are shown over top of a diagram of the full
length Smg protein. The conserved motifs are labeled a-f, while sequences containing one or
more phospho-serines are marked with +. B) Multiple sequence alignments of the six conserved
motifs a-f identified in Smg. The alignments include sequences from three Drosophilids, and the
mosquito Anopheles gambiae. The residues corresponding to the Drosophila melanogaster Smg
protein sequences are numbered. Phospho-serines within a conserved motif in the Drosophila
melanogaster Smg protein sequence are highlighted in grey. C) Two C-terminal phospho-serines
in the Smg protein are highlighted in grey.
67
68
that the mRNA decay mediated by NTsmg functions through a deadenylation-independent
mechanism, such as decapping or endonucleolytic cleavage. To explore this possibility, I would
take a tandem affinity purification and mass spectrometry approach to identify proteins which
interact with NTsmg. This approach could recover proteins involved with decapping or
endonucleolytic cleavage, which would support the hypothesis in which the Smg N-terminus
mediates transcript decay through decapping or endonucleolytic mechanisms. If binding of such
proteins was not seen in a mutant version of NTsmg that was defective for mRNA decay, this
would suggest the interaction is important for mRNA decay.
4.2.3 The mechanism of CTsmg function in mRNA decay
My assays of Hsp83, arrest, and BicC mRNA decay demonstrate that the Smg C-
terminus plays a role in transcript degradation, but does it function through deadenylation? While
insight into this question may be gained from the results of the assays investigating NTsmg in
mRNA decay, I would also take a similar approach to investigate CTsmg in mRNA decay and
deadenylation.
For example, if the results from the NTsmg assay outlined above show that the Smg N-
terminus does not mediate mRNA decay via deadenylation, then it would suggest that the CTsmg
induces deadenylation through recruitment of the CCR4/POP2/NOT deadenylase. I would
confirm this by testing a series of mutants in the Smg C-terminus for their ability to induce
deadenylation, and to interact with the CCR4/POP2/NOT complex. Mutants would be
constructed in the context of full length Smg, where I would generate a series of progressive C-
terminal truncations, as well as specifically mutate the two C-terminal conserved motifs.
In contrast, if the assays proposed to investigate NTsmg in mRNA decay and
deadenylation shows that NTsmg does bind the CCR4/POP2/NOT deadenylase, then it is
possible that the Smg C-terminus mediates mRNA decay via another mechanism. In this
scenario, I would generate a series of mutants in the Smg C-terminus in the context of the full
length protein in order to define the residues which function in transcript decay. This would
involve mutations of the two conserved motifs within this region, as well as progressive C-
terminal deletions. Once a region is mapped that is required for transcript decay, I would take a
tandem affinity purification and mass spectrometry approach to identify proteins which binds to
69
wild-type Smg, but not to a C-terminal mutant that is defective in mRNA degradation. Recovery
of decapping enzymes or endonucleases that meet this criteria would support a model whereby
the Smg C-terminus mediates mRNA decay through a deadenylase-independent manner and
indicate the mechanism through which the Smg C-terminal region functions in mRNA decay.
4.2.4 The role of the SSR2 in Smg function
The decay of Hsp83, arrest, and BicC mRNAs in smg47
mutant embryos rescued with
NTsmg and NTdSSR2 show that the SSR2 plays a transcript-specific role in regulating mRNA
decay activity of the Smg N-terminus during early embryogenesis. As suggested earlier, this may
occur through recruitment of a trans-acting factor by Hsp83 mRNA which binds to the SSR2 to
block transcript decay. Alternatively, it may be that arrest and BicC mRNAs recruit a trans-
acting factor which interacts with the SSR2 to block its inhibitory effect on transcript decay. To
ask which of these hypothetical models are true, I will identify proteins which interact with the
SSR2 by tandem affinity purification of NTsmg and NTdSSR2 proteins and mass spectrometry.
An alternative approach would be to capture proteins which bind to Hsp83, arrest, and BicC
mRNAs in smg47
mutant embryos rescued with NTsmg and NTdSSR2 by mRNA affinity
purification and mass spectrometry (Slobodin and Gerst, 2010, 2011). These approaches will
allow me to compare the binding partners recovered for NTsmg and NTdSSR2. Any proteins
whose interaction is lost when the SSR2 is deleted would be candidates as SSR2 binding
partners. Additionally, the mRNA affinity purification approach would identify SSR2 binding
partners which are transcript-dependent. For example, if an SSR2 binding partner is recovered
only with Hsp83 mRNA, it would support the model whereby a trans-acting factor is recruited
by Hsp83 to interact with the SSR2 and inhibit mRNA decay mediated by the Smg N-terminus.
In contrast, a protein which only binds SSR2 in the presence of arrest and BicC mRNAs, but not
Hsp83, argues in favour of the model where the trans-acting factor is recruited by arrest and
BicC mRNAs and interacts with SSR2 to allow mRNA decay mediated by the Smg N-terminus.
Further insight into the mechanism of this regulation would be gained from the identity and
known biological functions of the SSR2 binding partners identified.
4.2.5 Mechanisms of Smg-mediated translation repression
I have shown that both the Smg N- and C-terminal sequences are required to induce
transcript decay, but what are their roles in translation repression? To explore this question, I
70
would first ask whether the NTsmg protein is able to repress translation of nos mRNA, a well-
characterized target of Smg-mediated translation repression. Repression of nos mRNA by Smg
in the bulk cytoplasm of the embryo results in the expression of a gradient of Nos protein
emanating from the posterior. In smg mutant embryos, where nos mRNA is not translationally
repressed, Nos protein is ectopically and ubiquitously expressed. As such, I would take an
immunohistochemical approach to detect Nos proteins in smg47
mutant embryos rescued with
NTsmg. If Nos protein expression in smg47
mutant embryos rescued with NTsmg shows a
posterior gradient similar to wild-type embryos, this would indicate that, at least for nos mRNA,
that the translation repression activity of Smg lies in the N-terminus. If on the other hand, NTsmg
is defective for nos translational repression, this would suggest that C-terminal sequences are
involved in this process. I would then test the effect of progressive C-terminal truncations of this
region, as well as mutation of the two conserved motifs in this region on Smg-mediated
translational repression.
Subsequently, I would expand my analysis on Smg-mediated translation repression
beyond nos to other Smg target transcripts by employing polysome density gradients. This
technique separates translationally repressed, mRNP-associated mRNAs from poly-ribosome
(polysome) occupied, actively translated mRNAs. Indeed, work in our lab has identified, using
this method, a list of Smg target transcripts based on their shift from mRNP-association in wild-
type embryos to polysome-association in smg mutant embryos (J Dumelie and CA Smibert,
unpublished data). For my analysis, I would assay the mRNP and polysome association of the
top identified Smg target transcripts in wild-type embryos versus smg47
mutant embryos rescued
with NTsmg. A shift of these transcripts from mRNP-association in wild-type embryos to
polysome-association in the smg47
mutant embryos would suggest a loss in translation
repression. Thus, maintenance of these transcripts in mRNPs for NTsmg would argue that the
sequences in the Smg N-terminus mediate translation repression. This assay, together with the
immunohistochemical assay for nos expression, can show which Smg-terminal regions are
involved in translation repression.
If the assays proposed above show that sequences in the Smg N- and/or C-termini are
involved in translation repression, what are the binding partners through which they function? In
addition to Cup, Smg also mediates translation repression of nos mRNA through miRNA-
71
independent recruitment of Ago1 (Pinder and Smibert, 2013). Interestingly, I have preliminary
data showing that Ago1 co-immunoprecipitates with both full length Smg and the C-terminally
truncated smg1 protein. This maps the Smg/Ago1 interaction site to the protein sequences
encoded by the smg1 allele (amino acid 1-478), which contains SSR1, SSR2, and the first of the
four conserved motifs in the Smg N-terminus. To ask if the SSRs and conserved motif mediate
this interaction, I would test whether Ago1 co-immunoprecipitates with NTsmg, and whether the
interaction is disrupted when the SSRs or the conserved motif are deleted from NTsmg.
Disruption of the NTsmg/Ago1 interaction when only one of the SSRs or conserved motif is
deleted would map the deleted region as the Ago1 interaction site. However, if these deletions do
not disrupt the NTsmg/Ago1 interaction, it may be that the SSRs and conserved motif work in
concert to recruit Ago1. To follow this up, I would assay whether Ago1 binding is abolished
when the SSRs and conserved motif are deleted in combination. Alternatively, the protein
sequences between the SSRs and conserved motifs may function to recruit Ago1. Additional
mutants in which these protein sequences are deleted can be assayed to test whether they are
involved in Ago1 recruitment.
Finally, a tandem affinity purification and mass spectrometry approach with the FLsmg
and NTsmg protein could uncover novel Smg binding partners which are involved in translation
repression. This could identify mechanisms of Smg-mediated translation repression which are
additional to those involving Cup and Ago1. These novel mechanisms of Smg-mediated
translation repression can be tested using the assays proposed in this section to observe whether
Smg-mediated translation is affected in embryos mutant for these novel binding partners.
4.2.6 Mechanism of Smg protein degradation
The Smg C-terminus is necessary for timely degradation of Smg protein. In addition, the
protein encoded by the smg1 allele is further stabilized compared to NTsmg, suggesting that the
residues between amino acid 479 and 766 also play a role in Smg protein degradation. To map
the C-terminal sequences involved, I would test the effect of progressive C-terminal truncations
of this region, as well as mutation of the two conserved motifs in this region on the stability of
other intact Smg protein. Similarly, I would make progressive C-terminal truncations of the
NTsmg, as well as testing the effects of mutations in motifs b, c, and d (Figure 16) on the stability
of NTsmg to map other regions required for Smg protein degradation. Given the role of the Dop
72
kinase in Smg protein degradation, I would ask if regions of Smg which are required for its
degradation are also required for the Dop/Smg interaction, whether they are sites of Dop-
mediated phosphorylation, and if this phosphorylation is required for degradation of the protein.
4.2.7 Assessing additional smg mutant proteins
One aspect of Smg protein function involves its localization into foci throughout the bulk
cytoplasm of the embryo (Smibert et al., 1999; Zaessinger et al., 2006). However, the mechanism
by which these foci form, and their function in the embryo remain unclear. To ask which regions
of the Smg protein are involved in foci formation, I would first take an immunohistochemical
approach to visualize the ability of the NTsmg protein to localize into foci. If the NTsmg protein
does not localize into foci, it would indicate that the Smg C-terminus is required for foci
formation. Alternatively, NTsmg could form foci, indicating that it contains the required
sequences. Whichever region is implicated, I would map the elements involved using approaches
similar to those described above.
Furthermore, these immunohistochemical assays, which tests the role of the SSRs and
conserved motifs in Smg foci formation, will be performed in parallel with the assays proposed
previously, which investigates the roles of the SSRs and conserved motifs in mRNA decay and
translation repression. Together, the results of these assays may show a correlation of Smg-
mediated regulatory functions with Smg foci formation. For example, deletion of an SSR or
conserved motif which disrupts Smg foci formation and also impairs mRNA decay would imply
that Smg foci are sites of mRNA decay. In contrast, deletion of an SSR or conserved motif which
disrupts Smg foci formation and impairs translation repression would suggest Smg foci as sites
of translation repression.
To complement the above experiments, I would also ask if the various Smg binding
partners identified above also localize to Smg foci. The identity of these binding partners, the
regions of Smg required for their interaction, and the role of these regions in Smg function
should provide an indication of the function of these foci.
73
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