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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