Download Developmental Signaling by Noggin and Wnt in the Frog Xenopus

Developmental Signaling by Noggin and Wnt
in the Frog Xenopus
By
John Joseph Young
A dissertation submitted in partial satisfaction of the
requirements for the degree of
Doctor of Philosophy
in
Molecular and Cell Biology
in the
Graduate Division
of the
University of California, Berkeley
Committee in charge:
Professor Richard M. Harland, Chair
Professor Sharon Amacher
Professor Michael S. Levine
Professor Michael Freeling
Spring 2013
Developmental Signaling by Noggin and Wnt in the Frog Xenopus
© 2013
By John Joseph Young
Abstract
Developmental Signaling by Noggin and Wnt
in the Frog Xenopus
By
John Joseph Young
University of California, Berkeley
Professor Richard M. Harland, Chair
Xenopus has provided a powerful system to study cellular, developmental, and neurobiology. The availability of their embryos and the advent of modern molecular techniques
allowed investigators to revisit the observations of classical embryologists and begin to
determine the molecular mechanisms underlying germ layer formation and axis induction. My
thesis work took advantage of the frog Xenopus to first address the developmental role of
Noggin, a Bone morphogenic protein (Bmp) antagonist, and then to determine the mechanism of
Wnt-induced anterior-posterior patterning of the neural plate.
The frog Xenopus, an important research organism in cell and developmental biology, currently
lacks tools for targeted mutagenesis. In the first part of this work, I address this problem by
genome editing with zinc finger nucleases (ZFNs). ZFNs directed against an eGFP transgene in
X. tropicalis induced mutations that are consistent with results of non homologous end joining at
the target site, resulting in mosaic loss of fluorescence phenotype at high frequencies. ZFNs
directed against the noggin gene produced tadpoles and adult animals carrying up to 47%
disrupted alleles. Founder animals yielded progeny that carry insertions and deletions in the
noggin gene with no indication of off-target effects. Furthermore, functional tests demonstrated
an allelic series of activity among three germline mutant alleles. Breeding an identified null
allele to homozygosity resulted in tadpoles with deformaties in the cranial skeleton. Anatomical
analysis revealed severe reductions in Meckel’s cartilage with joint fusions. Gene expression
analysis via in situ hybridization for chondrogenesis regulating factors in noggin mutants
revealed a reduction in sox9 and col2a expression domains. Analysis of Bmp targets showed an
expansion of hand2, edn1, and msx2 in the pharygeal arches (PAs) of mutants. This suggested a
mechanism whereby incresed Bmp signaling inhibits chondrogenesis and ventralizes the PAs
resulting in the jaw deformities observed in mutants.
Neural development in amphibians occurs as a two-step process. First, ectodermal precursors
adopt a neural fate in the absence of Bmp signaling. A second signal is then required to pattern
the anterior posterior neuraxis. Signaling through Fibroblast growth factor (Fgf), retinoic acid
(RA), and Wnt have each been demonstrated to be both necessary and sufficient for inducing
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posterior fates in undifferentiated neural tissue. Wnt signaling in particular has been closely
studied. However, the mechanism by which this pathway induces posterior fates remains
unclear. To address this question, I used RNA-Seq to identify direct transcriptional targets in
neural tissue by activating Wnt signaling in Xenopus neural explants pretreated with the
translation inhibitor cycloheximide. Wnt-activated neural tissue resulted in over 200 genes with
expression increased greater than 2-fold when compared to anterior neural tissue. in situ
hybridization analysis of highly expressed transcription factors and RNA-binding proteins
showed posterior expression. Of particular interest, the transcription factor sal-like 1 (Sall1) and
sal-like 4 (Sall4) showed specific posterior neural expression suggesting a role in Wnt-induced
neural patterning.
The RNA-Seq screen found sall1 and sall4 expression to be induced by canonical Wnt signaling
!'#%&!","+ !
&'&#%&!'!'.%&'!'%"!"sall4 were
enriched in β-catenin chromatin imunoprecipitations. Knockdown of Sall4 resulted in the loss of
spinal cord marker expression and an increase in the expression of pou25, pou60 and pou91
(pouV genes), the three Xenopus homologs of the stem cell factor pou5f1/Oct4. Overexpression
of the pouV genes resulted in the loss of spinal cord identity, and knockdown of pouV function
restored spinal cord marker expression in Sall4 morphants. Finally, knockdown of Sall4 blocked
the posteriorizing effects of Fgf and retinoic acid signaling in the neurectoderm. These results
suggest that Sall4, activated by Wnt signaling, represses the pouV genes to provide a permissive
environment that allows for additional Wnt/Fgf/RA signals to posteriorize the neural plate.
2
To Nick Duesbery, Ph.D and Jeff McKelvey, Ph.D.
I’m grateful to have been your student, proud to be your colleague, and most of all,
honored to be your friend.
i
Table of Contents
Table of Contents......................................................................................................................ii
Table of Figures........................................................................................................................iv
Acknowledgements..................................................................................................................vi
Chapter 1: General Introduction.............................................................................................1
Classical Embryology.....................................................................................................1
The Molecular Era and Xenopus.....................................................................................2
Open questions................................................................................................................8
Goals of this thesis..........................................................................................................9
Chapter 2: Materials and Methods.........................................................................................11
Embryo and explant culture............................................................................................11
RNA and morpholino microinjections............................................................................11
Western Blotting.............................................................................................................12
Celery Extract Preparation..............................................................................................12
Mutation Detection by Celery Extract............................................................................12
Cartilage staining............................................................................................................13
Cycloheximide and dexamethasone treatments..............................................................13
Whole-mount in situ hybridization.................................................................................13
Embedding and Sectioning.............................................................................................14
RT-PCR and qPCR..........................................................................................................14
RNA-seq..........................................................................................................................14
ii
Chromatin immunoprecipitation...................................................................................14
Chapter 3: Analysis of the developmental role of noggin in Xenopus tropicalis
development via Zinc-Finger Nuclease mutagenesis...........................................................16
Introduction...................................................................................................................16
Results...........................................................................................................................17
Discussion.....................................................................................................................21
Chapter 4: Expression screen for direct targets of Wnt signaling in
neural tissue.............................................................................................................................46
Introduction...................................................................................................................46
Results and Discussion..................................................................................................47
Chapter 5: Spalt-like 4 mediates Wnt-induced neural patterning via repression of
pouV/Oct4 family members..................................................................................................................68
Introduction................................................................................................................................68
Results........................................................................................................................................69
Discussion...................................................................................................................................73
References...............................................................................................................................................97
Appendices.............................................................................................................................................118
I: RNA-Seq results from Chapter 3: Genes with >2-fold
expression (direct Wnt activation vs. anterior neural)..................................................118
II: List of PCR primers used in this work.....................................................................128
III: List of DNA plasmids used in this work................................................................131
iii
List of Figures
Figure 3.1: Disruption of the eGFP transgene in Xenopus tropicalis using ZFNs.....................24
Figure 3.2: Tolerance and activity of ZFNs targeting noggin in Xenopus tropicalis.................26
Figure 3.3: ZFN-driven editing of the noggin locus in Xenopus tropicalis...............................28
Figure 3.4: ZFNs induce heritable loss-of-function noggin alleles mutations...........................30
Figure 3.5: Knockdown of Chordin and Follistatin results in a loss of dorsal
structures in a subset of embryos produced by heterozygous noggin mutant adults..................32
Figure 3.6: Stage series of representative embryos produced by
heterozygous noggin mutant adults.............................................................................................34
Figure 3.7: Homozygous noggin mutant Xenopus tropicalis have severe
lower jaw deformities..................................................................................................................36
Figure 3.8: Expression of chondrogenic factors in wild-type and noggin
mutant tadpoles............................................................................................................................38
Figure 3.9: Expression of Bmp pathway targets in wild-type and noggin
mutant tadpoles............................................................................................................................40
Figure 4.1: Model of Wnt-induced patterning of the neural anterior-posterior
axis...............................................................................................................................................52
Figure 4.2: Temporal expression of anterior posterior neural markers in
Xenopus tropicalis........................................................................................................................54
Figure 4.3: TVGR activates canonical Wnt-signaling................................................................56
Figure 4.4: TVGR efficiently posteriorizes neuralized ectodermal explants..............................58
Figure 4.5: Cycloheximide treatment prior to TVGR induction enriches for
direct Wnt targets in neuralized ectodermal explants...................................................................60
Figure 4.6: Expression patterns of transcription factors identified in the screen for
direct Wnt targets..........................................................................................................................62
Figure 4.7: Expression patterns of RNA-binding factors identified in the screen for
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direct Wnt targets..........................................................................................................................64
Figure 4.8: Expression patterns of identified Wnt targets prickle1 and lmo7.............................66
Figure 5.1: sall4 is a direct transcriptional target of canonical Wnt-signaling............................77
Figure 5.2: Injected embryos express functional FLAG-tagged β-catenin..................................79
Figure 5.3: sall4 is expressed in the neurectoderm......................................................................81
Figure 5.4: sall1 is directly activated by canonical Wnt signaling and expressed
during early embryogenesis...........................................................................................................83
Figure 5.5: Loss of Sall4 results in a loss of posterior neural differentiation...............................85
Figure 5.6: cdx2 is directly activated by canonical Wnt signaling and not
affected by Sall4 knockdown.........................................................................................................87
Figure 5.7: Knockdown of Sall4 causes an increase in expression of the
pouV/Oct4 homologs.....................................................................................................................89
Figure 5.8: A second non-overlaping Sall4 morpholino results in similar phenotypes................91
Figure 5.9: Loss of spinal cord gene expression in Sall4 morphants requires an
increase in pouV/Oct4 expression..................................................................................................93
Figure 5.9: FGF and retinoic acid signaling fail to posteriorize Sall4 morphants........................95
v
Acknowledgements
First off, I want to thank Nikki, I could not have done any of this without you. I’m so excited for
our lives and all the adventures we’ll have together. Mom, you always said I could do this,
especially when I didn’t think I could. Thank you, I’ll always remember Melinda Mae. Dad,
thanks for all the encouragement and that great weekend of baseball, darts, and beer. Em, I’m so
lucky to have you as my sister. I have to thank my grandmother June, the first frog biologist I
ever knew. I love you all dearly.
My advisor Richard Harland, I knew I wanted to join your lab even before coming to Berkeley.
You’ve been an excellent mentor; kept me from flying too high, but propped me up when I
needed it. Thank you for giving me the freedom to explore my questions. I’m so proud to have
been in your lab.
To the members of my committee: Sharon Amacher, Mike Levine, and Mike Freeling, thank you
for your guidance, insight, and support.
I’m entirely grateful to the past members of the Harland lab crew: Andrea and James, I learned
so much about how to be a scientist from you, also how to play MarioKart, thank you for the
discussions and distractions. Mike, even if it can’t be Slayer, let’s take the Prius to another metal
show. Sara, thank you for scaring me first and then becoming a great friend and colleague. Isa
and Jen, I’m always ready for a beer lunch. My sincerest thanks to Jess, words truly fail here, I
can’t imagine grad school without you.
To my current labmates: Debbie, the warm, comfortable environment of the lab is in no small
part due to you. Thank you for making this an amazing place to work. Darwin, I’ll always think
of you when I need someplace to store a box of tubes and Dave, we’ll get that pig next time.
Caitlin and Sofia, you’re up next, I’m looking forward to hearing about all of your discoveries.
Peter, a fellow PBR lover, I’m so glad I got to know you. Hyeyoung and Lisa, thank you for
setting high standards for science in our lab. Stefanie, Cameron, and Rachel you’re part of the
best lab in MCB, I know you’ll do amazing science.
I have been fortunate to work with some wonderful undergraduate students here at Berkeley.
Gloria, Daniel, and Sofia, I have learned so much from you. Your futures are exciting and I wish
you the best in your careers.
I want to thank my lifelong friends Jason and Jeff. You are my real life heroes, thank you for all
of our times together, good and bad. You’ll always be my brothers.
Finally, I couldn’t have remained sane without Copsound: Brock, Justin, Dave, Alberto, Josh,
Blair and Mike. Thank you for giving me the opportunity to rock with you guys. Your creativity
and energy inspired me both in lab and at the space.
vi
CHAPTER 1
General Introduction
Classical Embryology
How an animal develops from a fertilized egg to a recognizable multicellular organism is a
fundamental question in developmental biology. This question has intrigued countless biologists
for thousands of years prompting many theories, both ingenious and ridiculous. Aristotle first
used the term epigenesis to describe chick development as a series of steps whereby structures
progressively induce the formation of other structures, ultimately giving rise to a complex
organism with multiple cell types and tissues. While this seems like basic knowledge in modern
times, his interpretations were swept aside in favor of the creationism-friendly preformationist
idea that tiny humunculi are present in the germ cells. Battles raged between the ovists and
spermists as to which gamete truly contained the humunculus until the mid 19th century when
Schleiden and Schwann’s theory that the fertilized egg formed a cell that through division gave
rise to all the cells in the body began to gain acceptance. Today, the combined theories of
Aristotle and Schleiden and Schwann form the core of modern developmental biology. Every
embryo contains the intrinsic information to build itself from within via cell division and
epigenesis, however the question remains, how?
Embryonic induction, where a specific tissue or cell(s) induces the fate of a different tissue or
group of cells, provides a general mechanism for epigenesis as described by Aristotle. The
experiments of Hans Spemann and Warren Lewis demonstrated the first embryonic induction
when they found that grafting an optic cup from a frog embyo to an ectopic location was
sufficient to induce lens formation in the overlying epidermis (Spemann, 1901) (Lewis, 1904).
Spemann then turned his attention from specific organ development to axis formation by grafting
pieces of gastrulae embryos to ectopic locations in host embryos. Spemann made his greatest
discovery when Hilde Mangold, a student in his lab, grafted the dorsal lip from a gastrula of the
lightly colored newt Triturus cristatus to the ventral region in a gastrula of the darkly pigmented
newt Triturus taeniatus. This graft generated an ectopic axis but the important discovery here
was that the secondary axis was comprised of a lightly pigmented notochord and floorplate and
darkly pigmented somites and neural tube. The darkly pigmented cells must have come from the
host and they made the conclusion that the grafted tissue induced the surrounding host tissue to
adopt a dorsal fate rather than a ventral one (Spemann and Mangold, 1924). This result
prompted the idea of regional specification where cells in the dorsal lip of the gastrula, known as
the organizer, specify the fates of other cells in the region (Spemann, 1938).
The discovery of mesendodermal tissue that induce surrounding cells to adopt alternate fates was
not limited to amphibians. Waddington’s discovery that transplantation of Hensen’s node from
chicks would induce ectopic neural tissue in host ectoderm showed this structure to be the avian
equivalent of Spemann’s organizer (Waddington, 1932; Waddington, 1933). The shield in teleosts
(Oppenheimer, 1936), and finally the node in mammals (Beddington, 1994) were also found to
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be capable of patterning surrounding tissue during gastrulation. These discoveries provided great
insight into how an embryo generates its pattern but they opened several more questions about
how these structures are themselves induced and how they exert their effects on neighboring
tissue.
Pieter Nieuwkoop’s discovery that ectoderm will change fate and give rise to mesodermal
derivatives when placed next to endoderm provided insight on how the embryo generates the
different germ layers (Nieuwkoop 1969). Furthermore, he noticed that the induced mesodermal
derivatives were different depending on the dorsal/ventral nature of the endoderm with which the
ectoderm was combined. Ventral endoderm induced blood and smooth muscle fates whereas
dorsal endoderm was capable of inducing nearly all tissues in the embryo (Nieuwkoop 1969)
(Boterenbrood 1973) (Dale and Slack, 1987). This observation led to the discovery of the socalled Nieuwkoop center, a portion of the dorsal endoderm that induces formation of the
organizer. Transplantation of the dorsal-most endodermal cells at the 64-cell stage from embryos
of the frog Xenopus laevis to the ventral side of irradiated embryos rescued axial structures in
what would otherwise develop into a ball of ventral tissue known as a “belly piece” (Gimlich and
Gerhart, 1984). Despite these incredible advances, the field of developmental biology would
have to wait for the molecular era before the precise mechanisms of cell specification and
embryonic inductions could be determined.
The Molecular Era and Xenopus
The field of experimental embryology was revolutionized by the adoption of the South African
Clawed Frog Xenopus laevis as a model system. In the early 20th century, the discovery that
injection of a pregnant female’s urine into the dorsal lymph sac of Xenopus induced ovulation
provided a robust and reliable test for pregnancy (Crew, 1939). This resulted in the export and
housing of these frogs in medical institutions worldwide (Gurdon and Hopwood, 2000). The
byproduct of this discovery was that amphibian eggs and therefore embryos could be made
available year round and in great numbers. Researchers no longer had to wait for the breeding
season to obtain their often limited experimental material. The availability of embryos and the
advent of modern molecular techniques allowed investigators to revisit the observations of
classical embryologists and begin to determine the molecular mechanisms underlying germ layer
formation and axis induction.
In amphibians, the endoderm forms mostly in the vegetal region, the mesoderm forms in the
marginal zone and the ectoderm comes from the animal region. Nieuwkoop’s experiments
suggested that factors from the endoderm induce the overlying ectoderm to adopt a
mesendodermal fate. Basic Fibroblast growth factor (bFgf) was identified as having weak
mesoderm inducing activity that was enhanced when combined with Transforming Growth
Factor-β (TGFβ) (Kimelman and Kirschner, 1987) (Slack et al., 1987). It was found that a pellet
of Xenopus tissue culture cells had the same mesoderm inducing properties as dissected
endoderm on ectodermal explants (animal caps) (Smith, 1987). This led to the discovery that the
TGFβ family member ActivinA was a potent mesoderm inducer. Treatment of animal caps with
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ActivinA alone was sufficient to induce mesodermal differentiation (Smith et al., 1990).
Furthermore, increasing doses of ActivinA resulted in more dorso-anterior characteristics of the
induced mesoderm (Green and Smith, 1990). While Fgf and ActivinA were demonstrated to
induce mesoderm in animal caps, it’s more likely that the Xenopus nodal-related (Xnr) factors
signaling through Smad2 are the molecules that induce mesoderm in the frog embryo. The Xnrs
are expressed in the marginal zone and overexpression of a Nodal-specific version of the
inhibitor Cerberus, Cerberus-short, blocks mesendoderm formation in embryos (Agius et al.,
2000; Joseph and Melton, 1997; Kessler and Melton, 1995; Piccolo et al., 1999). Nodal
induction of mesoderm is not limited to amphibians. The most direct evidence that Nodal
signaling induces mesendoderm in vertebrates was provided by forward genetic screens in the
zebrafish Danio rerio. Mutations in cyclops, squint and one-eyed pinhead, genes which encode
the nodal homologs and their receptor, result in a failure of mesendoderm and organizer
formation and can be rescued by injection of these factors(Feldman et al., 1998; Gritsman et al.,
1999). In both the chick and the mouse, Nodals induce mesoderm and their antagonist Lefty
restricts signaling to prevent multiple primitive streak formation (Bertocchini and Stern, 2002;
Conlon et al., 1994; Perea-Gomez et al., 2002; Skromne and Stern, 2001; Skromne and Stern,
2002; Zhou et al., 1993).
Nodal secretion alone from the endoderm, however, does not explain the induction of the
organizer. While high doses of ActivinA were able to induce dorsal mesoderm, this was not
likely the mechanism employed by the embryo. The organizer is comprised of cells that will
give rise to the notochord and head mesoderm in amphibians. Nodal signaling, while required
for mesoderm induction, is necessary for organizer formation but not sufficient. Therefore, a
dorsal modifying signal was proposed to be present in the embryo that would give the mesoderm
induced in the presumptive dorsal region the inductive activity of the organizer. Strong evidence
that this signal is mediated by Wnt signaling was provided when expression of Wnt8 on the
future ventral side of the Xenopus embryo was able to induce a second axis (Smith and Harland,
1991; Sokol et al., 1991). Lineage tracing the injected cells revealed cells that inherited the wnt
RNA comprised the notochord, pharyngeal tissue, and portions of the somites, reminiscent of
dorsal lip grafts done with different newt species by Spemann and Mangold (Spemann and
Mangold, 1924). Activation of the Wnt pathway results in the nuclear accumulation of β-catenin
which complexes with T-cell factor/Lymphocte enhancer factor (TCF/LEF) to activate
transcription of target genes, reviewed in (Logan and Nusse, 2004). Depletion of the maternal
pool of β-catenin transcripts resulted in embryos that lacked dorsal derivatives of the mesoderm
and ectoderm (Heasman et al., 1994; Wylie et al., 1996). Several wnt transcripts are maternally
deposited including wnt8b (Cui et al., 1995), wnt5a (Moon et al., 1993), and wnt11 (Ku and
Melton, 1993). Through the movements driven by cortical rotation, these molecules, initially
located at the vegetal pole of the egg, get displaced to the presumptive dorsal side during the first
cell cycle of the zygote. Similar to β-catenin-depleted embryos, depletion of wnt11 from oocytes
prior to fertilization resulted in embryos lacking dorsal structures (Heasman et al., 2000; Tao et
al., 2005). These results showed that Wnt signaling is the dorsal modifying signal in the
embryo . Recently, it has been proposed that maternally-provided wnt8 is necessary for
organizer induction in zebrafish (Lu et al., 2011).
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The combined action of Wnt and Nodal signaling on the presumptive dorsal side is responsible
for the Nieuwkoop center’s inductive activity in the blastula. These signaling pathways lead to
activation of specific transcription factors and signaling molecules whose expression domain will
come to define the organizer. At the blastula stage of Xenopus embryos, there is a gradient of
activated Smad2 that begins on the dorsal side and spreads ventrally (Lee et al., 2001). This
initial burst of Smad2 signaling on the dorsal side is the result of the early Wnt signal
cooperating with maternally-provided VegT. xnr3 is directly activated by Wnt/β-catenin
(McKendry et al., 1997) and stabilized maternal β-catenin on the dorsal side serves to prime the
promoters of xnr3,5, and 6 for activation via recruitment of Histone Methyltranferases (Blythe et
al., 2010). The combined activity of Wnts and Nodals on the future dorsal side serve to activate
organizer specific transcription factors in the dorsal mesoderm. The 5’ region of the paired-like
homeobox gene siamois (sia) contains a proximal element with TCF/LEF binding sites and the
closely related gene twin also has a distal element that is regulated by Smad2 (Houston et al.,
2002; Laurent et al., 1997; Nishita et al., 2000). Signaling by Wnt and Nodal synergistically
activates twin expression in the organizer only on the dorsal side(Nishita et al., 2000). Similarly,
the homeobox gene goosecoid (gsc) has a proximal region with TCF/LEF binding sites that is
also bound by Twin, which serves to mediate gsc expression in response to the Wnt signal
(Laurent et al., 1997). A distal enhancer with Smad2 binding elements also mediates gsc
expression but requires cooperative binding with members of the Mixer homeodomain family of
transcription factors (Germain et al., 2000). Bmp, however, restricts gsc to the dorsal mesoderm;
injection of noggin RNA into ventral mesoderm results in a broader gsc expression domain
(Eimon and Harland, 1999). Expression of gsc or sia on the ventral side of embryos results in
the induction of an ectopic organizer and the generation of twinned embryos (Cho et al., 1991;
Lemaire et al., 1995).
While the identification of organizer-specific transcription factors led to an understanding of how
the organizer is specified, it was the discovery of secreted molecules from this specialized group
of cells that provided the mechanism for axis induction observed by Spemann and Mangold in
their grafting experiments. Through expression screens using cDNA libraries constructed either
from dissected organizers or whole embryos treated with a dorsalizing agent such as LiCl,
several secreted factors were found to have organizer-like activity. The first such factor to be
discovered was noggin. It is normally expressed in the organizer and when expressed in UVventralized embryos, was capable of fully rescuing anterodorsal structures (Smith and Harland,
1992; Smith et al., 1993). Furthermore, treatment of animal caps with Noggin protein was
sufficient to induce a neural fate (Lamb et al., 1993). Within a short time, other molecules with
the same neural inducing activity were cloned including chordin (Sasai et al., 1994) and
follistatin (Hemmati-Brivanlou et al., 1994). These factors bind to and inhibit Bmps
(Zimmerman et al., 1996). Upon this discovery, Piccolo (Piccolo et al., 1996) suggested a
mechanism for organizer function whereby the organizer cells which give rise to the head
mesoderm and the notochord secrete Bmp antagonists to inhibit Bmp signaling in nearby tissue.
From these experiments, and earlier ones where the Activin receptor was inhibited (HemmatiBrivanlou and Melton, 1994), the default model for neural induction was proposed, stating that
4
ectoderm will adopt a neural (dorsal) fate in the absence of any inducing signal (HemmatiBrivanlou and Melton, 1997). Consistent with this model, genetic knockouts of noggin and
chordin result in mice lacking dorsoanterior-most structures however, they still retain neural
tissue(Bachiller et al., 2000). This result suggested that there are other Bmp antagonists that
function redundantly to induce neural tissue. Knockdown of three Bmp antagonists in Xenopus
tropicalis resulted in a complete loss of dorsal structures and yielded fully ventralized embryos
(Khokha et al., 2005). The presence of several Bmp antagonists that act redundantly in the early
embryo has made assessing their individual role challenging.
There is strong evidence for the default model. In vitro culture of excised animal caps results in
the adoption of an epidermal fate, however dissociation and reaggregation of caps result in a
neural fate (Godsave and Slack, 1989; Grunz and Tacke, 1989; Sato and Sargent, 1989). Neural
induction following dissociation can be blocked by adding Bmp to the culture medium (Wilson
and Hemmati-Brivanlou, 1995). These studies provided a model where Bmp signaling through
the Smad1/5/8 transcription factors serve to induce ventral fates while cells with low Bmp adopt
dorsal fates. The key experiment that supported this model was performed when a truncated
form of the activin receptor (which acted as a dominant negative for Bmp signaling) was
expressed in animal caps and neural tissue was induced (Hemmati-Brivanlou and Melton, 1992).
Taken together, these experiments support the interpretation that neural tissue is “induced” by the
removal of Bmp and is thereby the default differentiation pathway of the ectodermal precursors.
The default model can be applied to the other germ layers. Bmp signaling induces ventral fates:
blood, body wall muscle and hindgut in the mesoderm and endoderm, respectively. Dorsal fates
such as the somites and pharyngeal endoderm are induced when Bmp is blocked (Harland’s
chapter in (Stern, 2004). If one were to carry this model to a logical conclusion, then depletion
of the mesoderm and endoderm inducer along with Bmp inhibition would result in the entire
embryo adopting a neural fate . This intriguing hypothesis awaits experimental testing.
Despite broad acceptance of the default model for neural induction, some challenges have arisen
to the interpretation that Bmp inhibition alone is sufficient to induce ectoderm to adopt neural
fates. Injection of a dominant negative Fgf receptor blocked the neural inducing activities of
Noggin and Chordin which prompted the hypothesis that Fgf signaling is required for neural
induction (Launay et al., 1996; Sasai et al., 1996). The finding that simply cutting the animal cap
leads to activation of MAPK, the transducer of Fgf, called into question the interpretation of
experiments where animal caps were treated with BMP antagonists (LaBonne and Whitman,
1997). Furthermore, inhibition of Smad1 also required Fgf signaling to induce neural fates in
ventral epidermis (Delaune et al., 2005). This however, is due to the activity of Smad2, as
inhibition of Smad1 and Smad2 is sufficient to induce neural fates ventrally (Chang and Harland,
2007). An alternative explanation could be that Fgf-mediated activation of MAPK results in
phosphorylation of the linker region of SMAD1 causing its export from the nucleus and down
regulation of Bmp signaling (Fuentealba et al., 2007). Recently, it was proposed that the animal
cap is unsuitable for neural induction studies because of a pre-pattern imposed by other inductive
signals (Linker et al., 2009). This is unlikely to be the case, animal caps will form neural tissue
in the presence of a small molecule Fgf inhibitor when treated with a Bmp antagonist (Wills et
5
al., 2010). The source of the discrepancy is likely due to the use of sox2 as a marker for neural
tissue differentiation. While differentiated neural cells express sox2, a population of
uncommitted stem cells also express sox2 which revert to epidermis in the absence of Fgf (Wills
et al., 2010).
In addition to the Bmp antagonists, screens also identified secreted Wnt antagonists expressed in
a subset of organizer cells that give rise to the head mesoderm (Niehrs, 2004). The inhibitor
Dickkopf (Dkk), which blocks Wnt signaling by binding to the Wnt co-receptor Lrp5/6 and thus
preventing signal transduction (Semënov et al., 2001) is expressed in a subset of organizer cells
(Glinka et al., 1998). Overexpression of Dkk results in enlarged heads and reduced trunks
whereas loss-of-function experiments showed Dkk to be required for anterior neural
development (Glinka et al., 1998; Mukhopadhyay et al., 2001). Similarly, ectopic expression of
the Frizzled-related protein, Frzb induced anterior structures by binding to Wnt8 and preventing
it from binding to its receptors Lrp5/6 and Frizzled (Wang et al., 1997). Several additional
secreted Frizzled-related proteins were discovered in a screen for secreted proteins produced by
the organizer (Pera and De Robertis, 2000). As expected, these factors result in enlarged anterior
structures when overexpressed. The identification of the head inducer Cerberus as an inhibitor of
Wnts, Bmps, and Nodals (Piccolo et al., 1999) led to a model of axis induction by the organizer
where the Bmp antagonists induce a dorsal fate via repression of Bmp signaling and the anterior
is induced by inhibition of Wnt, Bmp, and Nodal signaling (reviewed in (Niehrs, 2004). The
inactivation of the repressor Tcf3 in zebrafish headless mutants further provided genetic evidence
that Wnt inhibition is required for head formation (Kim et al., 2000).
The default model, now supported by molecular evidence, is not completely novel. Johannes
Holtfreter and Pieter Nieuwkoop predicted that neural patterning was a two-step process
whereby the ectodermal precursors are first activated to adopt a neural fate (by default anterior in
nature), and then additional signals from the mesoderm posteriorize it to create the full anteriorposterior (A-P) pattern of the neural plate (HOLTFRETER, 1947; Nieuwkoop, 1952;
"62BD8<<=#A52?@
.'52Q;16;4A5.AA52<?4.;6G2?@20?2A2@*;A.;A.4<;6@A@@B442@[email protected]
*;A@64;.96;46@<;2@B05A?.;@3<?:6;43.0A<?5<D2C2?@2C2?.996;2@<32C612;02.9@<=<6;A21A<
43.;1?2A6;<60.061%@64;.96;[email protected];4;2B?.9A?.;@3<?:6;4.0A6C6AF
%12?6C213?<:)6A.:6;C6.%2A6;.9125F1225F1?<42;.@2%.9156@D2998;<D;.@.
A2?.A<42;D6A5:B9A6=9223320A@<;A5202;A?.9;2?C<B@@F@A2:D52;26A52??21B021<?6;0?2.@21
?2C62D216;!.12;!.12;'52?<92<3%6;$=.AA2?;6;4D.@?2C2.921D52;
@B6A./92:.?82?@D2?216@0<C2?21A<12A20A05.;[email protected]<;4A52.E6@'?2.A:2;A:2;A<3Xenopus
2:/?F<@D6A5%?2@B9A@6;:60?<02=5.9F.;1.;.;A2?6<?2E=.;@6<;<3@=6;.90<?1.;156;1/?.6;
:[email protected]<;4D6A5.?2=?2@@6<;<3.;A2?6<?;2B?.942;2@B?@A<;2A.9
<992:.;;2A.9
&6C22A.9
'?2.A:2;A<3;2B?.96G21.;6:.90.=@D6A5%?2@B9A@6;A522E=?2@@6<;
<3=<@A2?6<?;2B?.942;2@'52@2Q;16;4@12:<;@A?.A2A5.A%16?20A9FA?.;@3<?:@A52
;2B?20A<12?:$.=.9<=B9B.;16;A;2?
;720A6<;<3.1<:6;.;A;24.A6C23<?:<3A52%
receptor results in anteriorized Xenopus2:/?F<@D6A5?21B021A.69@D56920<;@A6ABA6C29F.0A6C2
3<?:@42;2?.A252.192@@A.1=<92@9B:/2?42A.9
'52@22E=2?6:2;A@=?<:=A21.:<129
D52?2/F%3<?:@.4?.162;A.9<;4A52;2B?.E6@D6A556452?0<;02;A?.A6<;@42;2?.A6;4:<?2
6
posterior fates. By using hox42;22E=?2@@6<;A<:.?8@=206Q0?5<:/<:2?2@6;A5256;1/?.6;<3
A52:<B@26AD.@16@0<C2?21A5.A6;0?2.@6;4%@64;.96;4?2@B9A@6;.;A2?6<??5<:/<:2?2@
.1<=A6;4=<@A2?6<?3.A2@<;9<;.;1%<@@.;A
&=206Q0.99F6;0?2.@21%@64;.96;4
resulted in ectopic hoxb1 ?5<:/<:2?2@=206Q02E=?2@@6<; and a loss of krox-20 in
?5<:/<:[email protected]
<D2C2?.05.992;42A<A56@:<1290<:2@3?<:A52
Q;16;4A5.AA52$=.AA2?;6;[email protected]<06.A21D6A5?2A6;<6112Q062;A>B.69@Raldh mutant
mice, and the raldh2G2/?.Q@5:BA.;Aneckless can all be rescued by a uniform concentration of
%242:.;;2A.9
.922A.9
!602A.9'52?2@<9BA6<;A<A56@.==.?2;A
=.?.1<ED.@3<B;16;G2/?.Q@5;<086;41<D;A52F=09.@@<30FA<05?<:2=2;GF:2@
D5605124?.12%?2@B9A216;/?<.12E=?2@@6<;<3%?2@=<;@6C242;2@.;1.3B99
=<@A2?6<?6G.A6<;<3A5256;1/?.6;2?;.;12G2A.9?2C6@21:<129;<D@B442@[email protected]
%6@=?<1B0216;A52=.?.E6.9:2@<12?:"6212??26A52?2A.9
.;16@@5.=216;A<.
gradient not by simple diffusion but by degradation anteriorly by the Cyp26 proteins.
@:2;A6<;21./<C2.;2:/?F<;60=.AA2?;6;4?<923<?43@64;.96;4D.@Q?@A612;A6Q216;
:2@<12?:6;1B0A6<;6:29:.;.;16?@05;2?
<D2C2?2C612;023<?43.0A6;4.@.
:<?=5<42;6;;2B?.9$=.AA2?;6;40.:2D6A5A5216@0<C2?FA5.A;2B?.96G21.;6:.90.=@2E=?2@@
increasingly more posterior neural markers with increasing doses of bFgf (Kengaku and
#8.:<A<
.:/.;1.?9.;1
'56@=<@A2?6<?6G6;4.0A6C6AF6@:216.A21A5?<B45A52
!$@64;.96;[email protected]@6;02A?.;@=9.;A21;2B?20A<12?:D6A5.1<:6;.;A;24.A6C23<?:<3%.@
6@B;./92A<42;2?.A2=<@A2?6<?;2B?.93.A2@+B2A.9
%6/[email protected]<;.99F
A52=<@A2?6<?6G6;4.0A6C6AF<3/43120?2.@[email protected]<3A5220A<12?:A?2.A:2;A<[email protected]
.;6:.90.=@?2@B9A@6;2E=?2@@6<;<3A52@=6;.90<?1:.?82?hoxb95<D2C2?A?2.A:2;A<[email protected]
12 caps only induces the more anterior gene enrailed2 (Lamb and Harland, 1995). This result is
consistent with a model where the posterior neurectoderm is in contact with the Fgf source early
in gastrulation and diffusion of the ligand reaches more anterior regions as the process of
4.@A?B9.A6<;=?<0212@&6;492:<920B92.;.9F@6@<343C6.RB<?2@02;020<??29.A6<;
@=20A?<@0<=F6;G2/?.Q@5?2C2.921A5.AA52964.;11633B@2@?.=619F3?<:@<B?020299@A5?<B45
2EA?.0299B9.?@=.02.;[email protected];B=A5?<B452;1<0FA<@6@6;?2@=<;16;40299@,B2A.9
This source/sink mechanism generates a gradient that in combination with the competence of
?2@=<;16;4;2B?20A<12?:=?<C612@.:205.;6@:3<?$=.AA2?;6;4<3A52;2B?.9=9.A2/F43
@64;.96;400<?16;49F.;43.@=96023<?:?2@B9A@6;.;2E=.;@6<;<3=<@A2?6<?;2B?.9:.?82?@
.;1.?2=?2@@6<;<3.;A2?6<?A6@@B2D52;<C2?2E=?2@@216;Xenopus5?6@A2;.;1&9.08
92A052?2A.9
Finally, Wnt/β-catenin signaling is the third transforming factor involved in posterior patterning
of the neural plate. Activation of this pathway during gastrulation represses anterior
development in contrast to its role of inducing the dorsoanterior axis in the early embryo.
Animal caps expressed hoxb9 (spinal cord) and krox20 (hindbrain) following treatment with both
Noggin and Wnt3a (McGrew et al., 1995). Introducing graded amounts of Dishevelled to
neuralized animal caps resulted in increasingly more posterior fates which suggested that graded
Wnt-signaling patterns the neural plate (Itoh and Sokol, 1997). Additional evidence to support
that a morphogen gradient of Wnt serves to pattern the neural plate emerged when
overexpressing the Wnt inhibitor Dkk in Xenopus embryos resulted in an expansion of the
anteriorly expressed genes bf-1 and otx2 and a posterior shift of krox20. Conversely, increasing
7
Wnt concentration results in the opposite phenotype: an anterior shift of krox20 and a repression
of anterior neural gene expression (Kiecker and Niehrs, 2001). Consistent with a graded level of
Wnt signaling in the neural plate, Kiecker and Niehrs (Kiecker and Niehrs, 2001) observed an
anterior-to-posterior gradient of nuclearly localized β-catenin in the neural plate of a gastrula
staged embryo. Several lines of evidence point to Wnts coming form the underlying paraxial
mesoderm to posteriorize the neural plate. In zebrafish, Wnt8 is expressed in the paraxial
mesoderm and knockdown results in a loss of posterior neural gene expression (Erter et al.,
2001; Lekven et al., 2001). Wnt3a in the dorsal paraxial mesoderm of Xenopus directly activates
meis3 in the hindbrain (Elkouby et al., 2010). Ultimately, knockout studies in mice provide
genetic evidence for Wnt-signaling in A-P patterning. Mice express wnt3a and wnt5a in the
paraxial mesoderm and mutants lack posterior structures (Greco et al., 1996; Yamaguchi et al.,
1999). Knockout of the Wnt antagonist dkk1 results in mouse embryos that lack forebrains
(Mukhopadhyay et al., 2001) while genetically increasing β-catenin in the forebrain results in
transformation to more posterior fates (Paek et al., 2012).
The above experiments suggested that Wnt acts as a classical morphogen to pattern the A-P axis
of the neural plate. While this may indeed be the case, visualization of a graded Wnt ligand has
not been reported. Furthermore, several findings have suggested that the ligand is poorly
soluble. Biologically active Wnt3a requires glycosylation and palmitoylation (Komekado et al.,
2007; Takada et al., 2006; Willert et al., 2003), post-translational modifications that make it
hydrophobic. While co-expression with Wnt antagonists results in greater diffusion of the Wnt
ligand in Xenopus (Mii and Taira, 2009), the different expression domains of these antagonists
suggest that the function of Wnt antagonists is unlikely to resolve the paradox of how an
insoluble protein can have long range effects. The duration of Wnt-signaling as apposed to the
concentration of the ligand provides an alternative explanation to a morphogen gradient of Wnt
patterning the amphibian neural plate. The more posterior regions stay in contact with the Wnt
source for longer periods during gastrulation than anterior or medial regions. Certainly a
uniform concentration of ligand signaling for different durations can have variable outcomes.
Premature inactivation of Sonic Hedgehog (Shh) signaling in the zone of of polarizing activity
(ZPA) results in a loss of posterior fates in the autopods of mice (Harfe et al., 2004). The
Australian two-toed skink expresses Shh in the ZPA for a shorter duration than its five-toed
relative (Shapiro et al., 2003). Therefore, it is possible that uniform doses of Wnt applied to
neurectoderm for different durations may elicit a full repertoire of anterior and posterior markers.
While this has yet to be carefully tested, it remains an intriguing possibility.
Open questions
Several questions of neural patterning remain unanswered. The lack of reverse genetic
mutagenesis techniques has confounded the study of noggin in amphibian development. Lossof-function studies were restricted to the use of morpholino oligonucleotides (MOs) that, while
effective in protein knockdown, are transient and not useful for examining later phenotypes. MO
knockdown of Noggin in Xenopus does not result in abnormal phenotypes while noggin mutant
mice have severe neural and skeletal deformities (Brunet et al., 1998; McMahon et al., 1998).
8
Either noggin is dispensable in amphibian development or current methods are not sensitive
enough to determine a role for this gene. The controversy over Bmp antagonist and neural
induction has cooled in recent years, however the contribution of individual Bmp antagonists in
amphibian development remains unanswered. The development of targeted mutagenesis
techniques via zinc-finger nucleases, TALENs, and CRISPR presents an opportunity to mutate
these genes and study their roles using genetic nulls.
The identification of Wnt, Fgf, and RA signaling as posteriorizers of the neural plate opens a
series of questions. What are the targets of these pathways and how are these targets regulated?
A handful of target genes have been identified, but there is yet no clear consensus on how graded
levels of these morphogens elicit specific expression domains of their targets. A thorough
understanding of the response elements that mediate expression will give insight to the
mechanisms of A-P patterning. Modern methods such as RNA- and ChIP-seq, combined with
the classical embryology techniques offered by Xenopus, make this organism an excellent model
for probing these questions.
Xenopus has proven to be an important model for developmental biology. Its large embryos are
well-suited for “cut and paste” experiments which provided some of the first demonstrations of
embryonic inductions. Expression screens and the identification of organizer-specific genes
made Xenopus extremely useful at the advent of modern molecular biology. With the dawn of
the genomics age, this genus continues to be a powerful system for understanding gene function
in development. The complete sequencing of the West African species Xenopus tropicalis
provided the first genome of an amphibian (Hellsten et al., 2010). This brought Xenopus
tropicalis into the genomics era and made whole-genome experiments possible. The recent
sequencing and assembly of the Xenopus laevis genome will provide the frog community as well
as the developmental biology community at large with powerful tools to uncover developmental
mechanisms and address these unanswered questions.
Summary of this thesis
The work in my thesis took advantage of both the classical embryology and genomic resources
offered by Xenopus. My goals were two-fold: (1) to establish reverse genetic strategies in the
frog at two proof-of-principle loci, including one implicated in embryo dorsalization and (2) to
utilize state-of-the art genomic approaches to investigate the role of Wnt in neural
posteriorization. I was fortunate to collaborate with the biotechnology company Sangamo
BioSciences that specializes in zinc-finger nuclease (ZFN) and TALEN design to induce
mutagenic DNA double-strand breaks at specific loci in a genome. I used this technology to first
demonstrate the efficacy by inactivating a Green Fluorescent Protein transgene. Next, I used
ZFNs designed to target the noggin locus. It’s well known that Noggin is sufficient to dorsalize
tissue, but evidence for its requirement in amphibian development was lacking. I was able to
successfully mutate the locus and generate several lines of Xenopus tropicalis carrying noggin
alleles with different levels of activity. Through breeding of a null allele to homozygosity, I
9
found noggin to be required for cranial skeletal development, specifically in the dorsoventral
patterning of the pharyngeal arches.
Having demonstrated a role for noggin in dorsoventral patterning, I next turned to A-P patterning
of the neural plate. As mentioned above, Wnt signaling is both necessary and sufficient to
induce posterior fates in neural precursors, yet the mechanism remains poorly understood. I
hypothesized that neural posteriorization via Wnt is mediated through transcriptional regulation
of target genes. Therefore, it was necessary to identify genes directly regulated by this pathway.
To that end, I carried out an expression screen using RNA-seq aimed at discovering genes that
are directly activated by Wnt in the neural plate. This screen was successful; I found over 200
genes that were upregulated in response to Wnt. In situ hybridization analysis of selected genes
showed a majority to be expressed in posterior regions of the embryo. Of particular note, two
Spalt-like (Sall) transcription factor family members identified in the screen, sall1 and sall4,
showed robust expression in posterior neural regions. The genes identified in this screen provide
a basis for understanding the link between activation of the Wnt pathway and posterior
patterning of the neural tube.
The final chapter of my thesis focuses on the function of Sall4, one of the Sall transcription
factors identified in the screen for direct neural targets of Wnt. Sall4’s role in mammalian stem
cell maintenance has been well documented but no role in A-P patterning has been described. I
found that sall4 is specifically expressed in the neurectoderm and TCF/LEF sites found in the
first intron of sall4 were enriched in β-catenin chromatin immunoprecipitations. Morpholino
oligonucleotide knockdown of Sall4 resulted in a loss of spinal cord development and an
upregulation of the pouV/Oct4 homologs. Ecotopic expression of pouV/Oct4 was sufficient to
block neural posteriorization and reducing pouV/Oct4 in Sall4 morphants rescued spinal cord
development. Finally, I found that Sall4 knockdown was epistatic to posteriorization by both Fgf
and RA signaling. This data presented a novel model of Wnt-induced neural patterning whereby
Wnt sends a permissive signal by activating sall4 in order to repress the pouV/Oct4 genes. The
neural plate is then competent to respond to instructive signals from Wnt/Fgf/RA following the
down-regulation of the pouV/Oct4 genes.
10
CHAPTER 2
Materials and Methods
Embryo and explant culture
Xenopus laevis embryos were collected, fertilized, and cultured according to (Sive et al., 2010)
and staged according to (Nieuwkoop, 1967). Xenopus tropicalis embryos were collected from
natural matings. To induce mating, female Xenopus tropicalis were injected with a priming dose
<3
B;6A@<35B:.;05<?6<;604<;.1<A?<=6;3<99<D21A52;2EA:<?;6;4/F./<<@A6;4
1<@2<3B;6A@5<?B9<;.;1=.6?21D6A5.:.926;720A21D6A5
B;6A@A52
=2C6<B@;645A;6:.9@D2?2A52;.99<D21A<.:=92E.;1=?<1B022:/?F<@<C2?.5<B?
period. Ectodermal explants (animal caps) were cut using fine watchmaker’s forceps from stage
9 embryos and cultured in ¾ NAM.
RNA and morpholino microinjections
All ZFN plasmids were linearized by restriction enzyme digest to produce transcripts with (AscI)
or without (NotI) .;&)=<9F.12;F9.A6<;@64;.9'?.;@0?6=A@D2?2@F;A52@6G21in vitro with a
&=:!2@@.42:!.056;286A:/6<; ZFN RNAs were microinjected into both blastomeres
of two-cell staged Xenopus tropicalis 2:/?F<@.9<;4D6A5=4<3:52??F%"D5<@2
RB<?2@02;A=?<1B0A@2?C21A<A?.08@B002@@3B99F6;720A212:/?F<@.;1@A.;1.?16G2:%"
amounts injected. Embryos tend to orient animal pole uppermost, so injection penetrated the
animal hemisphere, but mRNA was deposited near the center of the blastomere. Broad
6;52?6A.;02<3:%"D.@0<;Q?:21/FA5216@A?6/BA6<;<3:52??F3?<:6;720A21:%"
:/?F<@D2?2?.6@216;
!.?0P@%6;42?@<9BA6<;D6A542;A.:F06;!BA.;Anoggin RNA was
6;720A21C2;A?.9C242A.99F6;02992:/?F<@.9<;4D6A5=4LacZ RNA and cultured in 1/3
!.?0P@%6;42?@<9BA6<;D6A542;A.:F06;
Sall4 CS-108, Fgf8a CS-108, noggin CS-108, and β-catenin CS-108 were linearized with Asc1
and transcribed with a Sp6 mMessage mMachine kit (Ambion). The PouV genes (a gift from
Joshua Brickman), TVGR (Darken and Wilson, 2001), and nuclear β-galactosidase CS2+ were
linearized with Not1 and transcribed with Sp6. All RNAs were injected in either 5 or 10 ηL
bursts along with GFP and β-galactosidase RNAs to serve as tracers.
All morpholinos were injected in either 5 or 10 ηL bursts along with fluorescein-labeled control
morpholino (Gene Tools) to serve as a tracer. The Sall4 morpholino sequences are as follows:
morpholino 1: 5’- GCCAATTATTCCCTTTCTCCACCAC-3’ and morpholino 2: 5’GGTTCGGCTGCTTTCTCCTCGACAT-3’.
11
Western Blotting
Embryos were lysed in buffer containing 20 mM Tris pH 8.0, 50 mM NaCl, 2 mM EDTA, 1%
Triton X-100 and freshly supplemented with =?<A2.@26;56/6A<?A./92A@
[email protected]@D2?2
AF=60.99F0<;02;A?.A21A<
μL lysis buffer per embryo. $?<A26;@D2?2@2=.?.A21B@6;4
&&
$429@.;1/9<AA21<;A<;6A?<0299B9<@2:2:/?.;2@;.;A6/<1F?20<4;6G6;4A52 A.4
&64:.
169BA21
.;1.%$0<;7B4.A211<;82F.;A6:<B@2@20<;1.?F.;A6/<1F
.08@<; ./@169BA21
D2?2B@21A<12A20A A.4421=?<A26;@3?<:6;720A212:/?F<
[email protected]@;.;A6/<1F?20<4;6G6;40A6;&.;A.?BG169BA21
.;1.%$0<;7B4.A214<.A
.;A6?.//6A@20<;1.?F.;A6/<1F169BA21
D2?2B@21A<12A20A2;1<42;<B@0A6;[email protected]<.16;4
control.
Celery Extract Preparation for Cel-1 assays
4<33?2@[email protected]@(Apium graveolens var. dulce) were cut into 3-4 cm2 pieces and
?6;@21A5<?<[email protected]?99@B/@2>B2;A@A2=@D2?2=2?3<?:216;A52J?<<:
292?F=6202@D2?27B6021B@6;4.;#:24.:[email protected];47B602?.;1=.@@21A5?<B45.Q;2
:2@5@0?22;A<?2:<C2=.?A60B9.A2:.AA2?:9@<30292?F7B602D.@@=B;.AE43<?
:6;BA2@6;.=?20569921208:.;02;A?64B42D6A5.&?<A<?'52@B=2?;.A.;AD.@A?.;@32?21A<
.092.;/2.82?.;1'?6@9=.;1$!&D2?2.1121A<.Q;.90<;02;A?.A6<;<3
!
.;1
μ!?2@=20A6C29F'56@@<B9BA6<;D.@@A6??21@9<D29F3<?:6;D56924<3
.::<;6B:@[email protected];02;A?63B421.A
E43<?:6;BA2@'52@B=2?;.A.;A
[email protected];A?.;@32?21A<.092.;/2.82?.;1@9<D9F@A6??213<?:6;BA2@D56924<3
.::<;6B:@[email protected]'52@<9BA6<;D.@02;A?63B421.A
E43<?:6;BA2@'52
@B=2?;.A.;AD.@16@0.?121.;1A52=2992AD.@@B@=2;1216;:9@<3
!'?6@9!9
/B332?@B==92:2;A21D6A53?2@5$!&A<.0<;02;A?.A6<;<3
μ!'520292?F2EA?.0AD.@A52;
16.9FG21A5?<B45
!*#AB/6;4.4.6;@A <3
!'?6@9!9/B332?
'52/B332?D.@05.;4215<B?9F3<?5<B?@.;1A52;.99<D21A<16.9FG2<C2?;645A6.9FG21
2EA?.0AD.@A52;R.@53?<G2;6;μ .96>B<A@.;1@A<?21.AJ
Mutation Detection by Cel-1 from Celery Extract
;16C61B.9D5<92A.1=<92@<?A.69=6202@3?<:A.1=<92@@A.421.;19.A2?D2?29F@216;
μL
<3/B332?0<;A.6;6;4:!'?6@=:!".9:!'&&.;1
5292E®
#%.;1@B==92:2;A21D6A5µg/ml proteinase K. Lysates were
6;0B/.A216;.JD.A2?/.A5<C2?;645AD6A5=2?6<160C<?A2E6;42;<:60"D.@2EA?.0A21
/FQ?@A169BA6;[email protected]
μ <31<B/[email protected]?.;1A52;.116;4.;2>B.9C<9B:2
<3=52;<9059<?<3<?:6@<.:F9.90<5<9
'52@<9BA6<;D.@@5.82;C64<?<B@9F.;1
02;A?63B421<;.A./92A<=02;A?64B42.A
%!$3<?:6;BA2@'52.>B2<B@=5.@2D.@
A?.;@32??21A<.092.;AB/2.;1L <3!@<16B:.02A.A2.;1L <36@<=?<=F9.90<5<9D.@
.11213<99<D21/FC64<?<B@@5.86;4'52@.:=92@D2?2A52;02;A?63B4213<?
:6;[email protected]
%$!.A?<<:A2:=2?.AB?2'52@B=2?;.A.;AD.@16@0.?121.;1
L <32A5.;<9
[email protected]<D.@5A52=2992A3<99<D21/F02;A?63B4.A6<;.A
%$!3<?:6;BA2@'52
supernatant was decanted and the pellet allowed to air dry. Once dry, the pellet was resuspended
6;L <329BA6<;/B332?
:!'?6@9=<?2.05:BA.A6<;12A20A6<;?2.0A6<;
L 12
<3=B?6Q2142;<:60"3?<:.;6;16C61B.9D.@B@216;.L $%?2.0A6<;A<.:=963FA52
:BA.42;6G21?246<;<99<D6;4.:=96Q0.A6<;L D.@A?.;@32?21A<.092.;$%AB/2.;1D.@
denatured and re-annealed by incubating at 94°C for 5 minutes followed by cooling at a rate of
-2°@20A<°A52;.A.?.A2<3M
°C/sec until the reaction reached 25°C, then incubated at
J"2EAL <[email protected]<A52?2.0A6<;.;16;0B/.A21.AJ3<?
5<B?A<
.99<D3<?52A2?<1B=92E092.C.42'522;A6?2?2.0A6<;D.@A52;?2@<9C21<;.
.4.?<@2429
.;1=<@[email protected];21D6A52A5616B:/?<:612'<12A20A5<:<GF4<B@:BA.;A6;16C61B.9@:BA.;A
.:=960<;@D2?2:6E21D6A5A52$%=?<1B0A.;1@B/720A21A<12;.AB?.A6<;?2.;;2.96;4.;1
092.C.42C6.0292?F2EA?.0A*691AF=2.:=960<;@D2?2:6E216;A<612;A63F5<:<GF4<B@D691
AF=26;16C61B.9@
Cartilage staining
Tadpoles were fixed in 4% paraformaldehyde for 2-24 hours at room temperature in 4 mL vials.
Paraformaldehyde was decanted and the embryos were suspended in a sterile filtered solution of
acid/alcohol (70% ethanol and .37% HCl) containing 0.1% Alcian Blue. Vials were placed on a
rotator and gently mixed for 6-12 hours at room temperature when staining of the cartilage
elements becomes apparent. When staining was complete, the buffer was discarded and
tadpoles were resuspended in the acid/alcohol solution without alcian blue and rotated for 20
minutes at room temperature. This was repeated until the solution no longer had any blue tint.
Tadpoles were then rehydrated stepwise into water and then bleached in 1X SSC supplemented
with 1.2% hydrogen peroxide and 5% formamide for 1-2 hours on a white-light table. Vial caps
were removed to prevent excessive bubble formation. Following bleaching, tadpoles were
resuspended in a 2% KOH solution and rotated for 1 hour. Stained tadpoles were cleared by
successive 2 hour incubations in 2% KOH with increasing concentration of glycerol. Once
cleared, tadpoles were either directly imaged or flat-mounted on a microscope slide by fine
dissection and imaged.
Cycloheximide and dexamethasone treatments
Noggin RNA and an inducible Wnt agonist, TVGR RNA were injected animally into both
blastomeres of two-cell embryos. Embryos were cultured until stage 9 when animal caps were
excised and cultured with or without 10 µM dexamethasone (Sigma) to activate Wnt signaling.
To block protein translation, animal caps were pre-treated with 5 µM cycloheximide (Sigma) for
1.5 hours prior to dexamethasone addition. Animal caps were cultured until the stage 15
equivalent and total RNA was harvested using Trizol (Invitrogen).
Whole-mount in situ hybridization
Embryos were stained after whole mount in situ hybridization as described in (Harland, 1991).
β-galactosidase staining was carried out as described in (Fletcher et al., 2006).
13
Embedding and Sectioning
Embryos for sectioning were first equilibrated into a 30% sucrose solution and then transferred
into a PBS solution containing 20% sucrose, 30% BSA, 4.9% gelatin. Embryos for sectioning
were quickly transferred to fresh buffer supplemented with 1.5% glutaraldehyde and allowed to
harden in peel-away plastic molds. Embedded embryos were cut to about a 0.5cm3 block with a
razor blade and super-glued to an attack die from the game Heroscape®. Mounted embryos
were sectioned on a Pelco 101 vibratome while submerged in PBS.
RT-PCR and qPCR
RNA was isolated from either whole embryos or animal caps using Trizol according to standard
protocols and 1µg total RNA was reverse transcribed with either MMLV reverse transcriptase
(Promega) or iScript (BIO-RAD) for semi-quantitative or qPCR, respectively. Semi-quantitative
PCRs included trace amounts of 32P labelled dCTP (Perkin-Elmer) in the reaction and were
analyzed during the log-phase of amplification. qPCR reactions were amplified on a CFX96
(BIO-RAD) light cycler. ernithine decarboxylase (ODC) and eukaryotic elongation factor-1a1
(eef1α1) were used for internal controls. All primers annealed at 60°C and are listed in appendix
II.
RNA-seq
RNA-seq was performed according to standard protocols from Illumina as described in
(Dichmann and Harland, 2012). mRNA was purified from 10 µg total using oligo-dT Dynabeads
(Invitrogen) and fragmented using zinc ion fragmentation buffer (Ambion) for 1.5 minutes at
70°C. First strand synthesis was carried out according to standard protocols using Superscript II
(Invitrogen) and second strand synthesis was performed using DNA polymerase I (NEB).
cDNA fragment ends were repaired and ends were adenylated using Klenow, T4 DNA
polymerase and T4 PNK (NEB). Adaptors (Illumina) were ligated using T4 Ligase and Quick
Ligase buffer (NEB). AMPure XP beads (NEB) were used to select for fragments larger than
200 bp. The library was amplified with Phusion HF polymerase (NEB) and single-end 76basepair reads were sequenced on an Illumina Genome Analyzer II. All reads were mapped to an
index created from a collection of full-length Xenopus laevis mRNA sequences (http://
xgc.nci.nih.gov) using TOPHAT and BOWTIE (Langmead et al., 2009; Trapnell et al., 2009).
Analysis of transcript abundance differences was done using CUFFDIFF (Trapnell et al., 2010).
Chromatin immunoprecipitation
FLAG-β-catenin-injected embryos for immunoprecipitation were fixed in 1% formaldehyde/PBS
for 1 hr, quenched with 0.125 M glycine/PBS for 15 minutes followed by three 10 minute
washes in PBS. Lysis was performed according to (Blythe et al., 2009). Chromatin was sheared
on ice using a Branson Model 450 digital sonifier with a Model 102C probe for 24 ten second
bursts set at 30% amplitude. Immune complexes were pulled down using M2 FLAG antiboby
14
(Sigma) bound to anti-mouse magnetic beads (Invitrogen). Samples were washed, cross-links
reversed and DNA isolated according to ChIP protocols (Blythe et al., 2009). ChIP DNA was
quantified with SYBR-green PCR mix (BIO-RAD) on a CFX96 light cycler (BIO-RAD).
Enrichment was calculated by comparing the %input between samples. Uninjected embryos
served as a control for non-specific binding. xmlc2 (Blythe et al., 2009) and meis3 (Elkouby et
al., 2010) served as negative and positive controls for β-catenin binding, respectively.
15
CHAPTER 3
Analysis of the developmental role of noggin in Xenopus tropicalis
development via Zinc-Finger Nuclease mutagenesis
Introduction
Frogs of the genus Xenopus have been an important model organism for cell and developmental
biologists since the 1930s (Gurdon and Hopwood, 2000). X. laevis is the standard model, but due
to its allotetraploid genome, less suited for genetic approaches than the diploid X. tropicalis,
whose genome sequence has been determined (Hellsten et al., 2010). Whereas embryological
manipulations and gain-of-function experiments are major strengths of Xenopus, reverse genetics
are currently limited to the use of antisense reagents that provide transient and often incomplete
gene knock down (Eisen and Smith, 2008). The ability to introduce targeted, heritable mutations
that disrupt gene function has remained elusive.
This work provides a generally applicable solution to this problem: targeted gene disruption with
designed zinc finger nucleases (ZFNs). ZFNs are the fusion of the non-specific cleavage domain
of the Type IIS restriction enzyme FokI to a zinc-finger protein (Miller et al., 1985; Pavletich and
Pabo, 1991) that is engineered to bind a specific genomic locus in order to induce a targeted
double-strand break (DSB). Pioneering studies in oocytes of X. laevis (Bibikova et al., 2001)
and subsequent work in Drosophila (Bibikova et al., 2002) showed the mutagenic potential of a
DSB induced by ZFNs (reviewed in ref (Carroll, 2008; Urnov et al., 2010)). Resolution of ZFNinduced DSBs via non-homologous end joining (NHEJ) generates small insertions and deletions
which often produce null or hypomorphic alleles (Bibikova et al., 2002; Perez et al., 2008;
Santiago et al., 2008).
The Bone morphogenic protein (Bmp) antagonist noggin is expressed in Spemann’s organizer
and is a potent dorsalizing factor (Smith and Harland, 1992; Zimmerman et al., 1996). Together
with two other Bmp antagonists, Chordin and Follistatin, it serves to dorsalize the ectoderm and
mesoderm to give rise to neural and somitic derivatives, respectively (Khokha et al., 2005).
Targeted deletion of the noggin locus in mice results in perinatal lethality, spina bifida, and
skeletal deformities (Brunet et al., 1998; McMahon et al., 1998). However, antisense
morpholino oligonucletide (MO) knockdown of Noggin in Xenopus does not yield a phenotype.
Whether this is due to incomplete knockdown or a role for Noggin beyond the effective window
of MO activity remains unknown. Genetic inactivation of the locus is required to effectively
assay the function of Noggin in amphibian development, a method previously restricted to nonspecific, forward genetic screens and laborious mapping efforts.
I developed an effective protocol for gene disruption in X. tropicalis. Using ZFNs designed
against a reporter transgene and the noggin locus, the delivery and expression conditions for
ZFNs were optimized which resulted in high frequencies of somatic and germline mutations that
16
were transmissible to the next generation. Furthermore, ZFN induced mutations in noggin were
bred to homozygosity which yielded an unexpected phenotype.
Results
Xenopus eggs are large and easily manipulated (Sive et al., 2010), offering the opportunity to
deliver ZFNs via injection of mRNA, a method that has been successful in driving ZFN-induced
gene disruption in other organisms (Carroll, 2008; Urnov et al., 2010).
To develop conditions for gene disruption in Xenopus tropicalis, transgenic animals carrying a
single-copy GFP transgene were used(Hamlet et al., 2006). Wild type X. tropicalis eggs were
fertilized with sperm from a homozygous GFP transgenic male. The resulting heterozygous
embryos were injected with mRNA encoding ZFNs that target the eGFP coding region (Geurts et
al., 2009). Uninjected tadpoles express GFP robustly in the somites, lens, and head musculature
(Figure 3.1 A-C). Injection of 20 pg eGFP ZFN RNAs led to mosaic loss of fluorescence in
otherwise healthy tadpoles (Figure 3.1 D-F). At a higher dose of ZFNs, most cells had lost
fluorescence, suggesting efficient somatic mutation of the transgene (Figure 3.1 G-I).
To determine whether loss of fluorescence resulted from a ZFN-induced mutation in the eGFP
transgene, the target locus was genotyped using an assay based on the mismatch-sensitive
endonuclease, Cel-1 (Miller et al., 2007). This analysis (Figure 3.1 J) demonstrated that ZFNtreated, but not control, tadpoles had acquired a DNA sequence alteration in the stretch targeted
by the ZFNs. Sequencing subsequently revealed that individual tadpoles often carried multiple
distinct indels ranging from 5 to 20 bp centered over the ZFN recognition site (Figure 3.1 K), a
signature of mutagenic NHEJ. Taken together, these experiments show that ZFN mRNA
injection into the two-cell embryo yields tadpoles without detectable developmental defects and
exhibit both genetic and phenotypic mosaicism for the ZFN-targeted locus and trait, respectively.
To determine whether this approach can be used to disrupt an endogenous gene, ZFNs that target
the noggin locus were designed. Noggin is a Bmp antagonist that contributes to dorsal/ventral
patterning during gastrulation in Xenopus (Smith and Harland, 1992). Although its function in
later development has been studied in human patients (Marcelino et al., 2001) and mice
(Bachiller et al., 2000; Brunet et al., 1998; McMahon et al., 1998; Warren et al., 2003), its
developmental role in non-mammalian vertebrates remains poorly understood. Therefore,
mutant alleles of the endogenous noggin gene are required to probe its role throughout
amphibian development.
A panel of ZFNs targeting noggin was designed by Sangamo Biosciences Inc., screened in a
budding yeast proxy system (Doyon et al., 2008; McCammon et al., 2011), and cloned into an
expression construct that allows the synthesis of an efficiently translated mRNA with a
stabilizing polyadenylation signal (Turner and Weintraub, 1994). Embryos were injected in the
animal pole with the RNA deposited in the center of each blastomere at the two-cell stage, raised
to stage 40, and DNA was isolated from tadpoles that exhibited broad mCherry (i.e., tracer)
17
expression. Use of ZFNs that carry a wild-type FokI endonuclease domain yielded a significant
fraction of embryos with developmental defects (Figure 3.2 A). However, this was not observed
when RNA expressing the same zinc finger DNA recognition domains fused to the obligate
heterodimer forms of FokI was injected. The point mutations in FokI are made in the nuclease
domain to prevent the formation of a homodimeric and functional nuclease (Miller et al., 2007).
These mutations, E490K, I538K and Q486E, I499L, are made in the FokI domain of the left and
right ZFN, respectively (referred to as EL+KK). Even at the highest tested doses of such ZFN
mRNA, greater than 60% of the injected embryos developed normally (Figure 3.2 A).
To optimize the delivery of ZFNs, I also tested whether unstable, non-adenylated RNA, which
would be translated early and deliver a transient burst of ZFN, might be superior to the extended
expression of ZFNs from transcripts that are cleaved and polyadenylated after injection (Turner
and Weintraub, 1994). As expected, the non-adenylated transcripts led to considerably less
protein expression (Figure 3.2 C). Even at higher doses, they did not induce noggin gene
disruption at a frequency measurable by Cel-1. Thus, the prolonged presence of ZFNs from
adenylated RNA is superior for effective genome editing in Xenopus.
Because it is difficult to predict a priori the extent to which ZFN overexpression may cause
embryonic defects, Sangamo tested a panel of ZFNs in a yeast-based single-strand annealing
assay (Doyon et al., 2008) and I tested them in embryos (Figure 3.2 B, Table 3.1). Six ZFN pairs
shown to have activity in the yeast assay were chosen for testing in Xenopus. The ZFN pairs
found to be active in yeast and well tolerated in tadpoles also showed efficient genome editing in
Xenopus embryos as measured by sequencing noggin amplicons from injected tadpoles.
Since no Xenopus strains carrying noggin mutations existed, it wasn’t possible to screen for
phenotypes on a heterozygous background (Doyon et al., 2008), therefore I screened the ZFNs
for activity by genotyping the targeted region using the Cel-1 endonuclease (Figure 3.3 A). All of
the ZFNs that were well tolerated produced targeted gene disruption. Direct sequencing of the
nuclease-targeted region in tadpoles injected with ZFNs revealed a broad panel of insertions and
deletions ranging in size from 5 to 195bp (Figure 3.3 B), with frequencies of mutant amplicons
from 10-47%. Such high rates of somatic mutagenesis suggested that tadpoles might also carry
mutations in the germline, allowing the establishment of lines carrying novel noggin alleles.
I injected wild-type embryos at the two-cell stage with 100 pg of mRNA encoding ZFNs that
target noggin and were tolerated by greater than 60% of injected embryos (Figure 3.2 B).
Successful somatic genome editing in tadpoles and froglets was confirmed by isolating genomic
DNA from tail or toe clips, respectively, and genotyping the noggin locus by Cel-1 and
sequencing (Table 3.1). Injected embryos were raised to sexual maturity and outcrossed to wildtype animals. Offspring from this cross were raised to stage 40, lysed and analyzed via Cel-1 for
mutations in the noggin locus. Genotyping offspring from a cross using a ZFN-treated male
founder revealed that three of eighteen tadpoles had inherited a ZFN-induced Δ12 allele of
noggin (Figure 3.4 A,B). A second male founder produced six out of fifty embryos heterozygous
for a ZFN-induced three base pair insertion allele (Figure 3.4 A,B) and a third male produced
18
twelve of fifty tadpoles heterozygous for a four basepair insertion. The latter ZFN-induced
noggin allele induces a frameshift mutation that results in a premature stop codon at position 55.
Therefore, it is likely to behave as a null allele because the resulting protein lacks most of the
Bmp binding residues and all of the residues required for dimerization (Groppe et al., 2002).
The recovery of these mutations in the offspring of adult animals raised from injected embryos
demonstrates that ZFN-induced alleles of an endogenous gene can be transmitted to the next
generation. Of significant note, the parent and heterozygous tadpoles carrying mutant alleles
were indistinguishable from wild type siblings, indicating this is an effective approach to
establish lines of animals carrying novel alleles of investigator-specified genes. To test whether
ZFN-induced mutagenesis caused off-target mutations, gynogenotes from noggin ZFN injected
females that lacked mutant noggin alleles in their germline were made. These were siblings of
the germline mutated males, and from clutches that showed a high frequency of somatic
mutation. Gynogenesis diploidizes activated eggs by preventing the extrusion of the second
polar body and serves to homozygose recessive mutations (Khokha et al., 2009). While offtarget mutations would result in high frequencies of mutant embryos in gynogenotes produced
from these females, I did not detect such mutations; indeed 87% of gynogenotes were
indistinguishable from wild-type tadpoles and the remaining 13% had various mediolateral and
dorsoventral defects, consistent with reported phenotypes and frequencies from young wild-type
females (Grammer et al., 2005). This result demonstrates that potentially confounding off-target
mutations in founder animals are negligible.
Noggin induces ectopic dorsal tissue when expressed on the ventral side of Xenopus embryos
(Smith and Harland, 1992), which serves as an excellent test for whether the ZFN-induced
noggin alleles produce loss-of-function proteins. Full-length mutant alleles were cloned and
corresponding mRNAs for injection were synthesized. 5 or 10pg was injected into the ventral
vegetal blastomeres of 4-cell stage Xenopus laevis embryos (Figure 3.4 C). Embryos were
cultured to stage 28 and scored for the presence or absence of an ectopic axis (Figure 3.4 D).
The four basepair insertion allele failed to induce any ectopic dorsal tissues in embryos (Figure
3.4 H,I,J,J’), consistent with a loss-of-function frameshift mutation. LacZ mRNA was used as a
tracer and showed that injected cells populated the ventral posterior of injected tadpoles. The
∆12 allele induced ectopic axes in 8 and 16 percent of embryos when injected with 5 pg and 10
pg mutant noggin RNA, respectively. Interestingly, the induced axes in these embryos were
underdeveloped, suggesting that this allele functions as a hypomorph (Figure 3.4 M arrowheads).
Finally, the three basepair insertion allele was indistinguishable in activity from wild-type noggin
(Figure 3.4 N,O,P). These results demonstrate that specific loss-of-function mutant lines of X.
tropicalis can be generated via targeted ZFN mutagenesis. Furthermore, the mutations in noggin
form the basis of an allelic series that will be useful to examine noggin function in later
development.
Knocking down Noggin, Chordin, and Follistatin results in a loss of dorsal structures and yield
embryos without neural or somitic derivatives whereas knocking down any two in combination
results in a redution but not a complete loss of these structures (Khokha et al., 2005). To test if
19
the 4-basepair insertion allele is truly a null allele, heterozygous adults were crossed and the
resulting embryos were injected with morpholinos targeting Chordin and Follistatin. These
embryos were raised to stage 14 and assayed for sox2 and myoD expression. Uninjected
embryos expressed sox2 throughout the neural plate and myoD in the underlying mesoderm
(Figure 3.5 A,D). Chordin and Follistatin morpholino injection resulted in a reduction of both
sox2 and myoD in ~75% of the embryos (Figure 3.5B,E). However, in approximately 25% of
injected embryos, presumably those homozygous for the 4-bp insertion allele, there was a
complete loss of both sox2 and myoD expression (Figure 3.5C,E). These results, taken with the
failure of RNA bearing the 4-basepair insertion noggin allele to induce ectopic axes suggest that
the 4-bp insertion noggin allele is a true functionally null allele.
noggin null mice die at birth due to numerous defects including open neural tubes and several
skeletal abnormalities including chondrocyte hypertrophy and joint formation failure (Brunet et
al., 1998; McMahon et al., 1998). Conversely, morpholino knockdown of Noggin in Xenopus
does not yield a phenotype (Khokha et al., 2005). This discrepancy can be due to either
incomplete knockdown by the morpholino or because the requirement for Noggin occurs when
morpholinos are no longer effective. To distinguish between these possibilities, heterozygous
adults carrying the 4-bp insertion allele were crossed and the resulting embryos were cultured
and observed for phenotypic abnomalities. Unexpectedly, I did not observe defects in neural
tube closure in any of the embryos produced from heterozygous parents (Figure 3.6 A-J). No
defects were observed in any embryos until stage 45 where a dorsal-rostral protuberance was
noticed in a subset of the clutch . Sorting tadpoles based on the presence or absence of this
dorsal “horn” resulted in 2533 (75.1%) wild-type and 824 (24.9%) abnormal (Figure 3.6 K-N
and Figure 3.7 A-F). Genotyping 10 wild-type and 10 horned tadpoles revealed the horned
tadpoles were homozygous for the noggin null allele.
To determine the nature of the horn observed in noggin mutants, alcian blue staining was used to
viualize the cartilage skelton of the cranium. Staining and flatmounting cartilage preparations
from wild-type (Figure 3.7 G,G’) and mutant tadpoles revealed overall smaller cartilage
elements, deformed ceratobranchial cartilage, and most notably severely reduced Meckel’s
cartilage that are fused to the palatoquadrate in mutant tadpoles (Figure 3.7 H,H’). There are no
major differences in the superostral plate between the wild-type and mutant, indicating that the
horn observed in the mutants is due to a deformation and loss of ventral structures (i.e. Meckel’s
catilage and ceratohyals) rather than an overgrowth of the superostral plate. Mutant tadpoles did
not display sensory defects as they responded to gentle tapping on the culture dish. Finally, the
mutants began to die off rapidly by two weeks post fertilization (Figure 3.7 I), presumably due to
malnutrition because of an inability to feed effectively.
Next, expression of the neural crest gene sox9 was examined because it directly activates col2a,
which is required for cartilage differentiation(Lefebvre et al., 1997). A reduction in cartilage
differentiation could explain the overall smaller cranium size observed in mutants. Since the
mutant phenotype doesn’t manifest until after patterning and differentiation of the pharyngeal
arches, it was necessary to genotype tadpoles prior to analyzing gene expression. To that end,
20
the heads from stage 33 and stage 39 tadpoles were excised and processed for in situ
hybridization while the tails were genotyped. There was a marked reduction in sox9 expression
(Figure 3.8 A,B) and a corresponding loss of col2a expression in the mandibular arch of mutants
at stage 33 (Figure 3.8 C,D arrowhead), consistent with the observed reduction in Meckel’s
cartilage. However, there was no noticeable difference in col2a expression between wild-type
and mutant tadpoles at stage 39 (Figure 3.8 E,F).
Bmp signaling plays a major role in the dorsal-ventral patterning of the pharyngeal arches, which
in turn develop into the cranial skeleton. Previous work in zebrafish has shown that Bmp
signaling is sufficient to induce ventral arch fates in intermediate and dorsal regions (Alexander
et al., 2011; Zuniga et al., 2011). Given that Noggin functions as a Bmp antagonist, I examined
wild-type and mutant tadpoles for the expression of Bmp responsive genes that function in
pharyngeal arch development. bmp7 is expressed in the ventral portion of the pharyngeal arches
but did not show any appreciable differences between wild-type and mutant tadpoles at stage 33
or 39 (Figure 3.9 A-D). At stage 33, expression of the Bmp responsive gene msx2 (Hollnagel et
al., 1999; Tríbulo et al., 2003) showed slight dorasl expansion (Figure 3.9 E,F) but was
indistinguishable from wild-type at stage 39 (Figure 3.9 G,H). Another Bmp target, endothelin1
(edn-1) was not differently expressed at stage 33 between the two genotypes (Figure 3.9 I,J).
However, it showed a more diffuse expression pattern in the mutant at stage 39 (Figure 3.9 K,L).
Finally, the BMP target gene hand2 (Howard et al., 2000; Xiong et al., 2009) had a broader
expression domain in the mutants at stage 33 (Figure 3.9 M,N) and was ectopically expressed in
the mandibular arch of stage 39 mutants (Figure 3.9 O,P). In zebrafish, Hand2 is known to
repress bapx1 which is required for joint formation between Meckel’s cartilage and the
palatoquadrate (Miller et al., 2003).
Discussion
These results show that Xenopus can be added to the growing list of important model organisms
for which ZFN-encoding mRNA has allowed facile reverse genetics, including Drosophila
(Bibikova et al., 2001), zebrafish (Doyon et al., 2008; Meng et al., 2008), and the rat (Geurts et
al., 2009; Mashimo et al., 2010).
An important requirement for the use of ZFNs is a streamlined protocol to predict and implement
effective gene disruption. This work confirmed that a proxy assay in budding yeast is efficient in
identifying ZFNs that will function effectively in the developing embryo. Furthermore, I have
determined an expression vector architecture and dose of mRNA that allow optimal expression
of the ZFNs to generate both tadpoles and fertile adult animals that carry disrupted alleles of
target genes in both the soma and germline. Remarkably, the injection of ZFN mRNA into a
relatively small cohort of two-cell embryos was sufficient to raise adults carrying mutant alleles
in the germline. Our results demonstrate that with optimal husbandry (Grammer et al., 2005),
homozygous mutants can be generated within one year of the initial mutagenesis, and potentially
7 months if female founders are generated.
21
The Cys2-His2 zinc finger protein (Miller et al., 1985; Pavletich and Pabo, 1991) is the most
common DNA recognition motif in metazoa, and ZFNs can be engineered against any locus of
interest (Urnov et al., 2010). This work here showed that ZFNs built using an archive of prevalidated two-finger modules that either target eGFP or noggin and carry high-fidelity FokI
endonuclease domains (Miller et al., 2007) induced mutations at a high rate when injected at
doses that were well tolerated by the majority of injected embryos. Direct sequencing of both
targets showed a variety of indels that are likely to result in null alleles. Indeed, there was a
ZFN-induced loss of eGFP fluorescence phenotype in injected tadpoles that are heterozygous for
the eGFP transgene. The absence of a phenotype in founder noggin animals could be due to
several factors, including the mosaic nature of the injected founders, non-autonomy of secreted
Noggin, and a compensatory role of other Bmp antagonists in the early embryo (Khokha et al.,
2005). However, artificial ventral expression of mRNAs encoding the ZFN-induced noggin
mutants demonstrated that ZFNs induced both null and hypomorphic alleles of an endogenous
X. tropicalis gene. The normal development of gynogenotes derived from females with high
frequencies of somatic mutations in noggin (but no detectable noggin mutations in the germline)
shows that off-target mutations in the germline must be rare.
Breeding the induced noggin mutant alleles to homozygosity revealed that Noggin functions in
the development and differentiation of the pharyngeal arches. Specifically, Noggin appears to be
required to restrict expression of the Bmp target gene hand2. Mutant tadpoles express hand2 in a
broader domain, including the mandibular arch which gives rise to Meckel’s cartilage. This
provides a potential mechanism to explain the loss of joint formation between Meckel’s cartilage
and the palatoquadrate observed in mutant tadpoles. Hand2 represses the homeobox gene bapx1
(Miller et al., 2003). It is plausible that in noggin mutant tadpoles, the increase in Bmp signaling
due to a loss of Noggin, results in the expansion of Hand2 and a concomitant repression of
bapx1, causing the fusion of Meckel’s cartilage to the palatoquadrate (Figure 3.7 H,H’). This
may also explain the cause of death observed in mutants two weeks post fertilization. Meckel’s
cartilage forms the mandible and the severe reduction/loss of this structure in noggin mutants
impedes the tadpole’s ability to feed effectively and likely results in starvation. This is consistent
with the inverse sigmoidal survival curve observed in the mutants.
As mentioned above, noggin mutant mice show neural tube closure defects (McMahon et al.,
1998). While the skeletal deformaties are consistent between mutant frogs and mice, why are
there no defects in the neural tube observed in noggin mutant Xenopus tadpoles? One
explanation could be the presence of other Bmp antagonists. Chordin and Follistatin are
expressed in a similar domain during early embyogenesis of Xenopus (Khokha et al., 2005) and
could compensate for the loss of noggin in neural patterning. Indeed, there is redundancy in the
function of these Bmp antagonist during neural induction and dorsalization of the mesoderm
(Khokha et al., 2005). A second explanation could be that the paralog noggin2 partially
compensates for the loss of noggin (Fletcher et al., 2004).
Genome editing using ZFNs has the potential to enable numerous lines of experimentation that
were previously impossible with existing Xenopus methodology. Permanent, heritable mutations
22
will allow for the study of specific genes and later developmental processes without concern for
the off-target or transient effects associated with morpholino oligonucleotides (Eisen and Smith,
2008). Furthermore, the variety and size of ZFN-induced indels can be used to generate an
allelic series of mutations. We note that several noggin alleles we generated were deletions of
substantial size (~200 bp), indicating that ZFNs provide an attractive method for not only
disrupting specific coding sequences, but also for targeting regulatory elements in the genome.
Recently, alternative methods for genome editing have been developed that have replaced the use
of ZFNs . Transcription activator-like effectors (TALEs) from the plant pathogenic bacteria
Xanthomonas that consist of repeated motifs, each of which binds a single nucleotide (Boch et
al., 2009; Moscou and Bogdanove, 2009), have been fused with FokI nucleases to generate
TALE-Nucleases (Cermak et al., 2011; Christian et al., 2010). The modular nature of TALENs
allows for easy design and synthesis. Other groups have been successful in using TALENs to
generate biallelic mutations in Xenopus tropicalis (Lei et al., 2012). The CRISPR-CAS method
provides an additional alternative to nuclease fusion proteins for inducing targeted mutations.
This system uses an RNA molecule engineered to target specific sites that flank common repeat
sequences in the genome and recruit the Cas-9 endonuclease to induce double strand breaks
(Hwang et al., 2013; Jinek et al., 2012). The CRISPR-CAS system relies on basepair
complementarity and requires only two plasmids to synthesize targeting constructs. The
zebrafish genome has CRISPR sites every 8-128 basepairs thus, this system is likely applicable
to any gene of interest (Blackburn et al., 2013). My thesis work demonstrates that induced
mutations can result in heritable null mutations and allow for analysis of gene function in
amphibian development.
Due to the large size, abundance, and ready manipulation of their eggs and embryos, Xenopus
has provided important insights in both cell biology (King et al., 1996) and embryology (Harland
and Gerhart, 1997). Completion of the genome sequence of X. tropicalis brought this model
system into the genomics age (Hellsten et al., 2010), but the genetic engineering tools essential
for comprehensive study of biological mechanism were lacking. My studies add a robust method
for genome editing so that gene function can be analyzed throughout development.
23
Figure 3.1: Disruption of the eGFP transgene in Xenopus tropicalis using ZFNs.
(A-C) Uninjected tadpoles (UC). (D-F) Tadpoles injected with 20 pg of eGFP ZFN mRNA and
200 pg mCherry RNA (to monitor injection). (G-I) Heterozygous eGFP tadpoles injected with
50 pg eGFP ZFN mRNA and 200 pg mCherry RNA (tracer). (A,D,G) Brightfield. (B,E,H) eGFP
expression in tadpoles from A,D, and G, respectively. (C,F,I) Enlarged view of eGFP expression
in B, E, and H, respectively. (J) Cel-1 digestion of eGFP amplicons. Bands migrating at 345 bp
are full-length amplicons, Cel-1 cleavage products migrate at 246 bp and 99 bp. The fraction of
modified chromatids detected by Cel-1 are quantified as % NHEJ. “UC,” uninjected control. (K)
Sequence alignment of ZFN-induced mutant eGFP transgene alleles from tadpoles injected with
50 pg ZFN mRNA. Red nucleotides indicate insertions and dashes represent deletions.
Horizontal bold lines indicate ZFN binding sites.
24
25
% NHEJ
99 bp
246 bp
345 bp
J
C
B
A
Figure 1
16
1
11
2
20 pg
11
3
19
1
U.C.
U.C.
17
2
50 pg
F
E
D
24
3
UC
eGFP ZFN L
eGFP ZFN R
50 pg eGFP ZFNs
Gln Leu Gln Gln Pro Gln Arg Leu Tyr His Gly Arg Gln
I
H
50 pg eGFP ZFNs
Insertions
Indels
CAACTACAACAGCCACAACGTCTATATATCATGGCCGACA(+2)
GAACTACAACAGCCACAACGTGTGT--CATGGCCGACAAG(∆6,+4)
CAACTACAACAGCCACAACAA------CA--GCCGACAAG(∆12,+4)
CAACTACAACAGT--------------CATGGCCGACAAG(∆15,+1)
CAACTACAACAG----AA------------AGCCGACAAG(∆19,+3)
CAACTACAACAGCCACAAC----GGATCATGGCCGACAAG(∆6,+2)
CAACTACAACAGCCACAACGT---TATCATGGCCGACAAG(∆3)
CAACTACA------------TCTATATCATGGCCGACAAG(∆12)
CAACTACAACAG--------------------CCGACAAG(∆20)
CAACTACAACAG--------------------CCGACAAG(∆20)
CAACTACAACAG--------------------CCGACAAG(∆20)
CAACTACAACAG--------------------CCGACAAG(∆20)
CAACTACAACAGCCACAACGT-----------CCGACAAG(∆11)
CAACTACAACAGCCACAAC-----TATCATGGCCGACAAG(∆5)
CAACTACAACAGC--------------CATGGCCGACAAG(∆14)
CAACTACAACAG--------------------CCGACAAG(∆20)
CAACTACAACAGC--------------CATGGCCGACAAN(∆14)
CAACTACAACAG--------------------CCGACAAG(∆20)
CAACTACAACAG--------------------CCGACAAG(∆20)
CAACTACAACAG--------------------CCGACAAG(∆20)
CAACTACAACAGC--------------CATGGCCGACAAG(∆14)
CAACTACAACAGCCACAA--------TCATGGCCGACAAG(∆8)
WT CAACTACAACAGCCACAACGTCTATATCATGGCCGACAAG
Deletions
K
20 pg eGFP ZFNs
20 pg eGFP ZFNs
G
Figure 3.2: Tolerance and activity of ZFNs targeting noggin in Xenopus tropicalis.
(A) Optimization of ZFN delivery in Xenopus tropicalis. EL+KK: ZFNs with the EL and KK
modifications in the FokI domain (Miller et al., 2007), numbers represent different noggin ZFN
pairs (Table 3.2). WT: ZFNs with wildtype Fok1 nuclease domains. PA-: ZFN transcripts
lacking a poly-adenylation signal. UC: Uninjected control. (B) Comparison of activities of
different noggin ZFN pairs in the yeast activity assay and in injected tadpoles. Tadpoles were
injected with 100 pg of ZFN mRNA. Yeast activity values represented as a percentage relative to
expression of ZFNs targeting the human CCR5 gene (Perez et al., 2008). Activity in tadpoles as
calculated by the percent of mutant amplicons sequenced from injected embryos. ND: No Data.
(C) Western blot for FLAG tagged ZFN proteins.
26
A
100%
Tadpole % Phenotype
80%
60%
40%
20%
180
160
Yeast Assay % Control
100
50
20
eGFP ZFN EL+KK
100
50
100
Noggin
26EL+28KK
50
Noggin
60EL+66KK
Normal
Mild Axial Defects
Severe Axial Defects
Gastrulation Defects
Lethal
% Mutant Amplicons
Relative Activity in Yeast
50
20
100
Noggin
60+66 (WT)
50 pg ZFN
100
140
120
80
100
60
80
60
40
40
20
20
0
RNA
Noggin
60+66 (WT)
PA-
ND
ND
28EL+29KK
Tadpole % Phenotype
B
UC
Tadpole
0%
0
30EL+31KK
26EL+28KK
58EL+66KK
60EL+66KK
63EL+66KK
Noggin ZFN Pair
C
FLAG
UC
eGFP ZFN
Noggin
26EL+28KK
50
50
100
100
Noggin
60EL+66KK
50
100
Noggin
Noggin
60+66 (WT) 60+66 (WT)PA-
50
100
50
100
pg ZFN RNA
27
Figure 3.3: ZFN-driven editing of the noggin locus in Xenopus tropicalis. (A) Somatic
mutations in noggin detected by Cel-1. Bands migrating at 450 bp are full-length noggin
amplicons. Bands migrating at 300 bp and 150 bp bands are Cel-1 digest products. (B)
Sequence alignment of noggin alleles induced by indicated ZFN pairs. Red nucleotides indicate
insertions and dashes represent deletions. Horizontal bold lines indicate ZFN binding sites. EL
+KK: ZFNs with the EL and KK modifications in the FokI domain (Miller et al., 2007), numbers
represent different noggin ZFN pairs (Table 3.2).
28
A
100pg Noggin ZFN mRNA
Wild Type
58EL+66KK
60EL+66KK
63EL+66KK
450bp
300bp
150bp
Noggin ZFN L
B
Noggin ZFN R
Leu Ile Glu His Pro Asp Pro Ile Tyr Asp Pro Lys Glu Lys Asp
WT TGGACCTTATTGAGCATCCGGATCCTATCTATGATCCCAAGGAGAAGGATCTTA
58EL+66KK
60EL+66KK
63EL+66KK
TGGACCTTATTGAGCATCCGGATCC------------CAAGGAGAAGGATCTTA(¨12)
--------------------------ATCTATGATCCCAAGGAGAAGGATCTTA(¨181)
TGGACCTTATTGAGCATCCGGA---------------------GAAGGATCTTA(¨21)
TGGACCTTATTGAGCATCCGGATCC----TATGATCCCAAGGAGAAGGATCTTA(¨4)
--------------------------ATCTATGATCCCAAGGAGAAGGATCTTA(¨195)
-----------------------------TATGATCCCAAGGAGAAGGATCTTA(¨95)
--------------------------------GATCCCAAGGAGAAGGATCTTA(¨101)
TGGACCTTATTGAGCATCCGGATCC------------CAAGGAGAAGGATCTTA(¨12)
TGGACCTTATTGAGCATCCGGA---------------------------TCTTA(¨27)
-------------------------TATCTATGATCCCAAGGAGAAGGATCTTA(¨80)
TGGACCTTATTGAGCATCCGGATCC------------CAAGGAGAAGGATCTTA(¨12)
TGGACCTTATTGAGCATCCGGATCC------------CAAGGAGAAGGATCTTA(¨12)
TGGACCTTATTGAGC-----------------GATCCCAAGGAGAAGGATCTTA(¨17)
TGGACCTTATTGG--------------------------AGGAGAAGGATCTTA(¨26)
TGGACCTTATTGAGCATCC------------------CAAGGAGAAGGATCTTA(¨18)
----------------------------------------GGAGAAGGATCTTA(¨116)
TGGACCTTATTGAGCATCCGGATCCTATCCTATGATCCCAAGGAGAAGGATCTT(+1)
58EL+66KK TGGACCTTATTGAGCATCCGGATCCTATATCTATGATCCCAAGGAGAAGGATCT(+2)
TGGACCTTATTGAGCATCCGGATCCTATCTATCTATGATCCCAAGGAGAAGGAT(+4)
60EL+66KK TGGACCTTATTGAGCATCCGGATCCTATCCTATCTATGATCCCAAGGAGAAGGA(+5)
63EL+66KK TGGACCTTATTGAGCATCCGGATCCTATCCTATCTATGATCCCAAGGAGAGGGA(+1)
29
Figure 3.4: ZFNs induce heritable loss-of-function noggin alleles mutations. (A) Cel-1
digests of noggin amplicons from sibling heterozygous mutant and homozygous wild type F1
tadpoles produced from three mutant line founders. Bands migrating at 450 bp are full-length
noggin amplicons. Bands migrating at 300 bp and 150 bp are Cel-1 digest products. (B)
Sequence alignments of the targeted noggin locus from Cel-1-positive F1 mutants. Genomic and
translated sequences are shown for each mutant line. Asterisk indicates a stop codon. Red
nucleotides and amino acids indicate insertions and dashes represent deletions. (C) Schematic
of synthetic RNA injections into ventral vegetal blastomeres of 4-cell stage embryos to test
functionality of the induced mutant noggin alleles. (D) Quantification of secondary axis
induction following wild-type or mutant noggin RNA and scored for presence of ectopic dorsal
axes. Bars represent results of two (5 pg) or three (10 pg) independent experiments (±SD) White
bars show 5 pg RNA injections, black bars show 10 pg RNA injections. Two asterisks indicate
significantly different (p<0.01) from uninjected controls. (E-G’) Uninjected control embryos.
(H-J’) Embryos injected with 10 pg of 4 bp insertion mutant noggin and 200 pg LacZ RNA.
(K-M’) Embryos injected with 10 pg of 12 bp deletion mutant noggin and 200 pg LacZ RNA.
(N-P) Embryos injected with 10 pg of 3 bp insertion mutant noggin and 200 pg LacZ RNA. (QS) Embryos injected with 10 pg wild-type noggin and 200 pg LacZ RNA. (E,H,K,N,Q) Dorsal
view stage 19. (F,I,L,O,R) Dorsal view stage 28. (G,J,M,P,Q) Embryos stained with 12/101
antibody. (G.J) Lateral view. (G’,J’,M,P,Q) Dorsal view. Arrowheads show weak ectopic dorsal
axis induction.
30
31
Founder 1: 12bp Deletion
GATCCTATCTATGATCCCAAG
GATCC------------CAAG
EHPDPIYDPKEKDL
EHPD----PKEKDL
D
0
20
40
60
80
100
UC
5 pg
10 pg
4bp
Insertion
H
WT
Founder 3
**
3bp
Insertion
** **
WT
** **
Mutant or
Wild Type
Noggin RNA
Founder 2: 3bp Insertion
GGATC---CTATCTATGATC
GGATCTATCTATCTATGATC
EHPDP-IYDPKEKDL
EHPDLSIYDPKEKDL
12bp
Deletion
C
WT
Mut
WT
Mut
WT
H
H
WT
Founder 2
Founder 1
Founder 3: 4bp Insertion
WT CCGGATCC---TAT-CTATGA
Mut CCGGATCCCAAGATCCCATGA
WT EHPDPIYDPKEKDL
Mut EHPDKIP*
WT
Mut
WT
Mut
B
150bp
300bp
450bp
A
% Embryos with Ectopic Axis
UC
4bp Insertion
12bp deletion
3bp Insertion
WT
F
I
L
O
R
E
H
K
N
Q
S
P
M
J’
J
G’
G
Figure 3.5: Knockdown of Chordin and Follistatin results in a loss of dorsal structures in a
subset of embryos produced by heterozygous noggin mutant adults. (A-C) sox2 expression.
(D-E) myoD expression. (A,D) Uninjected control embryos. (B,C,E,F) Embryos injected with
20 ng Chordin morphino and 20 ng Follistatin morpholino. Dorsal views with anterior towards
the top.
32
Uninjected
Control
20 ng Chordin MO+
20 ng Follistatin MO
B
C
sox2
A
9/9
E
2/10
F
myoD
D
8/10
6/6
11/15
4/15
33
Figure 3.6: Stage series of representative embryos produced by heterozygous noggin
mutant adults. (A-J) Phenotypically wild-type embryos and tadpoles. (K-L) Wild-type
tadpoles. Total number of tadpoles and percentage within each phenotypic class. (M-N)
Abnormal tadpoles with dorsal “horn.” Total number of tadpoles and percentage within each
phenotypic class. (A,B,D,F,I,K,M) Dorsal views. (C,E,G,H,J,L,M) Lateral views. Anterior to
the left.
34
35
H
A
Stage 41
Stage 21/22
C
B
J
I
Stage 25/26
L
K
2533 (75.1%)
G
E
Stage 44
F
D
Stage 33/34
Stage 45
N
M
825 (24.9%)
Stage 39
Figure 3.7: Homozygous noggin mutant Xenopus tropicalis have severe lower jaw
deformities. (A-C) Wild-type stage 46 tadpoles. (D-F) Homozygous noggin mutant stage 46
tadpoles. Anterior towards the left. (A,D) lateral views. (B,E) dorsal views. (C,F) Ventral
views. (G-H) Flat-mounted cartilage from wild-type (G) and mutant (H) tadpoles. (G’-H’)
Schematic of skeletal elements in (G-H). (I) Survival curve of 100 wild-type (closed boxes) and
100 mutant tadpoles open triangles).
36
37
Wild Type
Mutant
Wild Type
Mutant
Infrarostral Cartilage
Ceratohyal
H’
H
Suprarostral Plate
Palatoquadrate
G’
E
D
G
B
A
Ceratobranchial
Meckel’s Cartilage
F
C
I
0
20
40
60
80
100
% Survival
8
9
11
12
13
14
15
Days post-fertilization
10
Wild type
Mutant
16
Figure 3.8: Expression of chondrogenic factors in wild-type and noggin mutant tadpoles.
(A-B) sox9 expression in the head of a representative wild-type (A) and mutant (B) tadpole at
stage 33. (C-F) col2a expression in the head of stage 33 (C-D) and stage 38 (E-F) tadpole.
(C,E) Representative wild-type tadpoles and (D,F) mutant tadpoles. Anterior to the left and
dorsal towards the top.
38
sox9
Wildtype
A
col2a
Mutant
B
Wildtype
Mutant
C
D
E
F
39
Figure 3.9: Expression of Bmp pathway targets in wild-type and noggin mutant tadpoles.
(A-D) Expression of bmp7 in the head of representative wild-type (A,C) and noggin mutant
(B,D) tadpoles. (E-H) Expression of msx2 in the head of representative wild-type (E,G) and
noggin mutant (F,H) tadpoles. (I-L) Expression of edn-1 in the head of representative wild-type
(I,K) and noggin mutant (J,L) tadpoles. (M-P) Expression of hand2 in the head of representative
wild-type (M,O) and noggin mutant (N,P) tadpoles. (A-B,E-F,I-J,M-N) Stage 33. (C-D,G-H,KL,O-P) Stage 38. Anterior to the left and dorsal towards the top.
40
bmp7
A
Wildtype
C
I
K
B
msx2
Mutant
D
Wildtype
edn1
J
L
E
Wildtype
G
Mutant
F
Mutant
H
Wildtype
hand2
M
N
O
P
Mutant
41
Table 3.1: $#noggin mutation induction and generation of germline mutants by
Zinc-Finger Nucleases in Xenopus tropicalis. Percent mosaic embryos in cohort calculated by
A52=2?02;A.42<3@6/96;42:/?F<@3<B;1A</2=<@6A6C23<?:BA.A6<;@.@12A20A21/F29
.@@.F@
Founder percent germline mutagenesis calculated by the percentage of heterozygous offspring
produced by founder in an outcross.
42
43
26EL+28KK
26EL+28KK
63EL+66KK
1
2
3
Founder (Males) Noggin ZFN Pair
116
568
300
Embryos injected
in cohort
100
60
16.1
% Mosaic
embryos in
cohort
2
17
4
Adults
raised in
cohort
24
12
16.7
Founder %
germline
mutagenesis
4bp Insertion
3bp Insertion
12bp Deletion
Mutation
Table 3.2: noggin ZFN recognition sequences. (;12?96;21@2>B2;02@6;160.A2-"/6;16;4
sites.
44
45
RSDNLSV RSANLTR RSDNLSV IRSTLRD TSGNLTR NRGNLVT
RSDALST ASSNRKT QSSDLSR TSANLSR RSDDLSE TNSNRKR
TTATTGAGCATCCGGATCctatcTATGATCCCAAGGAGAAG 25766KK
AATAACTCGTAGGCCTAGgatagATACTAGGGTTCCTCTTC 25763EL
RSDNLSV RSANLTR RSDNLSV IRSTLRD TSGNLTR NRGNLVT
RSDALST ASSNRKT QSSDLSR TSANLSR RSDTLSE TSANLSR
TTATTGAGCATCCGGATCctatcTATGATCCCAAGGAGAAG 25766KK
AATAACTCGTAGGCCTAGgatagATACTAGGGTTCCTCTTC 25760EL
RSDNLSV RSANLTR RSDNLSV IRSTLRD TSGNLTR NRGNLVT
RSDALST ASSNRKT QSSDLSR TSANLSR RSDTLSE TSANLSR
TTATTGAGCATCCGGATCctatcTATGATCCCAAGGAGAAG 25766KK
AATAACTCGTAGGCCTAGgatagATACTAGGGTTCCTCTTC 25758EL
TSSNLSR RSDNLSV RSANLTR RSDNLSV IRSTLRD
TSSNLSR RSDTLSE TSANLSR RSDYLST QNAHRKT
RSDNLSV RSANLTR RSDNLSV DNRDRIK QSSNLAR TSSNRKT
RSDALST ASSNRKT QSSDLSR TSANLSR RSDTLSE TSANLSR
25731KK
25730EL
ATCCGGATCCTATCTatgatCCCAAGGAGAAGGAT
TAGGCCTAGGATAGAtactaGGGTTCCTCTTCCTA
RSDNLSV RSANLTR RSDNLSV IRSTLRD TSSNLSR
QSSDLSR TSANLSR RSDTLSE TSANLSR RSDYLTK
Finger 1 Finger 2 Finger 3 Finger 4 Finger 5 Finger 6
TTATTGAGCATCCGGATCctatcTATGATCCCAAGGAGAAG 15026EL
AATAACTCGTAGGCCTAGgatagATACTAGGGTTCCTCTTC 15028KK
25729KK
25728EL
ZFN
AGCATCCGGATCCTAtctatGATCCCAAGGAGAAG
TCGTAGGCCTAGGATagataCTAGGGTTCCTCTTC
Noggin ZFN Binding Sequence (underlined)
CHAPTER 4
Expression Screen for Direct Targets of Wnt-Signaling in Neural Tissue
Introduction
2C29<=:2;A<3A52.:=56/6.;02;A?.9;2?C<B@@F@A2:6@A5<B45AA<[email protected]<?16;4A<A52
N.0A6C.A6<;A?.;@3<?:.A6<;O:<129'56@5F=<A52@6@Q?@A=?<=<@21/F"62BD8<<=
(NieuwkoopOthers, 1952b), postulates that neural tissue (by default anterior in nature) is induced
/FA52<?4.;6G2?Q?@A.;1A52;.116A6<;.9@64;.9@3?<:A52:2@<12?:=<@A2?6<?6G26AA<0?2.A2A52
3B99.;A2?6<?=<@A2?6<?$=.AA2?;<3A52;2B?.9=9.A2'526;1B0A6<;@A2=D.@Q?@A
:205.;[email protected];12?@A<<1D6A5A5216@0<C2?FA5.AA?2.A:2;A<320A<12?:.92E=9.;A@N.;6:.9
0.=@OD6A5A52:=.;A.4<;6@A"<446;6;1B02@;2B?.9A6@@B26;Xenopus (Lamb et al., 1993).
&B/@2>B2;A9F:<920B9.?05.?.0A2?6G.A6<;<3A52<?4.;6G2?6;.:=56/6.;@5.@@5<D;A5.AA52@2
0299@@20?2A2.;A.4<;6@A@<3A52:=.;1*;A=.A5D.F@2%</2?A6@.;1B?<1.. A role
for the Bmp antagonists Chordin &[email protected]
, Noggin (Lamb et al., 1993), and Follistatin
*69@<;.;12::.A6?6C.;9<B
in the induction step is well supported in Xenopus.
Reduction of these three antagonists results in a complete loss of neural tissue in Xenopus
5<85.2A.9<;@6@A2;AD6A5A52.0A6C.A6<;A?.;@3<?:.A6<;:<129;2B?.9A6@@B2
induced by Bmp antagonism will adopt an anterior fate in the absence of additional signals
2::.A6?6C.;9<B.;1!29A<;
.
'52./<C2Q;16;4@D2?23<99<D21D6A5A5216@0<C2?F<3@64;.96;[email protected];.0A6C.A21
in Xenopus,.?2@B3Q062;AA<=<@A2?6<?6G2A52;2B?.9=9.A2'52@26;09B12?2A6;<60.061B?@A<;2A
.9
<9:2A.9
%B6G69A./..;12@@299
&6C22A.9
*;A!0?2D2A
al., 1995) A<5.;1&<8<9
62082?.;1"625?@
and Fgf <E.;12::.A6
?6C.;9<B
92A052?2A.92;4.8B.;1#8.:<A<
.:/.;1.?9.;1
@64;.96;40A6C.A6<;<32.05<3A52@2=.A5D.F@6;16C61B.99F6;Xenopus was shown to
=<@A2?6<?6G2;2B?.9A6@@B2/F2E=.;16;4A522E=?2@@6<;1<:.6;@<3=<@A2?6<?;2B?.9:.?82?@.;1
D<?86;G2/?.Q@512:<;@A?.A212.05A</2;202@@.?F3<?A522E=?2@@6<;<3A52=<@A2?6<?;2B?.9
marker hoxB1 B1<52A.9.
!2:/2?@<3A52*;A3.:69F<3@64;.96;4964.;[email protected]/22;@5<D;A</2@A?<;4;2B?.9
=<@A2?6<?6G6;43.0A<?@;0.;<;60.9*;A@64;.96;4@20?2A21*;A/6;1@A<6A@?202=A<?@?6GG921
and LRP6 <4.;.;1"B@@2, leading to cytoplasmic enrichment and nuclear translocation
of β-catenin. Once inside the nucleus, β0.A2;6;0<:=92E2@D6A5' A<6;1B02A.?42A42;2
2E=?2@@6<;*;A.;1*;A.@64;.96;4D.@@5<D;A</2@B3Q062;AA<6;1B022E=?2@@6<;<3A52
hindbrain marker krox206;"<446;A?2.A21.;6:.90.=@.;1<C2?2E=?2@@6<;<3*;A.;A.4<;6@A@
0<B91@B==?2@@2E=?2@@6<;<3A52@242;2@6;D5<922:/?F<@6;160.A6;4A5.A*;A@64;.96;46@
?2>B6?213<?=<@A2?6<?;2B?.9=.AA2?;6;46;Xenopus 62082?.;1"625?@
. Furthermore, a
gradient of nuclear localized β0.A2;6;5645=<@A2?6<?9F.;19<D.;A2?6<?9FD.@</@2?C216;A52
neural plate of late gastrula/early neurula stage frog embryos 62082?.;1"625?@
,
92;16;4@B==<?AA<A525F=<A52@[email protected]*;A964.;[email protected];.:<?=5<42;4?.162;AA<=.AA2?;A52
46
;[email protected]?<:G2/?.Q@5@B442@[email protected]*;A3?<:A529.A2?.9:2@2;1<12?:
posteriorizes the neural plate ?A2?2A.9
<[email protected]@</22;@5<D;A5.A*;A.
from mesodem in the dorsal-lateral marginal zone of Xenopus embryos 6;1B02@2E=?2@@6<;<3
meis3,.42;2?2>B6?213<?56;1/?.6;12C29<=:2;A98<B/F2A.9
'56@5.@46C2;?6@2A<.
:<1293<?C2?A2/?.A2;2B?.9=.AA2?;6;4D52?2/F564592C29@<3*;A@64;.96;4=?<1B021/FA52
mesoderm, induces a more posterior neural fate such as spinal cord and intermediate doses
induce more medial fates like mid and hindbrain (Figure 4.1).
2@=6A2.D2.9A5<38;<D92142./<BAA52*;A=.A5D.FA52:205.;6@:/FD5605@64;.96;4
A5?<B45A56@=.A5D.F6;A52;2B?.9=9.A2?296./9F?2@B9A@6;A52?2@A?60A212E=?2@@6<;<3@=6;.90<?1
markers (e.g. hoxb9) posterior to hindbrain markers (e.g. krox20?2:.6;@B;092.?$?2C6<B@
D<?85.@12:<;@A?.A21A52=<@A2?6<?6G6;423320A<3*;A@64;.96;4F2A32DA?.;@0?6=A6<;.9A.?42A@
A5.A:216.A2A56@=?<02@@5.C2/22;612;A6Q21Here, I describe a screen that I performed to
16@0<C2?;<C29A?.;@0?6=A6<;.9A.?42A@<3*;A@64;.96;46;;2B?.9A6@@B2
Results and Discussion
Before assaying for Wnt-induced posterior neural gene expression, it is important to determine
the normal temporal and spatial expression of A-P neural markers at the onset of gastrulation
through neurulation. Therefore, Xenopus tropicalis embryos were fixed and stained for the
following factors: otx2, expressed in the forebrain and midbrain (Blitz and Cho, 1995), en2,
expressed at the midbrain-hindbrain boundary (Hemmati-Brivanlou et al., 1990; HemmatiBrivanlou et al., 1991), krox20, expressed in rhombomeres 3 and 5 (Bradley et al., 1993), and
hoxb9, a marker for the spinal cord (Wright et al., 1990). Consistent with the two-step model for
neural patterning, otx2 is expressed broadly throughout the presumptive neural plate at the onset
of gastrulation but becomes more restricted to the anterior with time (Figure 4.2 A-F). The more
posterior markers begin to be expressed at the onset of neurulation and the embryo exhibits a full
A-P pattern by the mid neurula stage (Figure 4.2 G-X). These results show that the neural plate
is fully patterned by stage 15 and therefore, an optimal stage to assay for Wnt targets in neural
tissue.
To allow temporal control over the activation of Wnt signaling, an inducible activator is required.
I used a fusion protein with the DNA binding domain of TCF fused to the transactivating domain
of VP-16 all in turn fused to a glucocorticoid receptor (TVGR) (Darken and Wilson, 2001).
Canonical Wnt signaling is sufficient to specify the dorsal axis during cleavage stages of
Xenopus embryos, and thus provides an excellent readout for Wnt activity (Smith and Harland,
1991; Sokol et al., 1991). To confirm the results of Darken and colleagues that TVGR efficiently
activates Wnt-signaling (Darken and Wilson, 2001), TVGR RNA was injected into the ventralvegetal blastomeres of 4-cell stage embryos and the resulting tadpoles were assayed for
secondary axis induction. Ventral-vegetal injection of TVGR alone or injection followed with
0.2% EtOH (vehicle) treatment resulted in normal development in 80% of injected tadpoles.
The remaining 20% showed only partial secondary axes (Figure 4.3A-C). Conversely,
dexamethasone (DEX) treatment of TVGR injected embryos resulted in 80% of injected tadpoles
with robust secondary axes induction as measured by the presence of eyes in the ectopic axis
47
(Figure 4.3A,D). The results from this experiment show that TVGR is a potent activator of Wnt
signaling upon DEX addition with minimal activation in the absence of the inducer.
In order to discover Wnt targets in neural tissue, ectodermal explants (animal caps) were induced
to become neural tissue via noggin expression followed by Wnt activation (Figure 4.4 A)
Therefore, the activity of TVGR in posteriorizing neural tissue was tested by assaying for known
A-P markers in animal caps under different treatments. As expected, uninjected animal caps
showed high expression of the epidermal marker epidermal keratin. Animal caps treated with
noggin alone expressed the anterior neural marker otx2 but not the epidermal marker epidermal
keratin or any posterior neural gene, demonstrating that this tissue adopted an anterior neural
fate. TVGR-injected animal caps that were neuralized by noggin showed robust expression of
the posterior neural markers krox20 and hoxb9 upon induction with DEX. While EtOH
treatment of TVGR-expressing animal caps express the hindbrain marker krox20, activation via
DEX is required to induce spinal cord fates as assayed by hoxb9 expression (Figure 4.4 B).
Consistent with these results, DEX treatment of neuralized animal caps expressing TVGR
induced the caps to undergo convergent-extension like morphogenesis that is consistent with
differentiation into spinal cord (Elul et al., 1997) (Figure 4.4 C). The observed morphogenesis
and neurulation are not due to mesodermal contamination of the ectodermal explants as there is
no expression of the mesodermal marker muscle actin.
Next, I validated the use of TVGR in neuralized animal caps to find direct transcriptional targets
of Wnt signaling. Detection of direct transcriptional targets requires inhibition of protein
synthesis to prevent activation of secondary targets following translation of Wnt-induced
transcripts. Cycloheximide (CHX) is a potent inhibitor of protein translation (Obrig et al., 1971)
and provides a convenient tool to prevent the expression of secondary targets upon activation of
a signaling pathway. To determine the dose of CHX I used was effective, I pretreated neuralized
animal caps that were injected with TVGR with CHX, then induced with DEX, and assayed for
known direct and indirect transcriptional targets of Wnt (Figure 4.5 A). CHX treatment prior to
activation of TVGR did not prevent activation of the direct transcriptional target meis3 (Elkouby
et al., 2010) but blocked expression of the indirect target hoxb9 (Domingos et al., 2001) (Figure
4.5 B). These results demonstrated that the conditions I used were sufficient to induce neural
tissue, posteriorize it via Wnt activation and finally, enrich for direct targets.
Next, total RNA from animal caps treated with noggin alone (anterior neural sample), neuralized
caps with activated TVGR (posteriorized neural sample) and neuralized animal caps treated with
CHX prior to TVGR activation via DEX addition (direct target sample) was harvested. The
RNA from these samples were used to construct RNA-seq libraries which were then 76-basepair
single-end sequenced. The resulting sequence reads were mapped to a collection of 10,088 nonredundant, full-length Xenopus laevis cDNA sequences (http://xgc.nci.nih.gov). Data analysis
was preformed by first aligning the reads using the programs TOPHAT(Trapnell et al., 2009) and
BOWTIE (Langmead et al., 2009) to map the reads to cDNA sequences and then using
CUFFDIFF (Trapnell et al., 2010) to determine differences in read quantities between samples.
This analysis identified 228 genes greater than 2-fold increased in the direct target sample when
48
compared to the anterior neural sample (Appendix 1). Enrichment of the direct target meis3
(Elkouby et al., 2010) as well as cdx2 (Wang and Shashikant, 2007) in the direct target sample
reads provided confirmation that this screen and analysis were successful in identifying Wnt
target genes and suggests that candidates are likely to be direct targets.
While I found over 200 genes with enriched expression when directly activated by Wnt, this
method of analysis was limited because the reads were mapped to a cDNA library in lieu of a full
genome. Any genes absent from this library would have been missed and it is likely that there
are more potential Wnt targets to be identified. With the recent sequencing of Xenopus laevis, it
is now possible to repeat the bioinformatic analysis using the genome and potentially reveal
more candidate target genes. Furthermore, comparing the results from the screen described here
to those obtained from β-catenin chromatin immunoprecipitation (ChIP) will provide a
complementary approach for detection of direct Wnt targets. ChIP-Seq will also identify the
enhancers and promoters that mediate Wnt regulation of the identified target genes allowing for
insight into the mechanism of gene regulation via Wnt signaling.
This screen aimed to identify novel factors that mediate Wnt signaling to posteriorize the neural
plate, therefore I prioritized the candidate genes that are likely to function in patterning by first
assaying expression of transcription factors, RNA binding proteins, and various signaling
proteins. These genes need to be expressed at the right developmental timepoint and relevant
tissue to play a role in Wnt mediated neural patterning. Accordingly, I assayed expression of
selected candidates via in situ hybridization in Xenopus tropicalis embryos during gastrulation
and neurulation to identify those that are expressed in the posterior neurectoderm. In addition to
the known neural Wnt targets identified in the screen, several of the selected transcription factors
had detectable expression in neurula stages. The transcription factors ahctf1 (Figure 4.6, A-C),
foxi4.2 (Figure 4.6, D-F), and churchill (Figure 4.6 G-I) were barely expressed over background
staining but show some weak staining in the posterior. zmiz1 is first detected in the posterior
neural plate at the onset of neurulation and becomes restricted to two posterior stripes in the
spinal cord anlage by stage 15 (Figure 4.6, J-L). In mice, ahctf1, the gene responsible for the
Elys mutation, and zmiz1 are expressed in the developing spinal cord (Beliakoff et al., 2008;
Kimura et al., 2002). However their roles in A-P patterning have not been established. max
(Figure 4.6 M-O), originally reported to be ubiquitously expressed (Newman and Krieg, 1999),
and znf384 are expressed in the presumptive neural plate at gastrulation and show enrichment in
the posterior neural region at neurulae stages (Figure 4.6 P-R). The Spalt-like genes sall1
(Figure 4.6 S-U) and sall4 (Figure 4.6 V-X) are expressed at gastrulation in the presumptive
neural plate and show robust expression in the posterior neural plate in later stages. The role of
the sall genes in human diseases (Kohlhase et al., 2002; Kohlhase et al., 1998), stem cell
maintenance (Zhang et al., 2006), and limb development (Neff et al., 2005) have been studied
but a neural patterning role has not been described. Unexpectedly, sox11 was identified in the
screen and is expressed in the neural plate but is absent from the most posterior region. This
could be the result of secondary repression wherein the function of a different factor that
normally represses sox11 expression in this region is relieved when the tissue was treated with
CHX (Figure 4.6 Y-Aa).
49
The splicing factor fus was shown to be crucial for gastrulation via regulation of multiple
developmental pathways (Dichmann and Harland, 2012). It is possible that RNA-binding
proteins may play a previously unappreciated role in A-P patterning. Both heterologous nuclear
ribonuclear proteins hnRNPk (Figure 4.7 A-C) and hnRNP H3 (Figure 4.7 D-F) are expressed in
the posterior neural plate. The genes encoding splicing factors sf3b4 (Figure 4.7 G-I) and sap130
(Figure 4.7 J-L) show specific expression dorsally around the blastopore at stage 12 which
becomes more enriched at the midneurula stage. These factors are known to interact with each
other (Kotake et al., 2007; Menon et al., 2008) and sf3b4 has been implicated in bone
development (Watanabe et al., 2007), a process that is also regulated by Wnt signaling (reviewed
in (Williams and Insogna, 2009). Two additional genes encoding splicing factors sfrs7 and sfrs6
were also identified in this screen (Figure 4.7 M-R). sfrs7 is specifically expressed in posterior
regions and sfrs6 also shows a posterior neural expression domain at stage 12. However the
staining is faint and does not show specific expression at stage 15. A role for RNA-binding
proteins and splicing factors in mediating neural A-P patterning has not been described or
implicated. The neural expression of identified RNA binding factors is strong evidence that
these proteins are important in neural patterning. It will be interesting to determine if the
splicing factors identified in this screen comprise a core of Wnt-regulated splicing proteins that
function in cell fate specification.
In addition to transcription and RNA binding factors, the planar cell polarity gene prickle1
(Wallingford et al., 2002) is expressed in the midline and enriched in the posterior (Figure 4.8 AC). Prickle homologs have been shown to mediate movements during gastrulation, but
morpholino studies in Xenopus and zebrafish have not revealed a role in neural A-P patterning
(Takeuchi et al., 2003). It remains to be tested if there is functional redundancy between the
prickle genes in neural patterning or if there simply is not a role for Prickle in this process.
Finally, the adhesion molecule Lmo7 (Ooshio et al., 2004)shows punctate expression in the
neural plate, although no function in neural development for this gene has been established
previously (Figure 4.8 D-F).
Many of the candidates identified in the screen are enriched in the posterior neural region
demonstrating the effectiveness of this method to identify novel targets of Wnt signaling.
However, not all of the candidates identified in the screen were expressed in the neural plate or
were detectable via in situ hybridization. One explanation could be that the expression of these
genes is below the threshold of detection by this method. This does not explain however, why
some genes found to be highly expressed in the RNA-seq data were not detected via in situ
hybridization. These could be false positives that result from the nature of the analysis. The
software packages used to align and quantitate reads also find splice junctions which could lead
to spurious results when reads are aligned to an index of spliced cDNAs as was done for this
screen. An alternative explanation is that the activation of Wnt signaling in the animal caps is
not physiological and results in activation of targets that are not normally expressed during early
or mid-neurula stages. However, this is likely to be a minority of the data set as 17 of 22
previously unknown posterior neural genes showed specific posterior expression. Another
50
possibility is that some of the identified genes are subject to repression via secondary targets.
Therefore, these genes would not be detected in wild-type embryos but would be highly
expressed in conditions where Wnt was activated in the presence of CHX.
While I prioritized transcription factors and RNA-binding proteins, there are other important
classes of genes identified in the screen that could play crucial roles in meditating A-P neural
patterning. For example, molecules involved in signal transduction and chromatin remodeling
likely have significant effects in Wnt-signaling and downstream gene expression. Further
analysis of candidates identified in this screen will provide a more complete understanding of
Wnt-mediated neural patterning.
51
Figure 4.1: Model of Wnt-induced patterning of the neural anterior-posterior axis.
Schematic of a proposed Wnt gradient and anterior-posterior patterning of the neural plate.
52
53
Midbrain
Hindbrain
Head Mesoderm
Source of BMP and
Wnt antagonists
2. Neural tissue is then posteriorized by a
gradient of FGF, RA, and Wnt.
1. BMP antagonists induce neural tissue from
ectodermal precursors.
Forebrain
Notochord
Source of BMP antagonists
Source: paraxial mesoderm
Wnt
Spinal Cord
Figure 4.2: Temporal expression of anterior posterior neural markers in Xenopus tropicalis.
In situ hybridization for otx2 (A-F), engrailed2 (en2) (G-L), krox20 (M-L), and hoxb9 (S-X).
Dorsal views with anterior towards the top.
54
55
otx2
en2
krox20
hoxb9
N
M
T
H
G
S
B
A
10.5
11
U
O
I
C
12
V
P
J
D
NF Stage
13
W
Q
K
E
14
X
R
L
F
15
Figure 4.3: TVGR activates canonical Wnt signaling. (A) Quantification of secondary axis
induction by ventral-vegetal injection of TVGR at the 4-cell stage. (B) Uninjected control
tadpole. (C) TVGR injected tadpole. (D) TVGR injected tadpole treated with 0.4% EtOH. (E)
TVGR injected tadpole treated with 10 µM DEX, red staining shows β-galactosidase (tracer).
56
57
0%
20%
40%
60%
80%
0.2% Ǎ0 0.4% Ǎ0
EtOH DEX EtOH DEX
ǒJ79*551$SJ/DF=51$
U.C.
Complete 2º Axis
Partial 2º Axis
Normal
TVGR/Dex Induction of Wnt Signaling
100%
A
SJ79*5Ǎ0'(;
E
SJ79*5(W2+
D
SJ79*5
C
U.C.
B
&RPSOHWHƒ$[LV
3DUWLDOƒ$[LV
1RUPDO
1RUPDO
Figure 4.4: TVGR efficiently posteriorizes neuralized ectodermal explants. (A) Strategy to
use TVGR to induce Wnt signaling in animal caps. (B) RT-PCR on 5 embryos or 25 animal caps
treated with the indicated reagents. -RT: reaction done in the absence of Reverse Transcriptase,
epi. ker: epidermal keratin (epidermis), mus. act.: muscle actin (mesoderm). When indicated the
following doses were used: noggin 10 pg, TVGR 4 pg, Dexamethasone: 10 µM (C) Animal caps
treated with the indicated reagents.
58
59
eef1a1
mus. act.
epi. ker.
hoxb9
krox20
en2
otx2
noggin
B
-RT
A
2-cell
Inject
noggin+
TVGR
isolate
RNA at
stage 15
equivalent
10 uM
DEX
noggin/TVGR+EtOH
UC
C
stage 9
Excise
animal
caps
noggin/TVGR+DEX
noggin
Figure 4.5: Cycloheximide treatment prior to TVGR induction enriches for direct Wnt
targets in neuralized ectodermal explants. (A) Strategy to use animal caps to screen for direct
transcriptional targets of Wnt signaling in neural tissue. (B) Semi-quantitative qPCR on either 5
whole embryos or 25 animal caps treated with the indicated reagents and used for RNA-seq
library synthesis. meis3 and hoxb9 serve as controls for direct and indirect targets of Wnt
signaling, respectively. When indicated, the following doses were used: noggin 10 pg, TVGR 4
pg, Dexamethasone 10 µM.
60
A
Inject
noggin+
TVGR
Excise
animal
caps
2-cell
5 uM
CHX
1.5hrs
10 uM
DEX
isolate
RNA at
stage 15
equivalent
stage 9
B
Animal Caps
Whole Embryo
noggin
-
+
+
-
+
+
+
TVGR
-
-
+
-
-
+
+
EtOH
-
-
-
-
-
+
-
Dex
-
-
-
-
-
-
+
-RT
hoxb9
meis3
eef1a1
5 µM Cycloheximide
hoxb9
61
Figure 4.6: Expression patterns of transcription factors identified in the screen for direct
Wnt targets. In situ hybridization of selected genes in Xenopus tropicalis. Dorsal-vegetal
views of stage 10.5 embryos, dorsal views of stage 12 and 15 with anterior toward the top. NF
stage: Nieuwkoop Faber stage (Nieuwkoop, 1967).
62
NF stage
12
15
B
C
G
H
I
M
N
O
S
T
U
Y
X
Aa
12
15
D
E
F
J
K
L
P
Q
R
V
W
X
sox11
sall4
sall1
max
znf384
zmiz1
churchill
ahctf1
A
10.5
foxi4.2
10.5
63
Figure 4.7: Expression patterns of RNA-binding factors identified in the screen for direct
Wnt targets. In situ hybridization selected genes in Xenopus tropicalis. Dorsal-vegetal views
of stage 10.5 embryos, dorsal views of stage 12 and 15 with anterior toward the top. NF stage:
Nieuwkoop Faber stage (Nieuwkoop, 1967).
64
NF stage
12
15
B
C
G
H
I
M
N
O
15
D
E
F
J
K
L
P
Q
R
sfrs6
sf3b4
sfrs7
12
sap130
hnRNPk
A
10.5
hnRNP H3
10.5
65
Figure 4.8: Expression patterns of identified Wnt targets prickle1 and lmo7. In situ
hybridization in Xenopus tropicalis for indicated genes. Dorsal-vegetal views of stage 10.5
embryos, dorsal views of stage 12 and 15 with anterior toward the top. NF stage: Nieuwkoop
Faber stage (Nieuwkoop, 1967).
66
NF stage
12
A
B
D
E
15
C
F
lmo7
prickle1
10.5
67
CHAPTER 5
Spalt-like 4 mediates Wnt-induced neural patterning via repression of pouV/
Oct4 family members.
Introduction
Pieter "62BD8<<=.;1F.969.16@[email protected]<=:2;A<3A52.:=56/6.;02;A?.9;2?C<B@
@F@A2:.?6@[email protected]<;.;1A?.;@3<?:.A6<;O"62BD8<<=
"62BD8<<=#A52?@
/
NieuwkoopOthers, 1952a). This hypothesis states that neural tissue is induced as an anterior
state by the organizer, and then posteriorized by additional signals from the mesoderm to create
A523B99.;A2?6<?=<@A2?6<?$=.AA2?;<3A52;2B?.9=9.A20A6C.A6<;<?;2B?.96;1B0A6<;
?2>B6?2@<;2:<?=5<42;60=?<A26;:=.;A.4<;6@A@@B05.@"<446; .:/2A.9
5<?16;&[email protected]
.;1<[email protected];2::.A6?6C.;9<B2A.9
3?<:A52<?4.;6G2?
A<6;1B02A521<[email protected]<12?:A<.1<=A.;2B?.93.A25<85.2A.9;1221.;F
manipulation that blocks Bmp signaling in ectoderm results in cells adopting anterior neural fates
2::.A6?6C.;9<B.;1!29A<;
.B1.96G.A6<;<?A?.;@3<?:.A6<;<3A52;2B?.9=9.A2
<00B?@C6.@64;.96;4/F?2A6;<60.061%9B:/2?42A.9
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<9:2A
.9
%B6G69A./..;12@@299
&6C22A.9
43<E.;12::.A6?6C.;9<B
92A052?2A.92;4.8B.;1#8.:<A<
.:/.;1.?9.;1
%6/[email protected]
.;1*;Aβ0.A2;6;<:6;4<@2A.9
?A2?2A.9
A<5.;1&<8<9
62082?.;1"625?@
!0?2D2A.9
2@=6A2A52612;A6Q0.A6<;<3A52@2@20?2A21
factors as mediators of anterior-posterior (A-P) neural patterning, the mechanism by which
A?.;@1B0A6<;<3A52@2@64;.9@?2@B9A6;16332?2;A92C29@<3 gene expression and discrete neural fates
remains poorly understood.
6C2;A526;A2?2@A6;A52?<92<3*;A@64;.96;46;.E6.9=.AA2?;6;4@B?=?6@6;49F32DA?.;@0?6=A6<;
3.0A<[email protected]:216.A2$16332?2;A6.A6<;5.C2/22;612;A6Q21'525<:2</<E42;2gbx2 is a
16?20AA.?42A<30.;<;60.9*;A@64;.96;4.;1=?6:.?69F@2?C2@A<6;1B02;2B?.90?2@A 62A.9
'5242;22;0<16;4A52' 3.:69F5<:2</<EA?.;@0?6=A6<;3.0A<?meis3,?2>B6?213<?
56;1/?.6;.;1;2B?<;.916332?2;A6.A6<;6@16?20A9F.0A6C.A21/F*;A.3?<:A521<[email protected]?.9
:.?46;.9G<;298<B/F2A.9
98<B/F2A.9
'520.B1.95<:<9<4@cdx1 and cdx4
.?216?20A*;AA.?42A@6;:<B@2$69<;2A.9$69<;2A.9$?6;<@2A.9
.;1
5.C2<C2?9.==6;4?<92@6;=<@A2?6<?12C29<=:2;A<3A52A5?2242?:9.F2?@..@.;[email protected]@
@[email protected]
C.;12)2;2A.9
;A52;2B?.9=9.A2<3Xenopus, simultaneous
8;<081<D;<31E
.;16@?2>B6?21A</9<08.1<=A6<;<3A52:<@A=<@A2?6<?;2B?.93.A2@
..@.;[email protected]@
'52&=.9A9682&.99=?<A26;@.?2C2?A2/?.A25<:<9<4@<3A52Drosophila=?<A26;&=.9A&=.9A6@
?24B9.A21/F20.=2;A.=92460@64;.96;[email protected]
.;1:BA.A6<;@6;spalt result in
5<:2<A60A?.;@3<?:.A6<;@I?42;@
'52?2.?23<B?:2:/2?@<3A52&=.9A9682&.99
3.:69F<3G6;0Q;42?A?.;@0?6=A6<;3.0A<?@6;C2?A2/?.A2@12296@.;1.??6<99
:2:/2?@0<;A.6;.;"A2?:6;.9G6;0Q;42?1<:.6;3<99<D21/FC.?6./92;B:/2?@<3
68
1<B/92A.;1A?6=92AG6;0Q;42?1<:.6;@&D22A:.;.;1!I;@A2?/2?4&.99
.;1
0.;3B;0A6<;.@26A52?A?.;@0?6=A6<;.9?2=?2@@<?@ .B/2?A5.;1%.B05:.; .B/2?A52A.9
B2A.9,.;42A.9
<?.0A6C.A<?@6232?2A.9-5.;42A.9
6:2A.9,.;42A.9B:.;:BA.A6<;@6;sall1 and sall4 cause the autosomal
1<:6;.;A'<D;2@?<08@.;1#8656?<@F;1?<:2@?2@=20A6C29FD5605.?2/<A505.?.0A2?6G21/F
96:/.;10<4;6A6C212320A@<[email protected]
<[email protected] is the best
studied of the spalt homologs because of its role in maintaining and inducing pluripotency. sall4
8;<08<BA:602162.A=?26:=9.;A.A6<;@A.42@1B2A<.3.69B?2<3A526;;2?0299:.@@!A<
=?<9632?.A2.;1:.6;A.6;=9B?6=<A2;0F&.8.86,B:<A<2A.9sall4 null embryos fail to
2E=?2@@A52@A2:02993.0A<?pou5f1#0A6;A52!?2@B9A6;46;6;0?2.@21cdx22E=?2@@6<;.;1
A?<=520A<12?:2E=.;@6<;.AA522E=2;@2<32:/?F<;60A6@@B2*B2A.9-5.;42A.9
B?A52?:<?28;<081<D;<3sall46;56/6A@6;1B021=9B?6=<A2;A@A2:6$&029942;2?.A6<;
3?<:Q/?</9.@A@<C2?2E=?2@@6;4oct3/4, sox2, and klf4; .;12E=?2@@6<;<3sall4 along with oct3/4,
sox2 and klf4?2@B9A@6;56452?23Q062;0F<36$&029942;2?.A6<;3?<:5B:.;12?6C21Q/?</9.@A@
'@B/<<8.2A.9
9<;6;4.;12E=?2@@6<;.;.9F@[email protected]?2C2.921sall2, sall3, and sall4A</22E=?2@@211B?6;4
early Xenopus 2:/?F<42;2@6@<992:.;;2A.9
#;.62A.9#;B:.2A.9
.;1D6A5A522E02=A6<;<3sall2#;.62A.92E=?2@@216;=<@A2?6<?;2B?.9?246<;@ <@@
of-function alleles of sall1, sall2 and sall4 result in mouse embryos with neural tube closure
12320A@H5:2A.9?2C2.96;4.?<923<?sall genes in neural differentiation and/or
:<?=5<42;2@6@2@=6A2A526?2E=?2@@6<;6;=<@A2?6<?;2B?.9?246<;@<3C2?A2/?.A22:/?F<@.
role for the sall genes in caudalization has not been elucidated.
Here, I show that sall46@?2>B6?213<?0.B1.96G.A6<;.;16:=<?A.;A9F@=6;.90<?116332?2;A6.A6<;
<3;2B?.9A6@@B2&=206Q0.99F@5<DA5.Asall4 represses the stem cell factor pouV/Oct4 in order
to release cells from an undifferentiated state.
Results
In the previous chapter, I described a screen to identify direct targets of Wnt in neural tissue.
That screen identified sall1 and sall4 as candidate Wnt targets that are involved in neural
posteriorization. Since the sall genes were identified in a high-throughput screen, it was
important to confirm that they were not false positives. To that end, I confirmed the results of
my screen via the animal cap assay described in Figure 4.5 and quantitated transcript abundance
with qPCR. Incubation with CHX prior to activation of TVGR in neuralized caps resulted in
increased cdx2 (Figure 5.6A), sall1 (Figure 5.4A), sall4, and meis3, but not hoxb9, expression
(Figure 5.1A). The activation of the sall genes by canonical Wnt activation in the absence of
protein translation strongly suggests that they are direct targets of Wnt signaling in neural tissue.
As a complementary approach to using CHX-treated animal caps, I used chromatin
immunoprecipitation (ChIP) followed by qPCR to examine whether β-catenin is bound to the
sall4 genomic locus. β-catenin forms an activation complex with TCF/LEF in response to Wnt
signaling to activate target genes (Behrens et al., 1996). Due to a lack of effective commercial βcatenin antibodies, I overexpressed a C-terminal FLAG-tagged version of X. laevis β-catenin.
69
Expression of the FLAG epitope was confirmed by immunoblotting (Figure 5.2A). The activity,
dosage and specificity of FLAG-tagged β-catenin were tested by rescuing the β-catenin
knockdown phenotype. β-catenin morphants lack dorsal structures, resulting in ventralized
“belly pieces” (Heasman et al., 2000). Injection of β-catenin morpholino resulted in over 60% of
embryos lacking dorsal tissue. Co-injection of FLAG-tagged β-catenin RNA restored dorsal
structures, demonstrating both the specificity and activity of this construct (Figure 5.2B).
Finally, I assayed dorsalization resulting from overexpression of FLAG-tagged β-catenin without
morpholino injection. Consistent with Yost and colleagues (Yost et al., 1996), injection of 500
pg of RNA encoding tagged β-catenin did not significantly alter dorsal structures as measured by
the dorsoanterior index (Kao and Elinson, 1985) (Figure 5.1B). I assume that the injected RNA
results in β-catenin that enters the normal pool of unstable protein, and whose amount is subject
to regulation by the normal signaling pathway (Yost et al., 1996).
The sall4 locus in Xenopus laevis contains four exons and three introns (Figure 5.1C). Scanning
for consensus binding sequences within 3 kb of flanking sequence of the sall4 locus revealed six
putative TCF/LEF binding sites (Elkouby et al., 2010; McKendry et al., 1997) within the first
intron. Three of these sites are tightly clustered within a 150 bp span at positions +2347, +2387,
and +2456 (relative to the predicted transcription start site) and are conserved in Xenopus
tropicalis. The conservation and clustering of these sites suggested that this region could be
regulated by β-catenin. Using FLAG antibodies for ChIP, this region was found to be
significantly enriched compared to a negative control (xmlc2) region (Figure 5.1D). A –2.7 Kb
region upstream of meis3 was used as a positive control for β-catenin binding (Elkouby et al.,
2010). Anti-FLAG immunoprecipitations in uninjected control embryos resulted in negligible
enrichment of any loci assayed. Taken together, activation via TVGR in the presence of CHX
along with β-catenin binding to TCF/LEF sites in the first intron provide strong evidence that
sall4 is a direct transcriptional target of Wnt signaling in the neurectoderm.
During gastrulation, sall4 is initially expressed in a broad domain throughout the marginal zone
and the animal pole (Figure 5.3A). At stage 10, sall4 expression is restricted to the sensorial
neurectodermal cells (cells that will give rise to the central nervous system) in animal dorsal
regions (Fig 3E). At the onset of neurulation, sall4 is strongly expressed throughout the sensorial
neurectodermal layer of cells in the neural plate (Fig 3B,F,G). Neural expression of sall4 in
stage 15 (mid-neurula) embryos is in the hindbrain and spinal cord anlage (Figure 5.3C,H,I).
Later stage neurulae (stage 18) strongly express sall4 in the posterior neural tube, hindbrain,
developing placodes, and epidermis (Figure 5.3D,J,K). The posterior neural expression of sall4
and regulation by β-catenin and TCF/LEF make it a candidate for mediating Wnt-induced
posteriorization of the neural plate.
sall1 is expressed in the dorsal ectoderm and involuting mesoderm during gastrulation (Figure
5.4 B,B’). Expression becomes restricted to the notochord and circumblastoporal collar at the
early neurula stage (Figure 5.4C-C’’). Similar to sall4, sall1 is expressed in the spinal cord
anlage at mid and late neurula stages (Figure 5.4D-D’’,E-E’’).
70
Given the neural expression of sall4 and its identification as a Wnt target in neuralized animal
caps, I hypothesized that loss of Sall4 would affect neural patterning. To test this, I used
morpholinos targeting sall4 and assayed for neural gene expression. Morphant embryos had
neural tube closure defects and began to disintegrate and die at mid-tailbud stages. The neural
tube closure defect is consistent with defects in neural patterning, so I assayed several markers of
neural differentiation. The pan-neural marker sox2 was expressed in the neural plate in
uninjected and Sall4 morphants, demonstrating that the dorsal ectoderm of morphants still
retained a neural identity (Figure 5.5A,B). Conversely, the expression of n-tub, a marker for
differentiating neurons, was markedly reduced but still present in morphants, suggesting that
Sall4 is required for the second wave of neurogenesis in the tailbud tadpole (Figure 5.5C,D).
Another marker for early motor neuron differentiation, nkx6.1, was expressed in the central
nervous system of morphants. Neural crest cells were present as determined by the expression of
snai2 (Figure 5.5E,F). Though present, these markers were expressed in a pattern more similar
to early neurulae, suggesting either a delay or failure of terminal differentiation. Sall4 morphants
expressed the dorsal mesoderm marker myoD in a similar pattern to uninjected control embryos,
therefore, neural defects were not due to a major loss of paraxial mesoderm (Fig. G,H).
It is well understood that Wnt is a caudalizing factor in neural patterning, therefore if Sall4
mediates this process I predicted that Sall4 morphants would lose posterior neural identity. To
test for this, Sall4 morpholinos were injected into one animal dorsal cell of 4-cell stage embryos
to allow for comparison between injected and uninjected sides. Strikingly, the injected side of
embryos showed a significant posterior shift in the expression of otx2 and krox20 relative to the
control side (Figure 5.5I,J,Q). This shift suggested that knockdown of Sall4 results in an
expansion of anterior neural identity at the expense of posterior neural differentiation, consistent
with a Wnt loss-of-function phenotype. Accordingly, the injected side had a significant reduction
in the expression domain of the spinal cord marker hoxb9 (Figure 5.5K,L). Furthermore, the
expression of two other markers for spinal cord differentiation, hoxc10 (Figure 5.5M,N) and
hoxd10 (Figure 5.5O,P) was significantly reduced following Sall4 knockdown (Figure 5.5R,S,T).
However, Sall4 knockdown does not equally reduce cdx2 expression, another identified direct
Wnt target (Figure 5.6B,C). These results demonstrate that Sall4 is required for specification of
posterior neural tissue.
The failure of Sall4 morphants to induce posterior fates in neural tissue suggested that the caudal
tissue remained in an undifferentiated stem-like state. Sall4 positively regulates the stem cell
factor Oct4/pou5f1 locus in the inner cell mass of mouse embryos to maintain pluripotency
(Zhang et al., 2006). Since the posterior neurectoderm failed to differentiate in Sall4 morphants,
I hypothesized that Oct4 prevents neural differentiation in the morphants because Sall4 was
negatively regulating pou5f1 in the neural tissue of Xenopus. There are three class V Poudomain proteins in Xenopus with similar sequence and syntenic organization to mammalian
pou5f1/Oct4: pou25, pou60, and pou91, henceforth referred to as pouV/Oct4 (Morrison and
Brickman, 2006).
71
If Sall4 negatively regulates pouV/Oct4, Sall4 morphants should have increased expression of
these factors. Indeed, knockdown of Sall4 in unilateral and bilateral injections resulted in
ectopic expression of pou25 (Figure 5.7A,B,C), pou60 (Figure 5.7D,E,F), and pou91 (Figure
5.7G,H,I). Accordingly, the increase in expression of pou25 and pou91 was the greatest in the
neural tube where sall4 is normally expressed. pouV/Oct4 expression in Sall4 morphants relative
to control embryos was quantified via qPCR and displayed a significant increase in all three of
the pouV/Oct4 genes (Figure 5.7O,P,Q). Co-injection of Xenopus tropicalis sall4 RNA that is
not targeted by the Sall4 morpholino resulted in a partial rescue of the pou25 expression level
and restored pou60 and pou91 expression to wild-type levels (Figure 5.7O,P,Q). These
experiments demonstrate that loss of Sall4 relieves repression of the pouV/Oct4 homologs,
resulting in an overexpression of pouV/Oct4 stem cell factors in the neural plate.
A second, non-overlapping Sall4 morpholino (MO2) was used as an additional control for
specificity. If the above results are indeed due to Sall4 knockdown, then this second morpholino
should result in the same phenotype. As seen with the first Sall4 morpholino, I found that
injection of the Sall4 MO2 resulted in a similar loss of hoxb9 (Figure 5.8A,B) and hoxd10
(Figure5.8C,D) expression as well as a posterior shift in oxt2 and krox20 expression (Figure
5.8A,B). The Sall4 MO2 also resulting in ectopic expression of pou91 (Figure 5.8E,F). These
results demonstrate that the morpholinos targeting Sall4 are specific and effective.
Next, I asked whether ectopic pouV/Oct4 expression is sufficient to block posterior neural
differentiation by injecting RNA for the three pouV/Oct4 genes unilaterally into embryos and
assaying A-P neural gene expression. Neural plate cells expressing ectopic pouV/Oct4 as traced
by the co-injected marker β-galactosidase did not express hoxb9, whereas cells on the uninjected
side expressed this spinal cord marker (Figure 5.7J,K,L,M,N). The increase in pouV/Oct4
expression in Sall4 morphants, together with the result that ectopic expression of these genes
inhibited posterior neural differentiation, provides evidence that the loss of caudal identity in
morphants is due to pouV/Oct4 overexpression.
The observed pouV/Oct4 increase following knockdown of Sall4 suggested a mechanism for the
loss of posterior neural identity whereby the ectopic pouV/Oct4 expression prevents
differentiation of neural tissue into spinal cord. I therefore reasoned that depleting pouV/Oct4 in
Sall4 morphants would restore posterior neural identity. To that end, morpholinos targeting the
three pouV/Oct4 homologs(Morrison and Brickman, 2006) were co-injected with Sall4
morpholinos. Consistent with results described above, knockdown of Sall4 resulted in a loss of
expression of the spinal cord markers hoxb9 (Figure 5.9A,E), hoxc10 (Figure 5.9B,F), and
hoxd10 (Figure 5.9C,G) but not in a loss of the pan-neural marker sox2 expression (Figure
5.9D,H). Co-injection of the pouV/Oct4 morpholinos with Sall4 morpholinos restored posterior
neural gene expression lost by Sall4 knockdown alone (Figure 5.9I,J,K). Though reduced, sox2
was expressed in the neural plate of Sall4-pouV/Oct4 morphants (Figure 5.9L). Finally,
comparing the length of the hox gene expression domains in Sall4 and Sall4-pouV/Oct4
morphants revealed a significant rescue of all three spinal cord markers in the quadruple
morphants (Figure 5.9N). pouV/Oct4 knockdown resulted in anterior defects, evidenced by
72
mispatterning of the otx2 and krox20 expression domains and a lateral expansion of hoxb9,
consistent with a role for the Oct4 homologs in repressing posterior neural identity (Figure
5.9M).
My results demonstrated that Wnt-signaling induces Sall4 expression, which represses pouV/
Oct4 allowing posterior neural differentiation. Yet posterior patterning of the neural plate also
occurs in response to Fgf and RA signaling. If Sall4 were required to repress pouV/Oct4 in order
to allow for caudal differentiation, then depletion of Sall4 should block posteriorization by Fgf
and RA. Therefore, I tested whether repression of pouV/Oct4 via Sall4 is required for both Fgfand RA-induced caudalization.
To address the question of whether Fgf-induced caudalization requires Sall4, uninjected and
Sall4 morphant embryos were injected with fgf8a RNA. Again, Sall4 knockdown resulted in a
loss of hoxb9 expression (Figure 5.10A,B) without major alterations to sox2 expression (Figure
5.10E,F). Overexpression of fgf8a in the dorsal ectoderm resulted in an expansion of hoxb9
(spinal cord) expression, lateral expansion of krox20 expression, and repression of otx2 (brain)
(Figure 5.10C) (Fletcher et al., 2006). Overexpressing fgf8a in Sall4 morphants resulted in otx2
repression, but also resulted in a loss of hoxb9 expression (Figure 5.10D). krox20 expression in
rhombomere 5 was severely reduced in the Sall4 morphants despite fgf8a overexpression while
rhombomere 3 expression remained expanded. These results support the conclusion that Sall4 is
required for Fgf-induced posterior neural differentiation.
To address the question of whether RA-induced caudalization requires Sall4, uninjected and
Sall4 morphant embryos were incubated in RA. Increasing RA signaling results in severe loss of
anterior neural tissue and expansion of posterior identities (Blumberg et al., 1997; Durston et al.,
1989; Ruiz i Altaba and Jessell, 1991; Shiotsugu et al., 2004; Sive et al., 1990). Uninjected
control embryos treated with 1 µM all-trans retinoic acid (ATRA) lacked otx2 and krox20 but had
hoxb9 expression (Figure 5.10I). Sall4 morphant embryos treated with 1 µM ATRA failed to
express otx2 and krox20, however, they also failed to express hoxb9 (Figure 5.10J). The
reduction of these markers was not due to a loss of neural tissue as sox2 expression was similar
among control embryos, embryos treated with ATRA, and Sall4 morphant embryos treated with
ATRA (Figure 5.10E,K,L). Thus, Sall4 is required for caudalization of the neural plate via both
Fgf and RA signaling.
Discussion
Wnt, Fgf, and RA signaling are caudalizing factors required for posteriorization of the neural
plate. However, the transcription factors identified to mediate patterning signals from these
pathways have largely been restricted to those specifying midbrain and hindbrain fates. In this
study, I show that canonical Wnt signaling directly activates sall4, which is required for spinal
cord differentiation. The primary role of Sall4 in Wnt-induced posterior patterning is the
repression of the pouV/Oct4 homologs. This repression is necessary for spinal cord
differentiation; Sall4 knockdown as well as pouV/Oct4 overexpression results in a loss of spinal
73
cord fate. Furthermore, the posterior defects in Sall4 morphants can be rescued via pouV/Oct4
knockdown. The data presented here suggest a model whereby the repression of pouV/Oct4 via
Sall4 provides a permissive environment, allowing cells in the neural plate to respond to
posteriorizing signals from Fgf, RA, and Wnt (Figure 5.10M).
Other studies have shown interactions between Wnt signaling and Sall factors. Sall2 (published
as XsalF) functions as a Wnt antagonist in anterior neural tissue (Onai et al., 2004). Our work
found sall4 to be directly activated by Wnt signaling in the neurectoderm. TCF/LEF binds to
and activates the sall4 gene in human cell culture lines at a conserved binding site within the
promoter (Böhm et al., 2006). This conservation is restricted to mammals: the chick Gallus
gallus and Xenopus lack TCF/LEF sites in the promoter regions of sall4. However, the first
intron of Xenopus laevis sall4 contains six putative consensus TCF/LEF binding sites, three of
which are found within a 150 bp region and conserved in Xenopus tropicalis, though not in
chickens. Our finding that this region is enriched upon β-catenin ChIP demonstrates that βcatenin is bound to it during late gastrulation at the appropriate time to transduce the Wnt signal
and activate sall4, resulting in the down-regulation of pouV/Oct4. In the future, mutagenesis of
the consensus binding sites will test whether they are indeed required for Wnt-induced sall4
expression.
Our results provide a novel mechanism of neural posteriorization wherein Wnt signaling
activates sall4 in order to repress inhibitors of neural differentiation. A key prediction of this
mechanism is that a requirement for Sall4 in adoption of posterior fates would not be restricted to
Wnt-induced posteriorization. Therefore, one would expect that the increase in pouV/Oct4
expression resulting from Sall4 knockdown would inhibit differentiation that was induced by
other caudalizing factors. Indeed, I found Sall4 knockdown prevented induction of hoxb9 by Fgf
or RA. Future experiments will test whether overexpression of pouV/Oct4 is sufficient to block
posteriorization by Fgf and RA.
While I show that sall4 is a target of Wnt signaling, it remains possible that sall4 is also
regulated by other signaling pathways. In flies, spalt is regulated by Dpp (BMP), Hedgehog and
EGF signaling, depending on tissue type (de Celis et al., 1996; Elstob et al., 2001; Sturtevant et
al., 1997). sall1 is regulated by both Fgf and Wnt signaling in the chick limb (Farrell and
Münsterberg, 2000) and sall4 is expressed in the Xenopus limbs during development and
regeneration (Neff et al., 2011). The expression pattern of sall4 at early neurula stages is broader
than what would be expected for a gene solely regulated by Wnt. This suggests possible
activation via multiple pathways. One possibility could be that sall4 expression is regulated
through different enhancers, each responsible for different expression domains. In the chick,
sox2 is expressed throughout the neural plate but is regulated by five different enhancers, each
responsible for a portion of the full expression domain (Uchikawa et al., 2003). Furthermore,
Fgf and Wnt signaling converge on one such enhancer, N-1, to mediate the most posterior
expression of sox2 in the neural plate (Takemoto et al., 2006).
74
Fgf signaling is sufficient to posteriorize neurectoderm (Christen and Slack, 1997; Fletcher et al.,
2006; Kengaku and Okamoto, 1995; Lamb and Harland, 1995), and I found this activity requires
Sall4. Therefore, it is possible that FGF and canonical Wnt signaling converge on an as-yet
unidentified enhancer to regulate sall4 expression. As in the sox2 N-1 enhancer, Wnt- and Fgfresponsive elements in the enhancers of pax3 and zic genes cooperatively regulate their
expression (Garnett et al., 2012), and both pathways are required for expression of these genes at
the neural plate border (Monsoro-Burq et al., 2005).
Multiple studies have identified Wnt signaling as a key factor in stem cell maintenance and
differentiation (Wang and Wynshaw-Boris, 2004). In amphibians, caudalization of the neural
plate via canonical Wnt signaling induces undifferentiated neural precursors to commit to
posterior fates. This induction requires repression of stem cell factors and the activation of
differentiating factors. In Xenopus, pouV/Oct4 play a conserved role in maintaining pluripotency
of uncommitted cells (Morrison and Brickman, 2006). pouV/Oct4 are first expressed animally in
cleavage stages and throughout the mesoderm and ectoderm of amphibian gastrulae (Frank and
Harland, 1992; Morrison and Brickman, 2006). Knockdown of pouV/Oct4 results in precocious
cell fate commitment in the three germ layers (Morrison and Brickman, 2006; Snir et al., 2006).
The spatiotemporal expression of pouV/Oct4 serves to prevent these cells from adopting
inappropriate fates due to premature response to differentiating signals. Accordingly, pouV/Oct4
overexpression prolongs the undifferentiated state (Archer et al., 2011; Morrison and Brickman,
2006). Our results suggest that pouV/Oct4 expression must be down-regulated in the
neurectoderm to allow for cells to respond to instructive Wnt/Fgf/RA signals and commit to
posterior fates.
Wnt signaling activates cdx1 (Pilon et al., 2007; Prinos et al., 2001) and in frogs, Cdx1 represses
pouV/Oct4 gene expression at the onset of gastrulation (Rousso et al., 2011). However,
knockdown of Cdx1 does not result in a loss of spinal cord differentiation, and combinatorial
knockdown of Cdx1/2/4 is required before hoxb9 and hoxc10 expression is reduced (Faas and
Isaacs, 2009). There is, however, a dramatic loss of hoxb9, hoxc10, and hoxd10 expression in
Sall4 morphants. In the absence of Sall4, pouV/Oct4 expression remains high, resulting in neural
cells being unable to commit to a posterior neural fate and differentiate into spinal cord. Our
results suggest that Sall4, and not Cdx1, acts as the main negative regulator of pouV/Oct4 in the
neurectoderm.
My findings are consistent with described roles for the cdx genes in Wnt-mediated neural
patterning. Several studies have shown the Cdx factors function to regulate posterior hox gene
expression in vertebrates (Gaunt et al., 2004; Gaunt et al., 2008; Isaacs et al., 1998; van den
Akker et al., 2002). Therefore, Wnt acts as an instructive signal through activation of cdx genes
to induce posterior hox genes, thereby transforming the neural precursors into a posterior fate.
Here, I find that canonical Wnt also induces the expression of sall4 to repress pouV/Oct4,
providing a parallel, permissive signal for the induction of posterior hox gene expression via Cdx
factors. Wnt still signals in posterior neural regions of Sall4 morphants - presumably activating
cdx genes - but the prolonged expression of pouV/Oct4 prevents hox gene expression and
75
adoption of spinal cord fates. Conversely, it’s likely that Wnt induces sall4 expression in Cdx
morphants, priming the neural plate to respond to other instructive signals that induce
posteriorization. This mechanism could explain the finding that knockdown of individual Cdx
homologs results in minor phenotypes.
The interaction between Sall4 and Oct4 has been well studied in cell culture and mouse embryos.
In the ICM of mouse embryos and in iPS cells, Sall4 positively regulates Oct4/pou5f1 expression
allowing for self-renewal and suppression of differentiation (Tsubooka et al., 2009; Zhang et al.,
2006). However, I found Sall4 knockdown resulted in increased pouV/Oct4 gene expression in
neural tissue, which suggests that Sall4 negatively regulates the Oct4 homologs in Xenopus
neurectoderm. One explanation for this apparent discrepancy is that cellular context affects
whether Sall4 will act as a transcriptional activator or repressor. The N-terminus of Sall4 recruits
DNA methyltranferases and the NuRD complex resulting in transcriptional inhibition of target
genes in HEK293 cells(Yang et al., 2012). Conversely, in embryonic stem (ES) cells, Sall4 was
found to directly bind to the promoter of pou5f1/Oct4 and activate transcription (Zhang et al.,
2006). Additionally, Sall4 regulates distinct circuitries between ES and extra-embryonic
endoderm cells, presumably due to the presence of different co-factors (Lim et al., 2008).
Therefore, it is likely that the regulation of pouV/Oct4 via Sall4 is conserved but is repressive in
the context of Xenopus neurectoderm. It remains unclear, however, if this regulation is direct or
indirect. While the mechanistic role of Sall4 in the neural plate is likely to be complex, our
experiments demonstrate Sall4 to be a pivotal factor in neural patterning.
76
Figure 5.1: sall4 is a direct transcriptional target of canonical Wnt-signaling. (A) qPCR on 5
whole embryos or 15-25 animal caps treated according to the conditions indicated on the X-axis.
The Y-axis shows relative expression to odc. meis3 and hoxb9 serve as controls for known direct
and indirect targets of Wnt-signaling, respectively. (B) Quantification of dorsalization in
uninjected embryos (open bars) and embryos injected animally with 500 pg FLAG-tagged βcatenin RNA (250 pg/blastomere) at the 2-cell stage (filled bars) as scored by the dorsoanterior
index (DAI). Error bars show 1 standard deviation from the mean. Images show a
representative uninjected (UC) embryo with an DAI of 7 (normal) and a representative embryo
with a DAI of 6 (kinked axis). (C) Schematic of the sall4 genomic locus in Xenopus laevis.
Blue boxes indicate exons and yellow circles indicate the location of putative TCF/LEF binding
sites. Black ovals show the locations of the zinc-finger domains. Numbers indicate the position
of putative binding sites relative to the transcription start site (TSS). Red octagon shows the stop
codon. (D) Chromatin immunoprecipitation of FLAG-tagged β-catenin in late gastrulae/early
neurulae. Open bars represent uninjected embryos and closed bars represent embryos injected
with 500 pg FLAG-tagged β-catenin (250 pg/blastomere at the 2-cell stage). Error bars represent
one standard deviation from the mean per cent input for each measurement. meis3 and xmlc2
serve as positive and negative controls for β-catenin binding, respectively. Means were
compared to the negative control (xmlc2) by one-way ANOVA followed by Tukey post-hoc
analyses (*: p<0.05).
77
A
B
450
sall4
8
DAI:7
7
DAI
6
350
5
UC
4
DAI:6
3
250
2
1
150
500 pg FLAG ȕ-catenin
0
50
C
6
sall4
hoxb9
5
+2387 +2882 +3885
+0 TSS
4
E3
E2
E1
E4
+2347 +2456 +3085
3
2
D
1
0
0.18
0.16
400
% Input
meis3
300
200
0.14
*
UC
FLAG
0.12
0.1
0.08
0.06
0.04
100
0.02
0
0
noggin
TVGR
EtOH
DEX
CHX
WE
-
+
-
+
+
+
-
+
+
+
-
+
+
+
+
xmlc2
meis3
sall4
intron 1
Animal Caps
78
Figure 5.2: Injected embryos express functional FLAG-tagged β-catenin. (A) Western blot
for the FLAG epitope in injected embryos. Actin serves as the loading control. (B)
Ventralization of embryos injected with β-catenin MO or co-injection of β-catenin MO with
different doses of FLAG-tagged β-catenin RNA. F-βcat: FLAG-tagged β-catenin.
79
A
UC
500 pg
FLAG-ȕ-catenin
1000 pg
FLAG-ȕ-catenin
ANTI-FLAG
ANTI-Actin
B
100%
Normal
90%
Partially
ventralized
80%
70%
Ventralized
60%
50%
40%
30%
20%
10%
0%
UC
250 pg 350 pg 500 pg
F-ȕcat
F-ȕcat F-ȕcat
+ 10 ng ȕ-catenin MO
750 pg
F-ȕcat
80
Figure 5.3: sall4 is expressed in the neurectoderm. (A-D) Whole-mount in situ hybridizations
of sall4 in Xenopus laevis embryos. (A) Whole mount stage 10 embryo stained for sall4, dorsovegetal view with the dorsal lip of the blastopore oriented towards the top. (B-D) Neurula stage
embryos are shown in dorsal view, with anterior toward the top. (E) Sagittal section of stage 10
embryo stained for sall4 expression, animal pole is to the top and dorsal is to the right. (F-G)
Transverse sections of stage 12 embryos stained for sall4, (F) anterior and (G) posterior. (H-I)
Transverse sections of stage 15 embryos stained for sall4, (H) anterior and (I) posterior. (J-K)
Transverse sections of stage 18 embryos stained for sall4, (J) anterior and (K) posterior. (E-K)
50µM sections, (F-K) all embryos are oriented with dorsal towards the top. SNE: sensorial
neurectoderm, No: notochord, S: somite, PM: paraxial mesoderm, PSM: presomitic mesoderm.
81
82
EE
A
10
G
F
G
F
B
PM
No
SNE
12
PM
I
I
H
H
C
No
PSM No
S
15
PSM
S
K
J
J
D
K
PSM
S
No
No
18
PSM
S
Figure 5.4: sall1 is directly activated by canonical Wnt signaling and expressed during
early embryogenesis. (A) qPCR on 5 whole embryos or 15 to 25 animal caps treated according
to the conditions indicated on the X-axis. The Y-axis shows expression relative to odc. (B-E)
Whole-mount in situ hybridizations of sall1 in Xenopus laevis embryos. (B) Whole mount stage
10 embryo stained for sall1, dorso-vegetal view with the dorsal lip of the blastopore oriented
towards the top. (B’) Sagittal section of stage 10.5 embryo stained for sall1 expression, animal
pole is to the top and dorsal is to the right. (C-D) Dorsal views of indicated neurula stage
embryos, anterior is oriented towards the top. (C’-C’’) Transverse sections of stage 12 embryos
stained for sall1, (C’) anterior and (C’’) posterior. (D’-D’’) Transverse sections of stage 15
embryos stained for sall1, (D’) anterior and (D’’) posterior. (E’-E’’) Transverse sections of stage
18 embryos stained for sall1, (E’) anterior and (E’’) posterior. (B’, C’-E’’) 50 µM sections, (C’E’’) dorsal oriented towards the top. No: notochord, S: somite, PSM: presomitic mesoderm.
83
A
10
450
B
sall1
350
10.5
B’
250
150
50
-
noggin
TVGR
EtOH
DEX
CHX
WE
-
+
-
+
+
+
-
+
+
+
-
+
+
+
+
Animal Caps
12
15
18
C
D
E
C’
D’
E’
C’’
D’’
C’
C’’
No
E’’
D’
E’
S
No
S
D’’
S
No
S
E’’
PSM
No
PSM
PSM
No
PSM
84
Figure 5.5: Loss of Sall4 results in a loss of posterior neural differentiation. (A-P) Wholemount in situ hybridization late neurula stage embryos shown in dorsal view with the anterior
toward the top. (A, C, E, G) uninjected control embryos and (B, D, F, H) embryos injected
bilaterally with 40 ng Sall4 MO (20 ng/blastomere at the 2-cell stage) . (A-B) Expression of
sox2 (C-D) Expression of n-tub. (E-F) Expression of snai2 and nkx6.1 (G-H) expression of
myoD. (I, K, M, O) Uninjected control embryos and (J, L, N, P) embryos injected with 20 ηg
Sall4 MO into the right animal-dorsal (A/D) blastomere. (I-J) Expression of otx2/krox20/hoxb9,
arrows show the relative anterior-posterior position of krox20 and the anterior limit of hoxb9. (KL) Posterior view of hoxb9 expression. (M-N) Posterior view of hoxc10 expression. (O-P)
Posterior view of hoxd10 expression. (Q-T) Quantification of A-P patterning defects associated
with Sall4 knockdown. (Q) Measurement of the length between the anterior-most expression of
otx2 and the first krox20 stripe in arbitrary units (AU) between the left (open bars) and right
(closed bars) side of uninjected control embryos and embryos injected with 20 ng Sall4 MO into
the right A/D blastomere. (R) Measurement of the length of hoxb9 expression domain in
arbitrary units (AU) between the left (open bars) and right (closed bars) side of uninjected
control embryos and embryos injected with 20 ng Sall4 MO into the right A/D blastomere. (S)
Measurement of the length of hoxc10 expression domain in arbitrary units (AU) between the left
(open bars) and right (closed bars) side of uninjected control embryos and embryos injected with
20 ng Sall4 MO into the right A/D blastomere. (T) Measurement of the length of hoxd10
expression domain in arbitrary units (AU) between the left (open bars) and right (closed bars)
side of uninjected control embryos and embryos injected with 20 ng Sall4 MO into the right A/D
blastomere. Error bars represent one standard deviation from the mean. Means were compared
between left and right sides by student’s T-test (*: p<0.05, **:p<0.01, ***:p<0.001).
85
86
I
UC
hoxd10
O
myoD
hoxc10
H
snai2/nkx6.1
G
M
E
F
hoxb9
n-tub
K
D
Sall4 MO (bilateral)
C
B
otx2/krox20/hoxb9
UC
sox2
A
P
N
L
Sall4 MO right A/D cell
J
Q
R
Length (AU)
S
T
0
5
10
15
20
25
30
35
0
2
4
6
8
10
12
0
2
4
6
8
10
12
14
16
18
0
5
10
15
20
25
30
Length (AU)
Length (AU)
Length (AU)
UC
UC
UC
UC
Left
Right
Sall4 MO
Right A/D Cell
*
Sall4 MO
Right A/D Cell
**
Sall4 MO
Right A/D Cell
***
Sall4 MO
Right A/D Cell
*
Figure 5.6: cdx2 is directly activated by canonical Wnt signaling and is not affected by Sall4
knockdown. (A) qPCR on 5 whole embryos or 15 to 25 animal caps treated according to the
conditions indicated on the X-axis. The Y-axis shows expression relative to odc. (B-C) cdx2
expression at stage 18 shown in dorsal view with the anterior toward the top. (B) Uninjected
control embryo. (C) Embryo injected with 20 g Sall4 MO in one animal-dorsal cell at the 4-cell
stage.
87
A
cdx2
450
350
250
150
50
noggin
TVGR
EtOH
DEX
CHX
WE
-
+
+
+
+
+
+
+
Animal Caps
UC
Sall4 MO right A/D cell
C
B
cdx2
+
+
+
+
8/8
5/6
88
Figure 5.7: Knockdown of Sall4 causes an increase in expression of the pouV/Oct4
homologs. (A-L) Whole-mount in situ hybridization of late neurula staged embryos shown in
dorsal view with the anterior toward the top. (A, D, G, J) Uninjected control embryos. (B, E, H)
Embryos injected with 20 ng Sall4 MO into the right A/D blastomere. (C, F, I) Embryos injected
bilaterally with 40 ng Sall4 MO (20 ng/blastomere at the 2-cell stage). (K-L) Embryos injected
with 150 ρg PouV RNA (50 pg pou25 RNA, 50 pg pou60 RNA, and 50 pg pou91 RNA) into the
right A/D blastomere. Red staining indicates β-galactosidase which is used as a tracer for RNA
injection. (M-N) Higher magnification view of indicated regions in K and L, respectively. (A-C)
Expression of pou25 (D-F) expression of pou60 (G-I) expression of pou91. (J-L) Expression of
otx2/krox20/hoxb9. (O) qPCR for pou25 in uninjected embryos, embryos injected with 40 ng
Sall4 MO (20 ng MO/blastomere at the 2-cell stage), and embryos injected with 40 ng Sall4 MO
+500 pg Xtsall4 RNA (20 ng MO/blastomere at the 2-cell stage+250 pg Xtsall4 RNA/dorsal
blastomere at the 4-cell stage). (P) qPCR for pou60 in uninjected embryos, embryos injected
with 40 ng Sall4 MO (20ng MO/blastomere at the 2-cell stage), and embryos injected with 40 ng
sall4 MO+500 pg Xtsall4 RNA (20 ng MO/blastomere at the 2-cell stage+250 pg Xtsall4 RNA/
dorsal blastomere at the 4-cell stage). (Q) qPCR for pou91 in uninjected embryos, embryos
injected with 40 ng Sall4 MO (20 ng MO/blastomere at the 2-cell stage), and embryos injected
with 40 ng sall4 MO+500 pg Xtsall4 RNA (20ng MO/blastomere at the 2-cell stage+250 pg
Xtsall4 RNA/dorsal blastomere at the 4-cell stage). (O-Q) Expression is measured relative to
odc. Error bars represent one standard deviation from the mean. Means compared to uninjected
control by one-way ANOVA followed by Tukey post-hoc analyses (*: p<0.05).
89
UC
A
Sall4 MO right A/D cell
B
Sall4 MO (bilateral)
C
O
4.5
pou25
*
Fold Change
3.5
2.5
1.5
0.5
pou25
UC
E
F
P
7
Fold Change
D
pou60
Sall4 MO
*
Sall4 MO
+Xtsall4 RNA
5
3
1
pou60
UC
G
H
I
Q
3.5
pou91
Sall4 MO
Sall4 MO
+Xtsall4 RNA
*
Fold Change
2.5
1.5
0.5
pou91
UC
UC
J
Sall4 MO
Sall4 MO
+Xtsall4 RNA
150 pg pouV RNA right A/D cell
K
L
M
N
otx2/krox20/hoxb9
90
Figure 5.8: A second non-overlaping Sall4 morpholino results in similar phenotypes. (A-F)
Whole-mount in situ hybridization if late neurula stage embryos shown in dorsal view with the
anterior toward the top. (A, C, E,) Uninjected control embryos and (B, D, F) embryos injected
with 20 ng Sall4 MO2 into the right A/D blastomere. (A-B) Expression of otx2/krox20/hoxb9
(C-D) Posterior view of hoxd10 expression. (E-F) Expression of pou91.
91
UC
A
Sall4 MO2 right A/D cell
B
otx2/krox20/hoxb9
C
D
hoxd10
E
F
pou91
92
Figure 5.9: Loss of spinal cord gene expression in Sall4 morphants requires an increase in
pouV/Oct4 expression. (A-M) Whole-mount in situ hybridization of late neuriula stage embryos
shown in dorsal view with anterior toward the top. (A-D) Uninjected control embryos. (E-H)
Embryos injected with 40 ng Sall4 MO (20 ng/blastomere at the 2-cell stage). (I-L) Embryos
injected with 40 ng Sall4 MO, 20 ng pou25 MO, 10 ng pou60a MO, 10 ng pou60b MO, and 20
ng pou91 MO (50 ng total MO/blastomere at the 2-cell stage). (M) Embryos injected with 20 ng
pou25 MO, 10 ng pou60a MO, 10 ng pou60b MO, and 20 ng pou91 MO (30 ng total MO/
blastomere at the 2-cell stage). (A, E, I, M) Expression of otx2/krox20/hoxb9. (B, F, J)
Expression of hoxc10. (C, G, K) Expression of hoxd10. (N) Quantification of posterior neural
gene expression as measured by length of expression domain in arbitrary units (AU). Open bars:
uninjected control embryos. Gray bars: embryos injected with 40 ng Sall4 MO (20 ng/
blastomere at the 2-cell stage). Closed bars: embryos injected with 40 ng Sall4 MO, 20 ng
Xlpou25 MO, 10 ng Xlpou60a MO, 10 ng Xlpou60b MO, and 20 ng Xlpou91 MO (50 ng/
blastomere at the 2-cell stage). Error bars represent one standard deviation from the mean.
Means compared to uninjected control by one-way ANOVA followed by Tukey post-hoc
analyses (***: p<0.001).
93
B
C
D
otx2/krox20/hoxb9
hoxc10
hoxd10
sox2
E
F
G
H
I
J
K
L
M
N
UC
Sall4 MO
Sall4 MO+pouV/Oct4 MO
35
30
25
Length (AU)
pouV/Oct4 MO
Sall4 MO
+pouV/Oct4 MO
Sall4 MO
UC
A
***
20
15
10
***
***
hoxb9
hoxc10
5
0
***
hoxd10
94
Figure 5.10: FGF and retinoic acid signaling fail to posteriorize Sall4 morphants. (A-L)
Whole-mount in situ hybridization of late neurula stage embryos shown in dorsal view with
anterior toward the top. (A, E) Uninjected control embryos. (B, F) Embryos injected with 40 ng
sall4 MO (20 ng/blastomere at the 2-cell stage). (C, G) Embryos injected with 50 pg fgf8a RNA
into the right animal dorsal (A/D) blastomere. (D, H) Embryos injected with 40 ng sall4 MO (20
ng/blastomere at the 2-cell stage) and 50 pg fgf8a RNA into the right A/D blastomere. (I, K)
Embryos treated with 1 µM all-trans retinoic acid (ATRA). (J-L) Embryos injected with 40 ng
sall4 MO (20 ng/blastomere at the 2-cell stage) and treated with 1 µM all-trans retinoic acid. (AD, I, J) Expression of otx2/krox20/hoxb9 (E-H, K, L) expression of sox2. (M) A model to
explain the role of Sall4 in Wnt-mediated neural posteriorization. Wnt activates sall4 then
represses pouV/Oct4 to allow for posteriorization via Wnt/Fgf/RA. Solid and dashed arrows
indicate direct and direct or indirect regulation, respectively.
95
UC
A
Sall4 MO +
50 pg fgf8a RNA
1 A/D Cell
50 pg fgf8a RNA
1 A/D Cell
Sall4 MO
B
C
D
F
G
H
otx2/krox20/hoxb9
E
sox2
G
Sall4 MO +
1 ȝM ATRA
1 ȝM ATRA
I
J
Wnt/ -catenin
M
Fgf/RA
Sall4
pouV/Oct4
Normal
Posterior neural
differentiation
(Hoxb9/Hoxc10/Hoxd10)
otx2/krox20/hoxb9
K
L
Wnt/ -catenin
Fgf/RA
Sall4
X
pouV/Oct4
Sall4
morphants
Posterior neural
differentiation
sox2
X
X
X
(Hoxb9/Hoxc10/Hoxd10)
96
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Appendix I
RNA-Seq results from Chapter 3:
Genes with >2-fold expression (direct Wnt activation vs. anterior neural)
Gene/Protein
Clone ID
Fold Increase
hnRNP H3
gi|52138902|gb|BC082630.1
1.51E+11
H3 histone, family 3B
gi|27503243|gb|BC042290.1
1.04E+11
glutamate ammonia ligase
gi|49256010|gb|BC073448.1
39422399227
protein phosphatase type 1 alpha,
catalytic subunit
gi|27695193|gb|BC041730.1
2824225487
ki-67
gi|115527315|gb|BC124560.1
1131777.541
copper chaperone for superoxide
dismutase
gi|50418348|gb|BC077488.1
3919.698435
foxI4.2
gi|50418055|gb|BC078036.1
1329.542265
ephrin-A4
gi|183985625|gb|BC166129.1
1297.844383
smad4
gi|54037962|gb|BC084196.1
1053.601949
cdx-2
gi|84105446|gb|BC111473.1
600.0062069
eukaryotic translation initiation factor 3
subunit 10
gi|35505403|gb|BC057711.1
414.3164277
churchill
gi|114107852|gb|BC123207.1
369.3076365
pip4k2a
gi|120537387|gb|BC129059.1
328.1431677
hnRNPk
gi|27882468|gb|BC044711.1
319.4817015
MGC83026
gi|49118646|gb|BC073670.1
226.469437
tpno2
gi|54673692|gb|BC084978.1
222.1449285
nol12
gi|114107789|gb|BC123345.1
151.6234281
epithelial V-like antigen 1
gi|50415563|gb|BC077583.1
147.2011472
sfrs6
gi|28422194|gb|BC044265.1
126.0892513
xirg protein-like
gi|213623421|gb|BC169722.1
87.788455
prickle1
gi|68533725|gb|BC098954.1
83.19938866
znf384
gi|50415185|gb|BC077403.1
69.76482898
rac-beta serine/threonine-protein kinase B
gi|47939912|gb|BC072041.1
62.12571541
118
Gene/Protein
Clone ID
Fold Increase
ccbl-2
gi|30046518|gb|BC051239.1
44.93558411
p80 katanin
gi|66910749|gb|BC097654.1
40.55422632
zeb2
gi|54648610|gb|BC084972.1
33.47771521
Zmiz1
gi|51513014|gb|BC080428.1
30.23438945
angiopoietin 4/5
gi|189442243|gb|BC167504.1
27.19110778
hcf-1
gi|52138923|gb|BC082658.1
26.78440995
ccr4-not transcription complex, subunit
10
gi|50416369|gb|BC077237.1
21.48403283
fam107a/b MGC78851
gi|51261937|gb|BC079918.1
21.17179772
nucleoporin Seh1B: MGC82845 protein
gi|49118558|gb|BC073561.1
19.13482551
pi3k related SMG1: hypothetical protein
MGC98890
gi|68226704|gb|BC098320.1
17.94963894
epsin-2: hypothetical protein MGC81482
gi|46249599|gb|BC068837.1
16.4173713
srsf7
gi|50603926|gb|BC077393.1
16.33581603
sf3b4
gi|28374169|gb|BC045264.1
15.37049865
pptc7: MGC81279 protein
gi|49257211|gb|BC071109.1
13.98198898
meis3
gi|54673770|gb|BC084920.1
13.07065969
origin recognition complex, subunit 6
homolog-like
gi|50603595|gb|BC077746.1
13.01809093
daxx: hypothetical protein LOC446279
gi|86577707|gb|BC112947.1
12.67764239
acsl4 hypothetical protein
LOC100174803
gi|189442239|gb|BC167498.1
11.62060714
necap2 MGC83534 protein
gi|50927256|gb|BC079728.1
10.9853218
timp3: tissue inhibitor of
metalloproteinases-3
gi|38014484|gb|BC060423.1
10.67580536
frizzled homolog 7
gi|27503170|gb|BC042228.1
9.299494092
serine/threonine/tyrosine-interacting
protein B
gi|54311224|gb|BC084791.1
9.188383287
ubadc1 hypothetical protein MGC115132
gi|62471528|gb|BC093557.1
8.970846126
cdca A7L transcription factor RAM2
gi|116487713|gb|BC126014.1
8.574819986
klf10: hypothetical protein MGC98877
gi|62089536|gb|BC092147.1
7.695378855
ivns1abp influenza virus NS1A binding
protein
gi|49898869|gb|BC076641.1
7.664198955
119
Gene/Protein
Clone ID
Fold Increase
MGC80567 protein
gi|50417996|gb|BC077854.1
7.544735234
lchn hypothetical protein MGC114999
gi|71050977|gb|BC098994.1
7.224153034
rabgap1l: hypothetical protein
MGC52980
gi|27694685|gb|BC043775.1
7.11745345
ptn1 pleiotrophin: MGC84465 protein
gi|49257697|gb|BC074426.1
6.911246415
arrb1 arrestin, beta 1
gi|49904092|gb|BC076815.1
6.832358987
txnrd3 Thioredoxin reductase 2
MGC81848 protein
gi|51704105|gb|BC081053.1
6.824096832
lims1-b LIM domain: hypothetical protein
MGC81174
gi|47939771|gb|BC072204.1
6.795291868
lmo7: LIM domain containing:
MGC180040
gi|197245592|gb|BC168520.1
6.755182581
arrdc3 arrestin containing hypothetical
protein MGC131006
gi|80476391|gb|BC108545.1
6.57050044
cant1 Calcium activated nucleotidase
similar to Ca2+-dependent endoplasmic
reticulum nucleoside diphosphatase
gi|27370857|gb|BC041215.1
6.486609662
d7 protein
gi|58702035|gb|BC090198.1
6.413210477
dact1 dapper 1 Antagonist of beta-catenin
FRODO
gi|50418314|gb|BC077380.1
6.403341734
rassf7 Ras assiciation domain containing
MGC78972 protein
gi|84105479|gb|BC111512.1
6.017970041
sox11 XLS13B protein
gi|47124741|gb|BC070707.1
5.989392572
myt1 cDNA clone MGC:196991
gi|213626262|gb|BC170264.1
5.974437792
zmiz2 MGC86475 protein
gi|51513014|gb|BC080428.1
5.658053905
zc3h7b zinc-finger CCCH-containing 7B
MGC80522 protein
gi|50418254|gb|BC077837.1
5.638059804
pcna similar to proliferating cell nuclear
antigen
gi|27371152|gb|BC041549.1
5.340877685
stx19 syntaxin 19 hypothetical
LOC494752
gi|52354747|gb|BC082852.1
5.239206209
hmg-box protein HMG2L1
gi|213625180|gb|BC169998.1
5.171640761
kif20a hypothetical LOC495414
gi|54648449|gb|BC084922.1
5.055010856
slc7a3 solute carrier family 7 (cationic
amino acid transporter, y+ system),
member 3
gi|27503399|gb|BC042222.1
4.989471538
120
Gene/Protein
Clone ID
Fold Increase
lmo7 cDNA clone MGC:180040
gi|197245592|gb|BC168520.1
4.861028301
mark2 MAP/microtubule affinityregulating kinase 2
gi|27694574|gb|BC043730.1
4.821716572
anp32b MGC80871 protein
gi|49118408|gb|BC073408.1
4.77985399
cyclin A2
gi|50417439|gb|BC077260.1
4.76329664
pppde2 peptidase domain containing
MGC84710 protein
gi|49256350|gb|BC074444.1
4.724302826
ctdp1 serine phosphatase
gi|62185666|gb|BC092306.1
4.712553945
ornithine decarboxylase-2
gi|28838468|gb|BC047954.1
4.690222394
ube2c hypothetical LOC496302
gi|57032917|gb|BC088818.1
4.676640452
Efr3a MGC83628 protein
gi|51950039|gb|BC082437.1
4.653269077
dlg7 discs large hypothetical protein
MGC116559
gi|68534624|gb|BC099363.1
4.501586994
stxbp3 hypothetical protein MGC115462
syntaxin binding protein 3 (stxbp3)
gi|72679360|gb|BC100235.1
4.472242676
acy-3: aspartoacylase-3
gi|116487526|gb|BC125990.1
4.452697089
ptdss2 cDNA clone MGC:179871
gi|197246680|gb|BC168517.1
4.234011971
tcf-7 transcription factor 7 (T-cell
specific, HMG-box)
gi|51261404|gb|BC079972.1
4.200569032
lsp1 lymphocyte specific protein
1hypothetical protein LOC100158340
gi|115528236|gb|BC124864.1
4.124150256
nphp3 nephronophthisis 3 MGC80264
protein
gi|50603779|gb|BC077320.1
4.066245125
med 15 Mediator complex subunit 15
ARC105 protein
gi|47123916|gb|BC070536.1
4.029208683
cyclin E3
gi|58701930|gb|BC090214.1
3.970372822
fam60a hypothetical protein MGC115222
gi|66910763|gb|BC097689.1
3.940864045
ahctf1 AT hook containing transcription
factor 1 MGC83673 protein
gi|49903664|gb|BC076775.1
3.892143367
rhebl1 Ras homolog enriched in brain like
1 hypothetical LOC495056
gi|54037975|gb|BC084211.1
3.882231045
rnf8a ring finger protein (C3HC4 type) 8
gi|28279439|gb|BC046256.1
3.801782364
ccnt2 cyclin T2 MGC81210 protein
gi|51895950|gb|BC081000.1
3.755306852
121
Gene/Protein
Clone ID
Fold Increase
tmed2 transmembrane emp24 domain
trafficking protein 2 coated vesicle
membrane protein
gi|28277265|gb|BC044095.1
3.747391508
mta1 metastatic associated 1 MGC83916
protein
gi|51950045|gb|BC082445.1
3.743645989
MAPK8/jnk1 mitogen-activated protein
kinase 8
gi|28422153|gb|BC046834.1
3.733178442
psmd4 26S proteasome subunit
gi|66910701|gb|BC097551.1
3.729782795
poldip3 polymerase delta interaction
protein 3 hypothetical protein
MGC114944
gi|62471555|gb|BC093543.1
3.720246762
dnajcb5 cDNA clone MGC:83536
gi|51703523|gb|BC081115.1
3.720172358
ncbp2 Nuclear cap binding protein 2
gi|49117074|gb|BC072902.1
3.701358817
fxdy domain containing ion transport
gi|125859119|gb|BC129686.1
3.694185141
ano5 Anoctamin 5 or Tmem16e
gi|50418049|gb|BC077486.1
3.642280513
Not Annotated
gi|62739385|gb|BC094151.1
3.628720112
ttc30a tetratricopeptide repeat domain
30a
gi|47938700|gb|BC072174.1
3.547737229
f2rl1 Coagulation factor 2 receptor like 1
gi|57033014|gb|BC088935.1
3.518659172
csda cols shock protein domain
containing A
gi|161611734|gb|BC155913.1
3.51654861
fus Fused in Sarcoma
gi|49522197|gb|BC074437.1
3.505453855
exo1 exonuclease 1
gi|54035217|gb|BC084102.1
3.494289274
cfp complement factor properdin
gi|50415018|gb|BC077925.1
3.468804465
ferritin light chain
gi|34785676|gb|BC057216.1
3.464575104
cdc25c
gi|213626377|gb|BC169346.1
3.456754005
slc44a1 solute carrier family 44 member
1
gi|52354612|gb|BC082837.1
3.306234736
pcf11 cleavage and poly-adenylation
factor
gi|50414592|gb|BC077233.1
3.277333059
slc9a1 or NHE3 solute carrier family 9
member 3
gi|157422994|gb|BC153791.1
3.274941479
anks1a Ankyrin repeat and sterile alpha
motif domain containing 1a
gi|47682305|gb|BC070831.1
3.249886264
122
Gene/Protein
Clone ID
Fold Increase
ap2b1 adaptor-related protein complex 1
beta 1 subunit
gi|120538239|gb|BC129531.1
3.240669681
Not Annotated
gi|76780224|gb|BC106027.1
3.21623043
ctnnd1 Catenin (Cadherin associated
protein) delta-1
gi|213623207|gb|BC169434.1
3.210767484
gcat Glycine C-acetyltransferase
gi|28704125|gb|BC047258.1
3.210735376
beta arrestin
gi|49256118|gb|BC072973.1
3.173896459
slc9a3r2
gi|55778573|gb|BC086464.1
3.167840103
ctdp1 (carboxy-terminal domain, RNA
polymerase II, polypeptide A)
phosphatase, subunit 1
gi|51950263|gb|BC082378.1
3.162965383
max bHLH
gi|47123961|gb|BC070710.1
3.144295944
mpv17l
gi|51261416|gb|BC079982.1
3.11285403
fibronectin 1
gi|49114986|gb|BC072841.1
3.110364743
sfrs5
gi|47717980|gb|BC070967.1
3.1059201
transmembrane protein 45B
gi|120538262|gb|BC129609.1
3.030355684
lysine (K)-specific demethylase 6A
gi|50603932|gb|BC077424.1
3.026903047
ralGDS/AF-6
gi|84105479|gb|BC111512.1
2.963378492
mek-2
gi|27694983|gb|BC043913.1
2.955122189
calpain 2, (m/II) large subunit
gi|39645066|gb|BC063733.1
2.924548179
phd finger protein 12
gi|46249573|gb|BC068803.1
2.89562217
pax interacting (with transcriptionactivation domain) protein 1
gi|50417566|gb|BC077588.1
2.822971349
mediator complex subunit 16
gi|62471580|gb|BC093546.1
2.822152806
xrmd-2 microtubule-associated protein
gi|58702063|gb|BC090235.1
2.803700074
tyrosine kinase 2
gi|49118136|gb|BC073112.1
2.790804764
methyltransferase like 3
gi|46249483|gb|BC068672.1
2.782222309
glycine amidinotransferase (Larginine:glycine amidinotransferase)
gi|28838491|gb|BC047973.1
2.746369891
syntaxin 5
gi|76779222|gb|BC106704.1
2.704962367
inhibitor of kappa light polypeptide gene
enhancer in B-cells, kinase beta
gi|47939754|gb|BC072192.1
2.686442963
G-2 and S-phase expressed 1
gi|62471553|gb|BC093540.1
2.683239948
123
Gene/Protein
Clone ID
Fold Increase
rbl1
gi|47123210|gb|BC070856.1
2.680418663
nucleoporin 93kDa
gi|27924241|gb|BC045089.1
2.672333338
embryonic ectoderm development
gi|50603665|gb|BC077425.1
2.655016847
ring finger and CCCH-type domains 1
gi|46250191|gb|BC068669.1
2.646867856
integrin, beta 5
gi|49899756|gb|BC076844.1
2.636182901
ataxin 2
gi|66910767|gb|BC097692.1
2.634583223
chromosome 19 open reading frame 2
gi|50415135|gb|BC077366.1
2.630865817
prp4 pre-mRNA processing factor 4
homolog
gi|51703477|gb|BC081044.1
2.62131998
protein phosphatase methylesterase 1
gi|50418398|gb|BC077600.1
2.617432826
orthodenticle homeobox 2
gi|50417481|gb|BC077357.1
2.616883223
chromosome 13 open reading frame 34
gi|49523107|gb|BC075159.1
2.599294339
dazap1
gi|50604139|gb|BC077252.1
2.585999275
fshd region gene 1
gi|49256477|gb|BC074376.1
2.555875944
serine/threonine kinase 11 interacting
protein
gi|47682952|gb|BC070809.1
2.553165597
carboxy-terminal kinesin 2
gi|54038135|gb|BC084431.1
2.538623487
survival of motor neuron 2, centromeric
gi|46249513|gb|BC068721.1
2.535840144
sal-like 1
gi|37590272|gb|BC059284.1
2.505331347
nima (never in mitosis gene a)-related
kinase 2
gi|27696903|gb|BC043822.1
2.503175185
zf-containing
gi|213623475|gb|BC169799.1
2.493496644
drebrin-like
gi|49257631|gb|BC074277.1
2.479066307
jumonji domain containing 6
gi|28277358|gb|BC045252.1
2.4687995
inhibitor of DNA binding 3, dominant
negative helix-loop-helix protein
gi|27696824|gb|BC044039.1
2.448101925
chaperonin containing TCP1, subunit 8
(theta)
gi|67678231|gb|BC097574.1
2.447348026
LIM domain containing preferred
translocation partner in lipoma
gi|62740239|gb|BC094110.1
2.445439839
cytochrome c-1
gi|71052231|gb|BC099350.1
2.442233526
kiaa0182
gi|120537359|gb|BC129052.1
2.438699731
124
Gene/Protein
Clone ID
Fold Increase
5-aminoimidazole-4-carboxamide
ribonucleotide formyltransferase/IMP
cyclohydrolase
gi|76779775|gb|BC106381.1
2.42732299
ribonucleoprotein A1a
gi|47938743|gb|BC072090.1
2.419006697
caspase 3, apoptosis-related cysteine
peptidase
gi|68533747|gb|BC098991.1
2.408087828
ubiquitin-conjugating enzyme E2G 1
(UBC7 homolog)
gi|28839012|gb|BC047985.1
2.407955386
protein tyrosine kinase 7
gi|148922111|gb|BC146640.1
2.387741643
integrator complex subunit 2
gi|47125091|gb|BC070524.1
2.387717766
prpf4b
gi|125858002|gb|BC129065.1
2.375801846
transmembrane protein 33
gi|49903380|gb|BC076764.1
2.371301594
non-SMC condensin II complex, subunit
D3
gi|49116983|gb|BC073714.1
2.363179599
sin3 homolog B, transcription regulator
gi|120538596|gb|BC129063.1
2.353559822
splicing factor, arginine/serine-rich 18
gi|47940261|gb|BC072160.1
2.350873591
mediator complex subunit 23 med23
gi|39645714|gb|BC063725.1
2.349851184
phospholipase A2-activating protein
gi|115528262|gb|BC124847.1
2.344309729
minichromosome maintenance complex
component 4
gi|49115033|gb|BC072870.1
2.342847336
nop2/sun domain family, member 2
gi|66912075|gb|BC097814.1
2.339817652
general transcription factor IIE,
polypeptide 2, beta 34kDa (gtf2e2)
gi|58403335|gb|BC089287.1
2.320004209
rho GTPase activating protein 19
gi|48734660|gb|BC072338.1
2.309370554
ccr4-not transcription complex, subunit
10
gi|46250097|gb|BC068748.1
2.298100702
lysine (K)-specific demethylase 3A
gi|47506877|gb|BC070982.1
2.296984096
zinc finger and BTB domain containing
44
gi|47124748|gb|BC070714.1
2.293259115
phosphatidylinositol glycan anchor
biosynthesis, class T
gi|52354598|gb|BC082818.1
2.284755462
heterogeneous nuclear ribonucleoprotein
A3
gi|213625122|gb|BC169881.1
2.283526595
Putative ortholog of von Hippel-Lindau
binding protein 1 (Prefoldin subunit 3)
gi|163916339|gb|BC157499.1
2.278221284
125
Gene/Protein
Clone ID
Fold Increase
nucleoporin 37kDa
gi|51703531|gb|BC081128.1
2.271537693
activating transcription factor 1
gi|61403334|gb|BC092037.1
2.266325959
nedd4 family interacting protein 2
gi|50924805|gb|BC079714.1
2.262854343
gi|33416619|gb|BC055957.1
2.260893298
proteasome (prosome, macropain) 26S
subunit, ATPase, 3
gi|28422358|gb|BC046948.1
2.253753391
family with sequence similarity 109,
member B
gi|47122977|gb|BC070645.1
2.237428018
translation initiation factor 4E family
member 3
gi|49257962|gb|BC071126.1
2.230893103
ets variant gene 4
gi|50417509|gb|BC077414.1
2.224884491
G kinase anchoring protein 1
gi|49118875|gb|BC073450.1
2.208726268
sall-like 4
gi|52138969|gb|BC082637.1
2.190818022
chromobox homolog 5
gi|32766466|gb|BC054962.1
2.18484743
ccr4-not transcription complex, subunit 6like
gi|47506927|gb|BC071015.1
2.17052701
uridine-cytidine kinase 2
gi|52354745|gb|BC082833.1
2.153018907
yy1 transcription factor b
gi|50925274|gb|BC079731.1
2.144522678
karyopherin alpha 4 (importin alpha 3)
gi|47122818|gb|BC070533.1
2.143067042
syntaxin 5
gi|76779222|gb|BC106704.1
2.132374185
PRP4 pre-mRNA processing factor 4
homolog B
gi|54038077|gb|BC084355.1
2.120332678
oxoglutarate (alpha-ketoglutarate)
dehydrogenase (lipoamide)
gi|49118216|gb|BC073213.1
2.110063412
acidic (leucine-rich) nuclear
phosphoprotein 32 family, member B
gi|27503409|gb|BC042250.1
2.104752746
AT hook containing transcription factor 1
gi|55250536|gb|BC086281.1
2.095665156
proline-rich nuclear receptor coactivator
2
gi|54038003|gb|BC084247.1
2.080782448
yy1
gi|50415555|gb|BC077581.1
2.079401267
ptk7
gi|38014809|gb|BC060500.1
2.074966481
H3 histone, family 3B (H3.3B)
gi|47506868|gb|BC070966.1
2.05094159
bromodomain containing 1
gi|49118425|gb|BC073421.1
2.046475407
126
Gene/Protein
Clone ID
Fold Increase
mllt6
gi|52354628|gb|BC082872.1
2.041361526
RAS oncogene family
gi|33416685|gb|BC056054.1
2.03028667
RAB6A, member RAS oncogene family
gi|28302337|gb|BC046683.1
2.027277987
transcription factor 3 (E2A
immunoglobulin enhancer binding factors
E12/E47)
gi|28422165|gb|BC046840.1
2.026584776
cell division cycle 20 homolog
gi|50370183|gb|BC076805.1
2.012178568
sema domain, transmembrane domain
(TM), and cytoplasmic domain,
(semaphorin) 6D
gi|213626595|gb|BC169687.1
2.010828849
lethal giant larvae homolog 1
gi|47123133|gb|BC070788.1
2.000831812
127
Appendix II
List of PCR primers used in this work
Primers used for Cel1 assays:
Gene
Forward
Reverse
xt
noggin
P''''''P P''''P
eGFP
5’-CAGTGCTTCAGCCGCTACC-3’
5’-CTGGTAGTGGTCGGCGAGC-3’
Primers used for RT-PCR and qPCR:
Gene
Forward
Reverse
cdx2
(qPCR)
5’-ACATACCGGGATCCAAGACA-3’
5’-CAGCCTGAGTCTGCTGGATT-3’
eef1a1
(RT-PCR/
qPCR)
5’-CCCTGCTGGAAGCTCTTGAC-3’
5’-GGACACCAGTCTCCACAC
GA-5’
en2 (RTPCR)
5’-CAGCCTGGGTCTACTGCAC-3’
5-CTTTGCCTCCTCTGCTCAGT-3’
epidermal
keratin
(RT-PCR)
5’-GACCTGGAAGGGAAGATCC-3’
5’-GAAGAGCCAGCTCATTCT
CAA-3’
hoxb9
(qPCR)
5’-TACTTACGGGCTTGGCTGGA-3’
5’-AGCGTGTAACCAGTTGG
CTG-3’
hoxb9
(RT-PCR)
5’-CTCCAGCAGCCAAATTCTCT-3’
5’-CAGTTGGCTGAGGGGTTG-3’
krox20
(RT-PCR)
5’-CCAGTGACTTTTGGTAGTTT
TGTG-3’
5’-TGGACGAGTAGGAGAA
ATCCA-3’
meis3
(qPCR)
5’-CAGGATACAGGGCTCACGAT-3’
5’-CTTGGGGCTGCTGTGTAATC-3’
meis3
(RT-PCR)
5’-ATGATCGTGATGGCTCTTCC-3’
5’-CCCTGTGCGATTAGATTGGT-3’
128
Gene
Forward
Reverse
muscle
actin (RTPCR)
5’-GACTCTGGGGATGGTGTGAC-3’
5’-AGCAGTGGCCATTTCATTCT-3’
odc (RTPCR/
qPCR)
5'-GGGCTGGATCGTATCGTAGA-3'
5'-TGCCAGTGTGGTCTTGACAT-3'
otx2 (RTPCR)
5’-TATCTCAAGCAACCGCCATA-3’
5’-AACCAAACCTGGACTCT
GGA-3’
pou25
(qPCR)
5’-GGGCCACCACTATCCCTAAT-3’
5’-GTGTGTAGCCCAGGGAC
ACT-3’
pou60
(qPCR)
5’-AGTTTGCCAAGGAGCTGAAA-3’
5’-GGACTCAAAGCGGCAG
ATAG-3’
pou91
(qPCR)
5’-ACTTATTTGCCCCGTCTCCT-3’
5’-CCCCATTCAGATCACTTGCT-3’
sall1
(qPCR)
5’-GAGAGGGGTCAAATCCATCG-3’
5’-GGAGGTGGTGGATTTTCA
TTC-3’
sall4
(qPCR)
5’-TGTCAAAGGATGAGCATTCG-3’
5’-CATGCGGTCAGAGGGTACTT-3’
Primers used for ChIP:
Gene
Forward
Reverse
meis3
5’-CACTGTAAGTTATTGCCTCAA
AGG-3’
5’-AGCTTGTAATACTTGTGGG
CTTT-3’
sall4
intron 1
5’-GGGAGTTGGAAGGTACAAAGC-3’
5’-AACCAAACAATAGACGAAA
AATAAA-3’
xmlc2
5’-TGGGATATTTTACTGAACACA
ATG-3’
5’-CGTCCTGTGCCACCTA
ATG-3’
129
Primers used for probe synthesis:
Gene
Forward
Reverse
Xl sall1
5’-CTTTCAAAGCATGGTGA
GCA-3’
5’-ATGGCACGATGGACAC
TGTA-3’
Xl sall4
5’-CTTGGTGCGCACTTAT
CTCA-3’
5’-GCCTCAGATTGTGTGGG
ACT-3’
Xl hnRNPH3
5’-GAAAATGCTCTGGGGAA
ACA-3’
5’-TCGTGTGTTGCAAATTC
CAC-3’
Primers used for Sub-cloning (Underline: RE site and lowercase: FLAG-tag sequence):
Gene
Forward
Reverse
Xt noggin
(Full length)
5’-GAATTCGGGCTCTGAAC
TTCCACTTG-3’
5’-GCGGCCGCTCAACATGAA
CATTTGCACTCA-3’
Xl FLAG-βcatenin
5’5’-GCTAGCGGCCGCTTActtatcg
GCATGAATTCCCACCATGGCAAC tcgtcatccttgtaatcCAAGTCAGTGTC
TCAAGCAGATCT-3’
AAACCAGG-3’
Xt sall4 (Full
length)
5’-CGATGTCGACGGACCAT
GTCGAGGCGAAAGCAGCC-3’
5’-ATCGATCCTCGAGTTActtatcg
tcgtcatccttgtaatcGTTCACCGCAAT
ATTTT-3’
130
Appendix III
List of DNA plasmids used in this work
Plasmids used for in situ hybridization probe sythesis:
Plasmid #
Gene
Vector
Cut
with:
Transcribe
with:
Note
96
hoxb9
pGEM
EcoRI
T7
97
myoD
pBS KS+
HinDIII
T7
167
krox20
pGEM4
BamHI
Sp6
302
otx2
pBS SK-
NotI
T7
1104
sox2
pCS2
EcoRI
T7
1346
hoxd10
pCS-107
SalI
T7
1425
hoxc10
pCS-107
SalI
T7
2048
Nkx6.1
pBS
XhoI
T7
2396
n-tubulin
pBSK
BamHI
T7
2491
snai2
pBS SK-
NotI
T3
2603
sall4
TOPO pCRII
KpnI
T7
Not full length
2604
sall1
TOPO pCRII
BamHI
T7
Not full length
2605
pou60
CS2
HinDIII
T7
2606
pou25
CS2
BamHI
T7
2607
pou91
CS2
BamHI
T7
2615
cdx2
pGEMT-EZ
SacII
Sp6
2616
hnRNP H3 TOPO pCRII
NotI
T7
154 (trop)
en2
TOPO pCRII
SpeI
T7
155 (trop)
krox20
TOPO pCRII
SpeI
T7
187 (trop)
hoxb9
pCMVSPORT6
EcoRI
T7
203 (trop)
msx2
pCS-107
EcoRI
T7
Not full length
131
Plasmid #
Gene
Vector
Cut
with:
Transcribe
with:
Note
274 (trop)
sox9
pCS-107
BsrgI
T7
287 (trop)
edn1
PCS-107
EcoRI
T7
288 (trop)
col2a
pCMVSPORT6
AgeI
T7
306 (trop)
bmp7
pCMVSPORT6
KpnI
T7
31 (trop)
otx2
pCS-107
EcoRI
T7
471 (trop)
hand2
pCS-107
EcoRI
T7
472 (trop)
ahctf1
pCS-108
SalI
T7
Image: 7552963
473 (trop)
churchill
pCMVSPORT6
EcoRI
T7
Image: 6980207
474 (trop)
foxi4.2
pCMVSPORT6
EcoRI
T7
Image: 5336445
475 (trop)
znf384
pExpress1
XbaI
T7
Image: 7017260
476 (trop)
zmiz1
pCMVSPORT6
EcoRI
T7
Image: 7793374
477 (trop)
max
pCMVSPORT6
EcoRI
T7
Image: 6988059
478 (trop)
sall1
pCMVSPORT6
EcoRI
T7
Image: 7677318
479 (trop)
sall4
pCMVSPORT6
EcoRI
T7
Image: 5307468
480 (trop)
sox11
pCMVSPORT6
EcoRI
T7
Image: 6979794
481 (trop)
hnRNPk
pCMVSPORT6
EcoRI
T7
Image: 5379554
482 (trop)
sf3b4
pCMVSPORT6
EcoRI
T7
Image: 5383583
483 (trop)
sap130
pCMVSPORT6
EcoRI
T7
Image: 7689619
484 (trop)
sfrs7
pCMVSPORT6
EcoRI
T7
Image: 6976658
485 (trop)
sfrs6
pCMVSPORT6
EcoRI
T7
Image: 5307339
486 (trop)
prickle1
pCS-107
EcoRI
T7
TEgg011j18
487 (trop)
lmo7
pCMVSPORT6
EcoRI
T7
Image: 7677402
132
Plasmids used for synthetic RNA synthesis:
Plasmid #
Gene
Vector
Cut with:
Transcribe
with:
Note
671
β-gal (Nuc.)
CS2+
NotI
Sp6
2304
Xt fgf8a
pCS108
AscI
Sp6
2444
TVGR
CS2+
NotI
Sp6
2550
25726EL
pCS-108
AscI
Sp6
2551
25728KK
pCS-108
AscI
Sp6
2552
eGFP-ZFN L
pCS-108
AscI
Sp6
2553
eGFP-ZFN R
pCS-108
AscI
Sp6
2554
25728EL
pCS-108
AscI
Sp6
2554
25758EL
pCS-108
AscI
Sp6
2555
25760EL
pCS-108
AscI
Sp6
2556
25763EL
pCS-108
AscI
Sp6
2557
25766KK
pCS-108
AscI
Sp6
2609
Xt sall4-FLAG
pCS-108
AscI
Sp6
2611
+4 bp noggin
pCS-108
AscI
Sp6
2612
–12 bp noggin
pCS-108
AscI
Sp6
2613
+3 bp noggin
pCS-108
AscI
Sp6
2617
25729EL
pCS-108
AscI
Sp6
2618
25730EL
pCS-108
AscI
Sp6
2619
25731KK
pCS-108
AscI
Sp6
2620
β-catenin-FLAG pCS-108
AscI
Sp6
2621
25760 WT
pCS-108
AscI
Sp6
Cut with NotI for
polyA-
2622
25766 WT
pCS-108
AscI
Sp6
Cut with NotI for
polyA133
Plasmid #
335 (trop)
Gene
noggin
Vector
pCS-107
Cut with:
Transcribe
with:
AscI
Sp6
Note
134