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Making a mouth: elucidating morphogenetic events of mouth development in the sea urchin
Lytechinus variegatus
Lauren K. Sibley
Under the supervision of Dr. David McClay,
Department of Biology, Duke University
May 2017
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Research Supervisor
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Faculty Reader
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Director of Undergraduate Studies
Honors thesis submitted in partial fulfillment of the requirements for graduation with
Distinction in Biology in Trinity College of Duke University
Abstract
Deuterostomes are bilaterian animals in which the blastopore, the site of gastrulation, becomes
the anus and the mouth develops secondarily. Deuterostome phyla include Chordata, such as vertebrates
like ourselves, and Echinodermata, such as sea urchins. The extent of homology among deuterostome
mouths is unknown. To address this question, this thesis compares three aspects of mouth
morphogenesis between frogs (Xenopous laevis), a vertebrate, and sea urchins (Lytechinus variegatus),
an echinoderm and basally branching deuterostome: 1) if mouth formation requires signaling from the
gut endoderm, 2) if Wnt signaling regulates basement membrane dissolution during mouth development,
and 3) if the mouth perforates by apoptosis. It was found through gut removal and isolation experiments
and by inducing exogastrulation that sea urchins may not require signaling from the gut for mouth
formation. Treating sea urchin embryos with C59, a Wnt signal-inhibiting drug, developed smaller
mouths as the level of Wnt-inhibition increased. Lastly, an apoptosis assay that immunostained embryos
for anti-caspase3 antibody revealed that sea urchin mouths may not open by apoptosis. We found that
these three aspects of mouth development contrast in the sea urchin and frog, supporting less homology
among deuterostome mouths.
Introduction
In deuterostomes, a super-phylum of bilaterian animals, the blastopore, the site of gastrulation,
becomes the anus and the mouth develops secondarily. Major deuterostome groups include
ambulacrarians (echinoderms and hemichordates), vertebrates, tunicates, and cephalochordates (Fig. 1).
The pharyngeal opening, not the external mouth, in vertebrates is thought to be homologous to the
mouths of the other deuterostome groups (Dickinson and Sive, 2009). I use “mouth” to refer to the
pharyngeal opening where the ectoderm and endoderm meet with no intermediate mesoderm.
“Stomodeum” refers to the patch of ectoderm that will become the mouth.
Fig. 1 Phylogenetic tree of bilaterian animals (adapted from (Bourlat et al., 2006; Martindale and
Hejnol, 2009))
Bilaterian animals are divided into two superphyla: protostomes and deuterostomes. Deuterostomes,
animals that develop the mouth secondarily to the anus, are further divided into chordates and
ambulacrarians. Echinoderms are basally branching deuterostomes and can, through comparison with
chordate groups, yield insights about the common deuterostome ancestor and the extent of homology
among derived deuterostome groups.
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The mouth opens at different locations on the body among deuterostome groups. In chordates,
the mouth opens at the border of the anterior neural ectoderm, while in ambulacrarians it opens at a
more posterior position (Christiaen et al., 2007). In addition, chordate mouths open on the BMPexpressing side of the dorsal-ventral axis, while ambulacrarian mouths open on the Nodal-expressing
side. There are two hypotheses for how this change in mouth location may have occurred. The first
hypothesis, proposed by Christiaen et al. (2007), states that the mouth changed location in derived
deuterostome groups, but the molecular regionalization of the embryo stayed the same (Fig. 2). They
hypothesize that the mouths of derived groups develop from the same gene regulatory networks found in
the common deuterostome ancestor, but moved to a different region. Thus, they hypothesize true
homology among deuterostome mouths. The second hypothesis, proposed by Kaji et al. (2016), states
that every relocation of the mouth indicates a new evolutionary event (Fig. 3). That is, mouth
development was reinvented independently several times in the deuterostome rather than co-opting preexisting molecular patterns from the common deuterostome ancestor, and that ambulacrarian,
cephalochordate, and vertebrate mouths are developmentally non-homologous. In sum, the contrasting
hypotheses proposed by Christiaen et al. (2007) and Kaji et al. (2016) ask the question: to what extent is
the mouth homologous among deuterostome groups?
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Fig. 2 The molecular landscape of the dorsal ectoderm may have been conserved between the
common deuterostome ancestor and the chordate ancestor (Christiaen et al., 2007)
It is thought that the mouth developed in a different position as the chordate ancestor evolved from the
common deuterostome ancestor. However, the regional expression of key dorsal ectodermal genes, such
as goosecoid (yellow), six3 (blue), dlx (purple), and pitx (pink), may have been preserved. It is possible
that the chordate ancestor, and the ancestors of other derived deuterostome groups, co-opted a preexisting ancestral molecular landscape to develop its mouth rather than invent an entirely new mouth. In
this way, deuterostome mouths may all be developed using similar signaling pathways, which may have
been present in the common ancestor, and morphogenetic events.
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Fig. 3 Mouth may have been reinvented for each derived deuterostome group (Kaji et al., 2016)
The change in mouth location among deuterostome groups may correspond with separate evolutionary
events described in dark grey dialog boxes. In this way, each derived deuterostome group may have used
different molecular and morphogenetic mechanisms to develop its mouth than in the common
deuterostome ancestor.
A better understanding of ambulacrarian mouths and their development is important to help
address the question posed by these two hypotheses. The sea urchin is a particularly powerful model
organism for studying mouth development because its embryos are optically clear, allowing easy and
effective imaging of cell movements involved with mouth development. The sea urchin is also a basally
branching non-chordate deuterostome, making it a useful tool for constructing hypotheses about the
common deuterostome ancestor and the extent of homology among deuterostomes (Fig. 1).
In this thesis, I compare three morphogenetic aspects of mouth development between the sea
urchin (Lytechinus variegatus), an ambulacrarian, and the frog (Xenopus laevis), a vertebrate. If the sea
urchin and the frog develop their mouths by similar mechanisms, then the hypothesis for extensive
homology among deuterostome mouths would be supported. Conversely, if sea urchins and frogs do not
appear to develop their mouths by similar mechanisms, then it is likely that vertebrates reinvented the
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mouth when they diverged from the common deuterostome ancestor (or, alternatively, sea urchins may
have reinvented the mouth from the common deuterostome ancestor). This would support the hypothesis
of at least three origins of the deuterostome mouth.
In this thesis, I ask whether sea urchin mouth morphogenesis employs any of the three
mechanisms known in the frog: 1) specification by inductive signaling from foregut endoderm, 2)
antagonism of Wnt signaling to permit basement membrane remodeling, and 3) apoptosis of overlying
ectoderm to perforate the mouth (Fig. 4). Normal mouth development in the frog depends on inductive
signaling from the endoderm (Dickinson and Sive, 2007). When the endoderm adjacent to the
stomodeum is removed from embryos, a small indentation forms at the stomodeum but no perforate
mouth develops (Dickinson and Sive, 2006). Normal mouth development in the frog also requires frzb-1
and crescent expression, Wnt signaling antagonists, at the mouth (Dickinson and Sive, 2009). These
genes are thought to regulate basement membrane dissolution and remodeling at the stomodeum in part
by directly regulating RNA expression of the basement membrane proteins laminin and fibronectin
(Dickinson and Sive, 2009). Perforation, or forming a mouth hole, is the last step of mouth development
in the frog and occurs by apoptosis at the stomodeum (Dickinson and Sive, 2006).
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Fig. 4 A descriptive model of mouth morphogenesis in Xenopus laevis (Dickinson and Sive, 2009)
In the frog, the mouth forms in an area where ectodermal and endodermal cells are in direct contact with
no intervening mesoderm. Adjacent cells of these two tissues express Wnt antagonists frzb-1 and
crescent and Wnt signaling is downregulated. As a result, the transcriptional levels of basement
membrane proteins are also downregulated and the basement membrane separating the ectoderm and
endoderm dissolve. The ectoderm then begins to thin via apoptosis and the stomodeum invaginates.
Finally, the mouth opens at the perforation step.
Very little is known about the morphogenetic mechanisms of mouth development in the sea
urchin. I aim to characterize mouth morphogenesis in sea urchins to not only contribute knowledge
about sea urchin development specifically, but also in the context of the evolutionary question of
homology among deuterostome mouths.
Sea urchin mouth development: An incomplete understanding
A molecular model for how the stomodeum is specified in the sea urchin has been proposed, but
remains incomplete (Fig 5). Stomodeal specification begins with the establishment of the dorsal-ventral
axis in the early embryo. Between fertilization and early cleavage, a maternally loaded redox gradient
and inductive signaling from the vegetal pole drives nodal expression selectively in the oral (ventral)
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ectoderm (Duboc et al., 2004; Li et al., 2013). Nodal appears to be a driver of not only oral ectoderm
patterning, but also stomodeal specification specifically. Nodal is thought to directly activate the
repressor not, which inhibits sip1 and ets4 at the stomodeum. Sip1 and ets4 repress the stomodeal gene
brachyury. When sip1 and ets4 expression is cleared from the stomodeum by the Nodal-activated not,
brachyury expression is turned on (Li et al., 2013). It is thought that in this way, Nodal drives the
activation of brachyury and other stomodeal-specific genes throughout the blastula stage. It has been
shown that if nodal expression is blocked, no mouth develops (Duboc et al., 2010).
Fig. 5 Sea urchin stomodeum-specific gene regulatory network (Davidson, 2015)
This gene regulatory network shows the proposed gene interactions necessary to specify the stomodeum,
a patch of oral ectoderm where the mouth will develop. Nodal (dark blue) is proposed to be the driver of
stomodeal specification as it promotes dowstream expression of brachyury (purple) and foxA (light
blue), two genes selectively expressed at the stomodeum. It is proposed that Nodal activates bra and
foxA indirectly by activating a repressor (not, not shown) of the repressors sip1 and ets4 (light grey).
An interesting question is to what extent stomodeal specification is autonomous in the sea
urchin. That is, does it require signaling from vegetal cells? It has been suggested that nodal expression
in the oral ectoderm depends on early inductive signaling from the vegetal pole (Duboc et al., 2004). If
the animal halves of embryos are isolated at the 8-cell stage, they entirely lack an ectoderm and no
mouth develops (Kominami et al., 2006; Wikramanayke et al., 1995). Kominami et al. (2006) showed
that if the third cleavage axis is shifted vegetally so that the animal half includes sub-equatorial
cytoplasm, isolated animal halves of embryos develop fully differentiated ectoderm, including a
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morphological mouth. Thus, after early cleavage, stomodeal specification may be an autonomous
process.
Very little is known about the morphogenetic details of mouth development in the sea urchin.
However, a descriptive model of the main morphogenetic steps of mouth development is summarized in
Figure 6. Toward the end of gastrulation, the stomodeum slightly invaginates into the blastocoel. With
guidance from filopodia at the top of the archenteron, the gut contacts the stomodeum (Hardin, 1990).
The basement membranes associated with the stomodeal epithelium and the tip of the foregut epithelium
then break down. The cells of the stomodeal and gut epithelia then rearrange and rebuild a shared
basement membrane as a single epithelium. Just how this fusion of these two epithelia occurs has not
been recorded. Lastly, the mouth perforates, or creates an opening, through cell rearrangement,
apoptosis, or both. Although the morphogenetic details of mouth development in the sea urchin are not
well-articulated, there is evidence that at least some morphogenetic events are autonomous to the
ectoderm (Kominami et al., 2006). However which or how many of these processes are tissueautonomous has not been studied.
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Fig. 6 The current understanding of sea urchin mouth morphogenesis
(A)The patch of oral ectoderm that will become the mouth, called the stomodeum (white), is specified
before it touches the gut (blue). (B) With guidance from filopodia (black lines in A), the gut contacts the
stomodeum. (C) Toward the end of gastrulation, the stomodeum slightly invaginates into the blastocoel
(inside of the embryo). (D) The basement membrane (green) between the oral ectoderm (magenta) and
gut endoderm dissolves and the two epithelia intercalate. (E) Lastly, the mouth perforates through an
unknown mechanism (cell migration or apoptosis). (F) To highlight the morphology of the mouth, a
control embryo was imaged on its side. The animal half of the embryo was marked with a green
fluorescent protein and the vegetal half was marked with a red fluorescent protein (artificially colored
magenta). The mouth is indicated with an arrow.
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The current study
In this thesis, I, with assistance from the McClay lab examined three specific morphogenetic
processes in sea urchin (Lytechinus variegatus) mouth development as described above to ask: 1) does
the mouth require induction from the gut to perforate (or vice versa), 2) does Wnt signaling regulate
basement membrane remodeling in the stomodeum, and 3) does apoptosis perforate the mouth? My
thesis is also motivated by a bigger-picture evolutionary developmental biology question: to what extent
is the mouth homologous among deuterostome groups? I approach this question by comparing these
three morphogenetic aspects of mouth development in the sea urchin (Lytechinus variegatus) and the
frog (Xenopus laevis). If the urchin and frog share developmental mechanisms of mouth development,
then the hypothesis of extensive homology among deuterostome mouths would be supported despite
their appearance in different embryonic regions.
I first asked if the sea urchin mouth requires an inductive signal from the gut endoderm or is
tissue-autonomous. As discussed above, there is evidence that some morphogenetic processes of sea
urchin mouth development occur tissue-autonomously. However, it has not been studied if basement
membrane remodeling in either the stomodeum or tip of the archenteron are autonomous processes. In
this thesis, I examined these two processes specifically. Two methods were employed to separate the gut
from the ectoderm and prevent signaling interactions between these tissues. The first method was to
induce exogastrulation by depriving embryos of calcium to interrupt normal invagination of the
archenteron (Okazaki, 1956). The second method was to remove archenterons by microsurgery and
culture separately isolated guts and gutless embryos. The embryos from both methods and the isolated
guts from the second method were immunostained for laminin, a basement membrane protein, to assay
basement membrane dissolution. If laminin breakdown was seen in any of these experimental embryos,
then tissue autonomy of basement membrane remodeling would be supported. Homology between sea
urchin and frog mouths would not be supported if basement membrane dissolution at the was found to
be autonomous processes.
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I then examined if late Wnt signaling regulates the basement membrane during sea urchin mouth
development. Early Wnt signaling is known to be critical in endomesoderm specification among all
animals. Blocking Wnt signaling early in a sea urchin embryo’s development causes failed
endomesoderm specification and no mouth forms (Duboc et al., 2004). However, little is known about
the effects of Wnt signaling in the context of mouth development. Embryos were treated with different
doses of C59, a drug that prevents secretion of Wnt ligands. These embryos were scored
morphologically and assayed for basement membrane remodeling using an anti-laminin antibody at the
pluteus larval stage, when control embryos had fully developed mouths. Homology between frog and
sea urchin mouths would not be supported if basement membrane proteins were upregulated and mouth
area decreased as the amount of Wnt antagonism increased.
Finally, I investigated if the sea urchin formed a mouth by apoptosis or cell migration, which has
not been previously studied. To test for apoptosis during mouth perforation, control embryos were
immunostained with an anti-caspase3 antibody, an executioner caspase involved in apoptosis. Live
imaging of control embryos was performed to look for evidence of cell migration during mouth
perforation. Lack of evidence of apoptosis at the ectoderm during sea urchin mouth perforation would
not support homology between frog and urchin mouths.
Methods
Lytechinus variegatus
Sea urchins (Lytechinus variegatus) were collected and shipped from Beaufort, North Carolina or from
Gulf Specimen Marine Labs in Florida during the winter months. Sea urchins were kept in tanks of
artificial seawater (ASW) at 22°C. New shipments of sea urchins arrived monthly.
Spawning and collection of gametes
Sea urchins were spawned by injecting 3 mL – 5 mL of 0.5 M potassium chloride (KCl) in at least two
spots in the soft tissue surrounding the mouth. Urchins were then shaken vigorously for 10 seconds. To
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collect eggs, the spawning urchin was placed on a 50 mL beaker filled with ASW. Unfertilized eggs
were only viable up to an hour after spawning. Sperm was collected by pipette and stored dry at 4°C for
up to 2 weeks.
Culture
Freshly spawned eggs were allowed to settle to the bottom of the 50 mL beaker. A sample of eggs was
examined under the microscope to ensure eggs were uniformly round with no spots (which indicates
immature eggs) or fertilization envelopes before fertilization. They were then washed three to four times
by decanting most of the water and replacing it with fresh artificial seawater. To activate sperm, 8 µL of
sperm were added to 1 mL of artificial seawater in a 1.5 mL tube and the tube was gently inverted.
Depending on the size of the culture, 0.5 mL to 1 mL of the diluted sperm and seawater mixture were
added to washed eggs. The mixture was pipetted up and down a few times to ensure the sperm was
evenly distributed around the eggs. After 2 minutes, a sample of eggs was examined under the
microscope for fertilization envelopes to ensure proper fertilization. Zygotes were then washed at least
two times before being added to a finger bowl filled halfway with artificial seawater. Depending on the
desired speed of development, the culture was incubated at temperatures ranging from 23°C (for faster
development) to 19°C (for slower development) (Fig 7). However, all times reported in the figures and
text of this thesis correspond with incubation at 23°C.
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Fig. 7 The speed of sea urchin (Lytechinus variegatus) embryonic development depends on
incubation temperature
The average time in hours post-fertilization (hpf) at which embryos reached certain developmental
stages was recorded at different incubation temperatures. Shaded boxes represent a lack of data. As
incubation temperature increases, the rate of development also increases. Each of the represented
incubation temperatures was used at some point during the data collection. However, the times reported
in the figures and text of this thesis correspond with an incubation temperature of 23°C.
Fixation
Equal volumes of embryos in ASW and 8% paraformaldehyde (PFA in 20mM EPPS) were added to a
tube. Embryos were kept at 4 degrees Celsius for at least 12 hours. Embryos were then washed at least
twice for 10 minutes with phosphate-buffered saline (PBS), removing most of the old solution with
every wash. Washed embryos were stored in methanol (MeOH) at -20°C.
Mounting embryos and live imaging
Glass slides (1.5mm thick) were coated with 1% protamine sulfate for 10 minutes and washed with
deionized (DI) water. Embryos were treated with a salt shock to remove embryos’ cilia, helping to
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immobilize embryos during live imaging. About 50 µL of 2x ASW were added to a well in a 9-well
glass dish. Using a mouth pipette, about 5 embryos were transferred to the 2x ASW. After 6 seconds,
750µL of 1x ASW were added to the well. Embryos were mounted in ASW containing parsley
extract/dillaoil (1:1000) to further inhibit movement, under a coverslip with modeling clay at each
corner. To prevent water evaporation during imaging, the edges of the coverslip were sealed with
VALAP (Vaseline, lanolin, paraffin 1:1:1) sealant.
Microscopy and image analysis
Embryos were imaged using a Zeiss Axio Imager upright microscope controlled by Zen software. The
time-lapse and Z-stack functions were used for live imaging. All images and time-lapse movies were
processed and analyzed using Fiji/ImageJ software.
Immunohistochemistry – Antibody staining
Between 5 and 10 embryos fixed in PFA/MeOH were transferred to a 9-well glass dish. Embryos were
washed 4 times with PBST (0.1% Triton-20 in PBS) for 5 minutes. In a 12-column Terasaki plate,
embryos were blocked in 8µL PBST-NGS (4% normal goat serum in PBST) for 1 hour at room
temperature. Embryos were then incubated in 8µL of diluted primary antibody in PBST-NGS overnight
at 4°C. Anti-laminin (rabbit IgG, Sigma) was diluted 1:250 and Endo1/5c7 (mouse IgG, monoclonal
(Wessel and McClay 1986) was diluted 1:100. Then, embryos were washed 3 times in 8 µL PBST for 5
minutes at room temperature. Embryos were placed briefly in 8 µL 1:5000 Hoescht (a nuclear marker) in
PBST and washed with 8 µL PBST for 5 minutes at room temperature. Embryos were incubated in 50%
glycerol in PBST for at least 10 minutes at room temperature before imaging. Embryos were imaged in
50% glycerol in PBST the same day.
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Microsurgery for gut-removal
Embryos at late gastrula, early prism, or early pluteus stage were mounted in a Kiehart chamber in
which the coverslips were treated with protamine sulfate to keep the embryos immobilized. A suction
pipet was inserted into the archenteron and suction was applied as the pipet was withdrawn from the
archenteron. In this way, the archenteron was inverted into the pipet. A glass needle then separated the
inverted archenteron from the remaining embryo with the cut made at or near the blastopore. The
archenteron was then transferred to a separate dish. Embryos and guts were cultured for 20 hours in
ASW at 23°C before fixation.
Inducing exogastrulation in embryos using calcium deficient sea water
Small cultures were prepared in a 6-well plastic dish by adding 9mL of calcium-free seawater to every
mL of embryo culture in ASW when embryos were at early blastula. Embryos were fixed at 26hpf and
40hpf.
Drug treatment with Wnt antagonist (C59 drug)
C59 is a cell-permeable drug that inhibits Wnt signaling by inhibiting porcupine (PORCN), a
transferase that adds palmitoleic acid to Wnt proteins. Palmityoylation by PORCN facilitates the
activation and cellular release of Wnt signal proteins. In this way, C59 drug treatment produces the same
effect as blocking the expression of all secreted Wnt genes (Cui et al., 2014).
In each well of a 6-well plastic dish, 5 mL of embryos in ASW were cultured. When embryos
were at mesenchyme blastula or early gastrula stage, different amounts of 1 mM C59 drug in dimethyl
sulfoxide (DMSO) (0.5 µL, 2.5 µL, 1 µL, or 5 µL) were directly added to the well and the plate gently
swirled for at least 10 seconds. Fertilization (no treatment added) and vehicle controls (1 µL DMSO)
were also included.
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Apoptosis assay: anti-cleaved caspase3 antibody
As a positive control assay, embryos at 25hpf were put in a petri dish and exposed to 10 minutes of UV
radiation using a gel box to induce DNA damage and apoptosis. Embryos were subsequently cultured at
room temperature for one hour before fixation. The hour of rest was intended to allow time for the
apoptosis signaling pathway to be activated. These UV-exposed embryos as well as control embryos at
25hpf were then fixed and analyzed using an anti-cleaved caspase3 antibody (rabbit IgG from Cell
Signaling Technologies product #9664) using the immunohistochemistry protocol above.
Results
Mouth morphogenesis occurs between 26hpf and 28hpf
Control embryos that were stained with an anti-laminin antibody and a nucleus marker provided a
time course of mouth development (Fig. 8). The basement membrane at the stomodeum and archenteron
began to dissolve at 26 hours post-fertilization (hpf). A perforate mouth was consistently observed
between 27hpf and 28hpf.
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Fig. 8 Basement membrane remodeling and dissolution occurs between 26 hpf and 28 hpf
Control embryos were fixed hourly and labeled with a basement membrane marker (anti-laminin
antibody, green), a mid- and hind-gut marker (Endo1 antibody, blue), and a nuclear stain (Hoechst,
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magenta). A side view and close-up oral view of embryos between 24 hours post-fertilization (hpf) and
27 hpf show that basement membrane begins to dissolve between 26 hpf and 28 hpf (white arrows). No
side view picture is included for the 26 hpf time-point. Scale bar represents 25 µm.
Mouth formation may not require contact between the stomodeum and endoderm
We sought to determine if mouth morphogenesis could occur tissue-autonomously or if it
requires signaling from the endoderm. The gut was removed by microsurgery from embryos at late
prism stage (at the end of gastrulation), and isolated in a separate dish. When imaged at 20 hours postsurgery (~40hpf), 67% of embryos showed little to no gut recovery and had mouths (Figure 9a).
Exogastrulation was also induced in embryos through calcium deprivation. At 26hpf, these embryos
showed no convincing evidence of mouth formation (Fig. 10b). At 40hpf, some exogastrulated embryos
formed what appeared to be an invagination of the ectoderm (Fig. 10d). However, it was unknown if
these invaginations had the stomodeal molecular identity. In addition, it was impossible to determine if
these structures were perforate like a mouth. Lastly, the phenotypes of exogastrulated embryos at 40hpf
were variable and only a small percentage of embryos showed this morphology.
Basement membrane breakdown at gut may occur without signaling from the ectoderm
To develop a functional through-gut, the top of the archenteron that touches the stomodeum must
form a hole. We investigated whether basement membrane breakdown at the gut and formation of a
hole require signaling from the ectoderm. Guts that were isolated from the embryo at the late prism stage
showed a wide range of phenotypes when examined 20 hours post-surgery (Fig. 9b). Some of these
isolated guts showed evidence of a laminin dissolution (Fig. 9b white asterisk). A majority of
exogastrulated embryos showed evidence of laminin breakdown at the gut (Fig. 10b-b’). However, there
was no evidence that the gut actually formed a hole (Fig. 10b’’).
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Fig. 9 A mouth develops after guts are removed and isolated from embryos
Guts were surgically removed from embryos at late pluteus stage and isolated in a separate dish.
Embryos were fixed about 24 hours post-surgery. Gutless embryos (A, z-stack) show little gut regrowth
(blue) compared to control embryos (A’). A perforate mouth is also seen in gutless embryos, suggesting
that the mouth is not formed by signal induction (laminin, green; Hoechst, magenta). (B) Isolated guts
showed a range of phenotypes from undifferentiated balls of endoderm to a phenotype that resembled a
tripartite gut. The identity of guts was confirmed by staining with a mid- and hind-gut marker
(Endo1/C57, not shown). It is possible, but unclear if these isolated guts formed a laminin hole (B, white
asterisk).
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Fig. 10 Exogastrulated guts may show laminin breakdown
Embryos were deprived of calcium in early development to induce exogastrulation (reverse growth of
the gut). (B) Although no mouth was conclusively seen in exogastrulated embryos, the basement
membrane on exogastrulated guts appeared to break down (B’, laminin, green). However, no hole
formed (B’’, Hoechst, magenta). (C, C’) A bottom view of an exogastrulated embryos shows that the
area of laminin breakdown was restricted to the side of the embryo (laminin, green; Hoechst, magenta).
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(D, D’’, z-stacks) Although the phenotype of exogastrulated embryos was fairly uniform at 26 hpf,
phenotypes at 40 hpf varied enormously and were difficult to interpret.
Inhibition of Wnt signaling decreases mouth size in a dose-dependent fashion
We sought to determine the role Wnt signaling plays in mouth morphogenesis. Treating embryos
with C59, a Wnt-inhibiting drug early in development greatly disrupted arm and gut growth and no
mouth formed (Fig. 11e). Treating embryos with C59 at late gastrula stage, after the endomesoderm was
specified, produced a phenotype almost identical to control embryos, but no mouth formed (Fig. 11e).
Treating embryos with different doses of C59 showed that as the drug dose increased, average mouth
size decreased from 265 microns (vehicle control) to 103 microns (1µM dose) (Fig. 11b). In addition, as
drug dose increased, the percent of embryos that developed mouths also decreased at doses greater than
0.5µM (Fig. 11c). At the highest dose of the Wnt-inhibiting drug, 5 µM, no embryos developed mouths
(Fig. 11a-d). In addition, as the concentration of C59 in drug treatments increased, the oral hoods of
embryos become narrower, less rectangular, and rounder (Fig. 11a).
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Fig. 11 Increased Wnt inhibition decreases sea urchin mouth size
Embryos were treated with the drug C59, which inhibits canonical Wnt signaling. (A-C) When added at
the mesenchyme blastula stage, as C59 dose increased, the mouth decreased in area (p = 1.65x10-8 by
ANOVA) and fewer embryos developed mouths at all. (D) At the highest dose of C59, no mouth forms.
(E) Treating embryos with the drug earlier in development interfered with endomesoderm specification,
as expected, and produced a more abnormal phenotype at the pluteus larval stage. When embryos were
treated at early gastrula, they developed a normal pluteus larva phenotype except they failed to develop
mouths.
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The sea urchin mouth does not appear to perforate by apoptosis
We sought to determine if the mouth forms a hole by cell migration or by programmed cell death.
We first attempted to observe mouth perforation through live imaging of embryos using differential
interference contrast (DIC) microscopy (Fig. 12). Cells at the stomodeum appear mesenchymal from
frame 1 to frame 50. From frame 75 to the final frame of the movie, the cells appear more cuboidal and
characteristic of epithelial cells. However, the exact moment of mouth perforation could not be observed
from this angle or z-plane. Control embryos stained with anti-caspase3 antibody did not show apoptosis
at the oral ectoderm (Fig. 13a-a’). I conclude that the lack of signal was not due to a faulty assay since
embryos that were exposed to 10 minutes of UV radiation did show high levels of caspase3 signal (Fig.
13b-b’).
Fig. 12 The mouth may perforate by cell migration
A control embryo was mounted on its side and live imaged using differential interference contrast (DIC)
microscopy. Five out of 103 frames are represented above. It is unclear from a single z-plane what
morphogenetic processes, such as those involved mouth perforation, are occurring. However, by the end
of the movie (frame 103), what appears to be a cuboidal epithelium appears (indicated by a black
bracket). It is likely this is the mouth.
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Fig. 13 The sea urchin mouth does not appear to perforate via apoptosis
Embryos were stained with a DAPI nuclear stain (blue) and an anti-caspase3 antibody, a marker of
apoptosis (red). A surface view (A, B) and cross-sectional view (A’, B’) of embryos are shown. (A, A’).
Control embryos show no apoptosis in the ectoderm where the mouth perforates. (B, B’) For a positive
control assay, embryos were treated with 10 minutes of UV radiation to promote apoptosis.
Discussion
This thesis had two main motivations. First, I aimed to delve into the details of mouth
morphogenesis with hope to add to the current limited model of sea urchin mouth development. Second,
by comparing three aspects of mouth development in frogs (Xenopus laevis) and in sea urchins
(Lytechinus variegatus), I aimed to provide insight about the extent of homology among deuterostome
mouths. The three specific morphogenetic aspects of mouth development we investigated were: if the
mouth can form without signaling from the archenteron, if Wnt signaling regulates basement membrane
remodeling during mouth development, and if apoptosis perforates the mouth. If the urchin and frog do
not share many mechanisms of mouth morphogenesis, then homology among deuterostome mouths
would be supported. In this case, it is instead likely that different deuterostome groups independently
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reinvented the mouth at different locations on the embryonic body plan. The results addressing each of
the three aspects of mouth formation are discussed below.
Mouth formation and basement membrane breakdown at the gut may be tissue-autonomous
A majority of embryos that had their gut removed at late pluteus stage developed a perforate
mouth, suggesting that mouth formation does not depend on signaling with the gut. However, some of
the guts in these embryos were removed after the endoderm and ectoderm had already met, diminishing
the credibility of the conclusion that the mouth perforates autonomously. We tried to account for this
problem by removing guts at an earlier time point, the late gastrula stage, but by the time the mouth
perforated, most of the guts re-invaginated and re-established themselves in these embryos.
We therefore turned to a second approach to address this question. Embryos were induced to
exogastrulate through treatment with calcium-deficient seawater. Imaging these embryos proved
difficult because exogastrulation through this technique also seemed to radialize the ectoderm. It was
impossible to discern the location of the stomodeum and to orient the embryo on the slide accordingly.
We conclude with caution that mouth formation can occur without signaling from the endoderm.
This contrasts with mouth development in frogs, which requires inductive signaling from the gut
endoderm (Dickinson and Sive, 2007). In this way, these results do not support homology between frog
and sea urchin mouths.
We also examined if basement membrane remodeling at the gut, a necessary step for developing
a functional through-gut, could also occur tissue-autonomously. Guts that were surgically removed from
embryos and isolated in a separate dish were too fragile and damaged to yield useful results. Analysis of
the exogastrulated guts proved more illuminating. Exogastrulated guts tended to show laminin
breakdown, suggesting that dissolving the basement membrane at the tip of the archenteron does not
depend on contact with the ectoderm.
If the basement membrane can dissolve tissue-autonomously at the ectoderm during mouth
formation and at the gut during through-gut formation, what is the signal that controls these processes?
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Future studies could investigate this question by first performing in situ hybridization for foxA, a gene
that is normally expressed throughout the gut and stomodeum, on gutless or exogastrulated embryos. If
foxA expression remains in both the gut and stomodeum in these embryos, then it may be a candidate for
controlling basement membrane dissolution. Other genes that show expression in the gut and
stomodeum, such as brachyury, could also be strong candidates. FoxA could then be knocked down in
embryos using an anti-sense morpholino. To selectively knockdown foxA in the gut, the animal half of a
control embryo could be combined with the vegetal half of an embryo injected with a foxA morpholino.
The reciprocal could be done to knockdown foxA in the stomodeum selectively. These perturbation
experiments would be particularly insightful for determining what gene or molecule provides the signal
for the basement membrane dissolution necessary for sea urchin mouth development.
Wnt signaling may be required for sea urchin mouth development
Treating embryos with the Wnt-signal inhibiting drug, C59, at mesenchyme blastula decreased
mouth size in a dose-dependent fashion. This contrasts from the frog where antagonism of Wnt signaling
by frzb-1 and crescent promote larger mouths (Dickinson and Sive 2009). In this way, Wnt signaling
appears to play the opposite role in sea urchin mouth development than in frog mouth development. This
provides further support against homology among deuterostome mouths.
It is clear from my results that Wnt-signaling inhibition plays an important role in mouth
development. It is important, however, to validate these results by over-expressing Wnt-signaling,
through a drug for example, and recording if mouth size increases as expected. In addition, the
mechanisms by which Wnt-signaling regulates mouth size cannot be addressed by the results of this
thesis. A logical first step is to investigate if Wnt-signaling at the stomodeum regulates basement
membrane protein transcription as it does in the frog. To address this question, in situ hybridization
against the RNA of basement membrane markers could be performed on embryos treated with C59 drug.
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Urchin mouth does not likely perforate via programmed cell death
Embryos that were stained with an anti-caspase3 antibody showed no apoptosis at the oral
ectoderm, suggesting that the sea urchin mouth does not perforate via apoptosis. However, having only
analyzed a single time point, I am skeptical to entirely disregard the possibility that apoptosis perforates
the urchin mouth. Analyzing additional time points with an anti-caspase3 antibody are necessary to
strengthen this conclusion. In addition, conclusions should be verified through other methods for
detecting apoptosis such as TUNEL staining, which marks nicks in fragmenting DNA with labeled
nucleotides. I believe Syto12 would be the most useful method for detecting apoptosis for future studies,
since it can be used on living cells and could complement live imaging of embryos during mouth
development. Whether the sea urchin mouth perforates by cell rearrangement must be further explored.
Live imaging of embryos during mouth perforation is the most direct way of addressing this question.
The frog mouth is thought to perforate by apoptosis, so the results of this experiment provide even more
evidence against homology between frog and sea urchin mouths (Dickinson and Sive 2009).
Conclusion
Regarding the evolutionary biology question that motivated this thesis, these data do not support
true homology between frog and sea urchin mouths. Therefore, it is likely that there are at least three
origins of the deuterostome mouth. From a pure developmental biology perspective, my thesis serves as
a starting point for adding more detail to the current model of sea urchin mouth development. The future
experiments already discussed would be helpful in determining the mechanisms of specific
morphogenetic events. However, live imaging is imperative for truly understanding mysterious
developmental processes like the “fusion step” of sea urchin mouth development. It is unknown how
exactly the archenteron and oral ectoderm epithelia “fuse” and reorganize to construct a single epithelia.
A powerful technique for visualizing the “fusion step” is to image fluorescent chimeric embryos
(Fig. 14). In these embryos, a green fluorescently marked animal half of an embryo is combined with a
red fluorescently marked vegetal half. As the embryo develops, the ectoderm will be marked green and
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the gut will be marked red. Live imaging these embryos could clearly document the cell movements
involved with the fusion of these two tissue layers that is imperative for sea urchin mouth development.
Fig. 14 Transplants with GFP and RFP-injected embryos can document the formation of a new
mouth epithelia from gut and ectoderm cells
Zygotes were injected directly after fertilization with either a green fluorescent protein (GFP) or red
fluorescent protein (RFP). At the 32-cell stage, the animal half from one embryo injected with one color
was combined with the vegetal half of an embryo injected with the other color. As the chimeric embryo
develops, all endoderm and mesoderm will be one color (red in the figure above) and the ectoderm will
be another color (green in the figure above). The image on the left shows a surface view of the embryo
with ectodermal cell membranes clearly outlined in green. On the right, the same embryo is shown
further down the z-axis.
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Acknowledgements
This research was funded by Sigma Xi Grants-in-aid of Research. I thank Allison Edgar for her
guidance, detailed feedback, and for constantly challenging me. Thank you also to Dr. David McClay
for his guidance, unwavering support, and surgical expertise. Thank you to Andrew George for his
technical support with live imaging, and image processing. Lastly, thanks to Leslie, Esther, and
Raymond for making the lab such a positive, friendly place.
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