Molecular and Morphological Characterizationof Segmentation in Artemia franciscana
Item Type text; Electronic Dissertation
Authors Blachuta, Beata J
Publisher The University of Arizona.
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MOLECULAR AND MORPHOLOGICAL CHARACTERIZATION OF SEGMENTATION IN ARTEMIA FRANCISCANA
By
Beata J. Blachuta
_____________________
A Dissertation Submitted to the Faculty of the
DEPARTMENT OF MOLECULAR AND CELLULAR BIOLOGY
In Partial Fulfillment of the Requirements For the Degree of
DOCTOR OF PHILOSOPHY
In the Graduate College
THE UNIVERSITY OF ARIZONA 2009
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THE UNIVERSITY OF ARIZONA GRADUATE COLLEGE
As members of the Dissertation Committee, we certify that we have read the dissertation prepared by Beata J. Blachuta entitled Molecular and Morphological Characterization of Segmentation in Artemia franciscana and recommend that it be accepted as fulfilling the dissertation requirement for the Degree of Doctor of Philosophy ____________________________________________________________Date: 12/18/08 Dr. Lisa Nagy ____________________________________________________________Date: 12/18/08 Dr. Gail Burd ____________________________________________________________Date: 12/18/08 Dr. Ted Weinert Final approval and acceptance of this dissertation is contingent upon the candidate's submission of the final copies of the dissertation to the Graduate College. I hereby certify that I have read this dissertation prepared under my direction and recommend that it be accepted as fulfilling the dissertation requirement. ____________________________________________________________Date: 12/05/08 Dissertation Director: Dr. Lisa Nagy
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STATEMENT BY AUTHOR This dissertation has been submitted in partial fulfillment of requirements for an advanced degree at the University of Arizona and is deposited in the University Library to be made available to borrowers under rules of the Library. Brief quotations from this dissertation are allowable without special permission, provided that accurate acknowledgment of source is made. Requests for permission for extended quotation from or reproduction of this manuscript in whole or in part may be granted by the head of the major department or the Dean of the Graduate College when in his or her judgment the proposed use of the material is in the interests of scholarship. In all other instances, however, permission must be obtained from the author.
SIGNED: Beata J. Blachuta
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ACKNOWLEDGEMENTS I am thrilled to put this chapter of my life behind me. However, I can not close the book on my time in Tucson without thanking the wonderful people that kept me sane while I struggled and fought ‘the great battle’. I am eternally grateful to the two most wonderful professors with whom I had the pleasure to teach. Both Dr. Thomas Lindell and Dr. Jonathan Flax were both a pleasure to work with. I enjoyed our many wonderful philosophical conversations, and will be forever grateful for the constant advice and moral support. You were my adoptive mentors and kindred spirits. Thank you. If this chapter has a cheering section, it surely is led by Barb Johnson. Thank you Barb for your constant support – financial, logistical, and most importantly, emotional. Your unfailing support and kindness kept me going in the darkest moments. Thanks to the lab folk who I had the pleasure of working with, namely Will Sewell, Maey Gharbiah, Bob Reed, Jessica Crance, Cristen Kern Hays, Jessy Wandelt, Matthew Terry, Ayaki Nakamoto, Julia Bowsher. You made me laugh, took my stuff out of the incubator when necessary, and were great lunch and coffee buddies. Most importantly, you were my mentors, my advisors and my scientific peers all rolled into one wacky package. I leave Tucson with a slew of new friends I will treasure forever. I will never forget our time together. Thanks to Jessica Crance, Drew Erickson, Lynda Hu-Donie, Ben Donie, Suzy Kim, Matt Knatz, Meghan McChesney, Jennifer Morris, Juliette Moore, William Sewell, Rebecca Spokony, Kelley Stanko, Susanne Stringfield, Andi Wardinsky, and Nicole Washington. You were a great support system and I treasure our friendships. I also thank Lori Pro for her friendship and for opening her doors to me in time of need. Thank you for taking in the strays, we enjoyed our time with you immensely. I was fortunate to have a Tucson family – the Garners. Thanks to Norm, Krista, Karly, and Karrin. You were the best adoptive family a girl can have. Thanks for all the great times, for feeding me, for giving me shelter, for giving me dogs, and for being the most supportive and coolest people ever. I cannot believe how lucky I was to have met you all. Finally, I would like to thank my family. My sisters, Eva and Anna have always been my musketeers – one for all and all for one. And as far as supportive parents go, I definitely got won the lottery. My parents, Danuta and Miroslav have not only supported me financially through this endeavor, but have also always encouraged me to keep fighting until the battle ends. So as I finally come to the end of this journey, I thank you for your unfailing support and dedicate this thesis to both of you.
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TABLE OF CONTENTS LIST OF FIGURES AND TABLES ……..………………………………………………6 ABSTRACT.........................................................................................................................7 CHAPTER 1: INTRODUCTION........................................................................................8
Overview..................................................................................................................8 What is a segment? ...………...…………………………………………………...8 Evolution of Segmentation………………………………………………………..9 Chordate Segmentation…......................................................................................11
Annelid Segmentation................................................................... ........................36 Concluding Remarks and Rationale for Study ......................................................38 CHAPTER 2: GAMMA SECRETASE INHIBITION ARRESTS SEGMENTATION IN THE BRANCHIOPOD CRUSTACEANS THAMNOCEPHALUS PLATYURUS AND ARTEMIA FRANCISCANA …………….………………………………………….…….46 Abstract …….…………………………………………………..…………..……46 Introduction …………………………………………..…………………….....…47 Methods....……………………..……………………………………….....……...51 Results……….………………………..……………………………………....….53 Discussion……………………………..………………….………………...……58 CHAPTER 3: SUMMARY OF CONCLUSIONS AND FUTURE DIRECTIONS ...….80 APPENDIX A: MORPHOLOGY OF SEGMENTATION………………..……………84 APPENDIX B: MORPHOLOGY OF DAPT MUTANTS ………………………..…….91 APPENDIX C: PARTIAL SEQUENCE OF ARTEMIA NOTCH…..…………………...94 APPENDIX D: WINDOW TREATMENTS OF ARTEMIA LARVAE WITH DAPT.....97 WORKS CITED....………..…..……………………………………..…………………101
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LIST OF FIGURES AND TABLES FIGURE: 1.1 Hypotheses for the evolution of segmentation………………………………40 1.2 Artemia development and body plan overview…………………………….. 42 1.3 Summary of the Delta/Notch signaling pathway ……………………………44 2.1 Artemia development ……………………………………………….……….68 2.2 Timing of the addition of Artemia segments……………………….………..70 2.3 DAPT affects Artemia segmentation in a dose dependent manner…….…... 72 2.4 DApT treatment does not affect larval growth………………………….…...74 2.5 Ubx/abdA expression in the posterior growth zone of DAPT treated larvae..76 A1.1 Newly hatched larva has no evidence of trunk segments …………………87 A1.2 Segment five and six morphology ………………………………………...89 A2.1 Cellular organization in the posterior of DAPT treated larvae ……………92 A3.2 Partial Artemia Notch amino acid sequence ………………………………95 A4.1 12 hour DAPT treatment window at 12-24 hours after hatching …………98 TABLE:
2.1 Loss of Notch signaling on segmentation in vertebrates and arthropods…...78
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ABSTRACT
In the animal kingdom, only the annelids, arthropods and chordates are segmented.
Whether the common bilateran ancestor to these three phyla was segmented, remains
debated. One way to address the origins of the evolution of segmentation is to compare
the molecular mechanisms underlying this complex process between the phyla and across
each phylum. This thesis first examines what we already know about segmentation in
each of the three phyla, and compares the models of segmentation in each phylum as well
as between the three. Then, the role of γ-secretase mediated signaling in segmentation
was examined in the branchiopod crustacean, Artemia franciscana. These findings were
further compared to another crustacean Thamnocephalus platyurus. Both of these species
develop their thoracic segments sequentially from anterior to posterior, and exposure to a
γ-secretase inhibitor slows segmentation in a dose dependent manner, but does not affect
the overall growth. My results suggest that Delta/Notch signaling is an essential for
segment patterning in these two species, although it may not function as a molecular
oscillator, as is the case in vertebrates. Similar findings in other arthropods suggest that
the role of Notch in segmentation is not as unique to vertebrates as once thought. Finding
such similarities in the molecular pathways that pattern segments across segmented phyla
suggests that the Urbilaterian may have indeed been segmented.
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CHAPTER 1: INTRODUCTION
Overview
segmented organisms make up about a third of the described phyla (Nielsen, 2001), They
are very diverse and successful in a large variety of habitats. The way organisms develop
segments, or functional body units, is as interesting as it is complex. This dynamic and
complex process requires a high degree of molecular coordination, such as specific
temporal and spatial regulation of gene expression, and targeted degradation of regulatory
proteins. As researchers sample a wide variety of species and compare how segments are
patterned, they can begin to ask questions about the evolutionary origins of segmentation,
and whether it speaks to the relatedness of all segmented organisms.
What is a segment?
Simply, a segment is one of a number of repetitive units into which something can be
divided. Animal segments are a series of repeated structures along the anterior-posterior
axis of the body that share several morphological and functional characteristics (see
Scholtz, 2002 for review). Units that define true segments are comprised of both
ectoderm and mesoderm, and include muscles, nerves, coeloms (mesodermal hollow
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spaces), blood vessels, excretory glands, and a pair of appendages (Scholtz, 2002; Seaver,
2003; Tautz, 2004; Wilmer, 1990). Although there are many examples of animals with
serial repetition, such as flatworms with serially repeated gut structures and nematodes
with cuticular rings, true segments are unique to only annelids, arthropods and chordates,
each part of a different clade of bilaterans (Figure 1.1A) (Balavoine, 1998; De Robertis
and Sasai, 1996; Valentine and Collins, 2000).
Evolution of Segmentation
We do not know whether the last common bilateran ancestor to the annelids, arthropods
and chordates, known as the Urbilaterian, was segmented (De Robertis and Sasai, 1996;
Valentine and Collins, 2000). Based on the fossil record, the Urbilaterian is thought to
have been worm-like (Valentine, 1994). If this vermiform animal was segmented, then
the process of segmentation may have evolved only once, giving rise to segmentation in
all three phyla (Figure 1.1A) (Kimmel, 1996). This plausible hypothesis was the basis
for grouping the annelids, arthropods and chordates as closely related clades, until the
protostome-deuterostome distinction asserted that most bilaterans are more related to
either the arthropods/annelid group or the chordate group, than these groups were related
to each other (Grobben, 1908). This separation of the arthropods and annelids from the
chordates, led to the belief that segmentation may have evolved twice and the
Urbilaterian may not have been segmented (Figure 1.1A) (Brusca, 1990; Clark, 1981). In
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accordance with this view, segmentation was used to unite the annelids and arthropods in
a clade, which excluded unsegmented phyla (Eernisse, 1992). However, both molecular
and morphological data suggest that arthropods and annelids are more closely related to
unsegmented phyla than they are to each other (Adoutte et al., 1999; de Rosa et al., 1999;
Eernisse, 1992). This finding supports the hypothesis that segmentation evolved three
separate times during the course of evolution (Figure 1.1A) (Wilmer, 1990).
Understanding the molecular regulation that governs the patterning of segments, and
identifying parallels, or the lack thereof, among these three phyla can be used to
distinguish between the three models for the evolutionary origins of the segmentation
process. However, comparisons to regulatory networks made between species have to
keep in mind the potential qualification that after segmentation evolved, the program was
likely modified differently in different species, without changing the outcome of a
segmented organism (Salazar-Ciudad et al., 2001). As such, some differences are likely,
and the degree to which the same molecular program is similar between species becomes
significant.
Perhaps the most well known example of how useful molecular information can be in
our understanding of whether certain structures are related across phyla is that of eye
development. Vertebrate and insect eyes were not though of as homologous structures
until the discovery that eyeless and pax6 genes program eye development in both flies
and mice (Quiring et al., 1994). Indeed, if we find that common pathways govern
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segmentation in annelids, arthropods and chordates, then it is likely that the Urbilaterian
was segmented and segmentation evolved only once. However, it is also possible that we
find similarities in the molecular program for segmentation between the arthropods and
annelids, and a completely different program in the chordates, supporting the hypothesis
that segmentation evolved independently in protostomes and deuterostomes. Finally,
finding mostly differences among the regulatory networks that pattern segments in
arthropods, annelids and chordates, lends itself to the possibility that segmentation
evolved separately in these three phyla (reviewed in Davis and Patel, 1999).
Most studies aimed at understanding regulatory networks governing segmentation focus
on a single species. Since different species are more amenable to different experimental
protocols than others, information obtained from different species is put together to
obtain a general model of the process in all organisms in the same phylum. This is
certainly the case in chordates, where understanding of the regulation of segmentation is
much richer when information about the process from studies in chick, mouse and
zebrafish is combined into one model.
Chordate Segmentation
In the vertebrates, segments along the anterior-posterior axis of the body develop as
blocks of tissues called somites. Somites form from the pre-somitic mesoderm (PSM), on
either side of the dorsal midline of the animal. Somites form at very regular time
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intervals in an anterior to posterior direction, suggesting the existence of an internal clock
which regulated this process (Dubrulle and Pourquie, 2004a; Pourquie, 2001; Pourquie,
2003). The ‘clock and wavefront model’ is the most widely accepted model explaining
the periodic nature of vertebrate segmentation (Cooke and Zeeman, 1976). The ‘clock’
component is a molecular oscillator which consists of waves of gene expression along the
PSM from posterior to anterior. Each wave is initiated in the posterior of the PSM,
progresses anterior, and is narrowed to a band that marks the location of the next somite
to form (Palmeirim et al., 1997). The ‘wavefront’ component of the model is the anterior
to posterior gradient of FGF expression and the posterior to anterior gradient of Retinoic
Acid, which together control the spacing mechanism of somite boundaries (Delfini et al.,
2005; Dubrulle et al., 2001; Dubrulle and Pourquie, 2004b; Goldbeter et al., 2007). The
next two sections review the mechanisms by which the ‘wavefront’ and ‘clock’ models
are thought to establish segment boundaries.
The’ Wavefront’ Component of Vertebrate Segmentation
The ‘wavefront’ component of vertebrate segmentation refers to the polarization of the
PSM, where molecular gradients establish positional information along the A-P axis
(Dubrulle et al., 2001; Maroto et al., 2005; Palmeirim et al., 1997). Highest in the
posterior of the PSM, a gradient of FGF creates a permissive environment for oscillation
of the downstream targets of Notch/Wnt signaling (Dubrulle et al., 2001; Sawada et al.,
2001). As the axis elongates and cells move anteriorly, they experience gradually
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decreasing levels of FGF levels along the portion of the PSM where segments are being
patterned, called the determination front. They ultimately reach a region along the PSM
where FGF levels drop below threshold and the cell can no longer respond to the
oscillating signals (see (Dubrulle and Pourquie, 2004a) for review). The number of cells
in the determination front during the length of one cycle of oscillations thus regulates the
size of each segment. Recent studies suggest that Wnt signaling also acts to establish a
posterior to anterior gradient in the PSM (Aulehla et al., 2003; Aulehla et al., 2008).
Furthermore, the PSM is also polarized from anterior to posterior by a gradient of retinoic
acid (RA), which acts to arrest oscillations in the anterior portion of the PSM, stopping
the signal at the location where the next segment will form (Dubrulle et al., 2001;
Dubrulle and Pourquie, 2004b; Sawada et al., 2001).
The ‘Clock’ Component of Vertebrate Segmentation
The first molecular evidence for the theoretical model of a segmentation clock was the
discovery in chick that the mRNA expression of the basic helix-loop-helix (bHLH) gene,
c-Hairy1 oscillated at regular intervals along the PSM, and the timing of each oscillation
corresponded to the length of time necessary for the addition of one somite (Palmeirim et
al., 1997). Since then, other hairy/ Enhancer of split related bHLH genes have been
found to cycle in chick, mouse and zebrafish (Bessho et al., 2003; Holley et al., 2000;
Oates and Ho, 2002) suggesting that the role these genes play in the timing of
somitogenesis is conserved in vertebrates. These molecules cycle within individual PSM
cells, and these single cell oscillations are then coordinated between neighboring cells to
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form a dynamic wave-like domain of expression (Maroto et al., 2005; Palmeirim et al.,
1997). As a broad wave of gene expression moves anterior, it narrows, responding to the
FGF and Retinoic Acid gradients, until a band of expression is formed just posterior to
the already patterned somite and signals the formation of the next somite (Dubrulle et al.,
2001; Maroto et al., 2005; Palmeirim et al., 1997).
Hairy is a downstream target of the Delta/Notch signaling pathway (see Figure 1.3 for a
summary of the Delta/Notch signaling pathway), which has been shown to be required
for proper segment formation in vertebrates (Conlon et al., 1995; Hrabé de Angelis, 1997;
Huppert et al., 2005). However, the mechanism by which Notch and Delta regulate hairy
oscillation may vary between species. In zebrafish, deltaC expression is cyclic;
suggesting that this Notch ligand is regulated hairy expression, leading to Notch
mediated signaling in the neighboring cell. The activation of Notch then leads to the
expression of hairy, and subsequently deltaC in that cell, hence causing a wave of hairy
expression from once cell to the next (Jiang et al., 2000; Oates et al., 2005).
Directionality of hairy oscillation is ensured by the fact that hairy inhibits its own
expression, whereby cells that expressed hairy quickly shut it down and can not express
more until it is degraded, limiting the signal only to neighboring cells that have not yet
expressed the gene (Lewis, 2003; Oates and Ho, 2002). In chick on the other hand, there
is no evidence of oscillation of Notch or Delta genes. Furthermore, the only evidence for
oscillation of these genes in mouse is the fact that Delta-like 1 oscillations were detected
with a riboprobe to the intronic region of the gene. (Maruhashi et al., 2005; Shifley et al.,
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2008). However, in both of these systems the expression of the glycosyl-transferase
lunatic fringe, a modulator of Notch signaling, has been shown to oscillate (Forsberg et
al., 1998; McGrew et al., 1998; Morales et al., 2002; Prince et al., 2001). No oscillating
lunatic fringe homologue has been identified in zebrafish. Interestingly, loss of Notch
signaling in vertebrates does not result in the loss of segments, instead affecting the
symmetry of the somite pairs and the integrity of segment borders between adjacent
somites along the anterior-posterior axis (Conlon et al., 1995; Takke and Campos-Ortega,
1999). Taken together, these data suggest that even though components of the Notch
pathway oscillate and are therefore part of the ‘clock’ program, the fact that these
oscillations can be maintained independently of Notch activity and that a loss of Notch
activity does not result in a loss of segments means that Notch is not the pacemaker of the
clock it was once thought to be (Holley et al., 2002; Jouve et al., 2000; Morales et al.,
2002). Other hypotheses regarding the possible roles of Notch signaling in the PSM
include: (1) establishment of boundaries between adjacent somites (Barrantes et al., 1999;
Saga, 2007; Takahashi et al., 2000), and (2) mediating communication between
neighboring cells to synchronize oscillations (Horikawa et al., 2006; Jiang et al., 2000)
(Ozbudak and Lewis, 2008).
In addition to components of the Notch pathway, members of the Wnt pathway also
oscillate in mouse (Aulehla et al., 2003; Ishikawa et al., 2004). These Wnt pathway
oscillations are out of phase with and slightly ahead of the Notch pathway oscillators.
Since disruption of the Wnt signaling leads to a disruption of oscillating Notch pathway
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components, and Wnt pathway targets have been shown to oscillate even when Notch
signaling is impaired, the Wnt pathway likely acts upstream of the Notch pathway
(Aulehla et al., 2003; Satoh et al., 2006). Furthermore, a microarray analysis of the PSM
transcriptome in mouse revealed a large network of cyclic genes in the Notch, Wnt and
also FGF signaling pathways (Dequeant et al, 2006). FGF pathway oscillators are in
synch with Notch pathway oscillators, and therefore slightly behind oscillations of Wnt
pathway components (Dequeant et al, 2006). Mutations that disrupt the Wnt pathway
stop oscillations of FGF signaling molecules (Dunty et al., 2008). However, in this
mutants, Axin2, a member of the Wnt signaling pathway, and members of the Notch
signaling pathway continue to oscillate, suggesting that Wnt signaling may play a
permissive rather than instructive role in the segmentation clock (Aulehla et al., 2008;
Dunty et al., 2008). Since the components of FGF signaling that have been shown to
oscillate are predominantly negative regulators of the FGF pathway, it has been
postulated that this signaling pathway may be involved in driving the molecular clock
(Dale et al., 2006; Dequeant et al., 2006; Niwa et al., 2007).
In sum, the current model of the molecular clock regulating vertebrate segmentation
segmentation relies on a network of molecular signals from three signaling pathways
(Dequeant et al, 2006) Although there are differences between species, proper timing
and formation of trunk segments involves oscillatory signaling of the Notch, Wnt, and
FGF pathways. Since mutations in components of these pathways do not result in a
complete arrest in segmentation, it is likely that these pathways compensate for each
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other offering robustness to the process of segmentation (Dequeant et al., 2006; Dequeant
and Pourquie, 2008; Riedel-Kruse et al., 2007). Though these ‘failsafe’ mechanisms
allow for a greater degree of reliability for proper segment development, the underlying
complexity has made deciphering the roles of all of the components of the molecular
clock difficult (Dequeant et al, 2006).
Segment Counting
The number of segments that an organism forms is species specific but variable among
vertebrates (Richardson et al., 1998). If all vertebrates use a similar clock-wavefront
mechanism to pattern their segments, then how can there be differences in segment
number between species? The ‘wavefront’ of FGF and Wnt gradients, which provides a
permissive determination front for segmentation, travels caudally as segments are added
and the axis elongates (Aulehla et al., 2003; Sawada et al., 2001). However, as the final
segments are added, the PSM shrinks due to a gradual extinction of the signals that
maintain the permissive nature of these cells to the segmentation process (Cambray and
Wilson, 2007). In chick and mouse embryos, this shrinking process is aided by extensive
cell death in the tail bud, and provides a spatial boundary for the addition of further
segments (Sanders et al, 1986). Interestingly, a recent study of segmentation in snakes,
which have a greater number of segments than the chick, mouse or zebrafish, showed that
the rate at which somites are added does not account for the larger number of segments
(Gomez et al., 2008). However, the average cell generation time, and more significantly,
the development rate of the snake embryo were much higher (Gomez et al, 2008). Taken
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together, these data show that the segmentation clock is faster over developmental time in
snakes than in the other vertebrates (Gomez et al, 2008).
Since the segmentation clock is comprised of a complex network of molecular regulators
that act in concert to pattern both the spatial and temporal addition of segments, it is not
surprising that this process is highly conserved among vertebrates. The species specific
variability therefore depends on changes of other processes associated with segmentation,
such as the overall rate of development in the snake (Gomez et al, 2008), variability of
which may be under the control of differential expression of just a few growth factors,
and thus easily manipulated during the course of evolution. With this degree of
conservation in the vertebrate segmentation program, a large amount of similarity to this
molecular program is expected in the signaling pathways that govern segmentation in
arthropods and annelids if the Urbilaterian was indeed segmented.
Arthropod Segmentation
Drosophila Segmentation
The fruit fly, Drosophila melanogaster, has served as the primary model for arthropod
segmentation since the 1980s, when many of the transcription factors involved in the
regulatory cascade that patterns segments were identified in through a genetic screen
(Nusslein-Volhard et al., 1980; Nusslein-Volhard and Wieschaus, 1980). The mechanism
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of segmentation in this insect still remains the best described arthropod segmentation
program. This hierarchic gene cascade systematically subdivides the Drosophila embryo
into increasingly smaller domains, until each unit is the width of one embryonic segment
(parasegment) along the anterior-posterior axis of the embryo (Deutsch, 2004). This
cascade begins with maternally provided coordinate gene products, including bicoid and
nanos, which establish the anterior and posterior poles of the embryo, respectively
(Driever et al., 1989; Driever and Nusslein-Volhard, 1989; Wang and Lehmann, 1991).
These transcription factors diffuse to form gradients along the embryo, and their
combined regulatory activity (both positive and negative) regulates the expression of the
gap genes, including tailless, orthodenticle, giant, Kruppel and knirps (Eldon and
Pirrotta, 1991; Hader et al., 1998; Knipple et al., 1985; Lehmann, 1984; Liaw and
Lengyel, 1993; Nauber et al., 1988; Wieschaus et al., 1984). Gap genes are expressed in
several segment-wide bands and act to not only narrow their expression domains, but also
coordinate the expression of pair-rule genes (Carroll and Scott, 1986; Frigerio et al.,
1986; Gergen and Butler, 1988; Goto et al., 1989; Hafen et al., 1984; Harding et al.,
1989; Howard et al., 1988; Kilchherr, 1986). The pair-rules are expressed in seven
stripes along the anterior-posterior axis, and are the first genes in the cascade to delineate
segment-wide boundaries by either the presence or absence of pair-rule gene expression.
The primary pair-rule genes, including hairy, runt and even-skipped, are expressed in
direct response to gap and maternal coordinate gene signaling (Pankratz, 1993). These
genes in turn help to define the expression domains of the secondary pair-rules including
fushi tarazu and paired (Pankratz, 1993). The pair rule genes then act to activate the
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segment polarity genes engrailed and wingless, which give anterior and posterior identity
to each segment (Baker, 1987; DiNardo et al., 1985; DiNardo and O'Farrell, 1987;
Kornberg et al., 1985).
One of the key features of Drsophila development which allows for the nearly
simultaneous patterning of segments is the fact that while segments are patterned the
embryo is a syncytium (reviewed in (Bate, 1993; Campos-Ortega, 1985). During this
syncitial blastoderm stage, which follows fertilization, the embryo undergoes 13
synchronous nuclear divisions. In this stage, there are no cell membranes separating the
nuclei, allowing for transcription factor mediated signaling without the use of cell surface
receptors.
Comparisons between the segmentation program in chordates and Drosophila would
deduce that the common ancestor to these two phyla was most likely not segmented,
since they do not share any segmentation mechanisms. However, the fly is a highly
derived hexapod, and hence not be the best model for generalizations of the mode of
segmentation in the arthropods (see Figure 1.1B for the evolutionary relationship between
arthropod groups) (Averof, 1995). Unlike the nearly simultaneous development of
segments in the Drosophila embryo, most arthropods are sequentially segmenting
developers and develop at least the posterior segments in a sequential pattern from
anterior to posterior from the growth zone located in the posterior region of the
unsegmented trunk. This is similar to the progression of vertebrate segmentation. It is
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most likely that the ancestral arthropod was sequentially segmenting (Davis and Patel,
2002; Stern, 1990; Tautz, 1994).
Nevertheless, what we know about Drosophila segmentation can be used as a stepping
stone to learning more about segmentation in other arthropods, through a candidate gene
approach. An important factor to consider when comparing the regulation of segment
patterning in sequentially segmenting developers is that the patterning of segments occurs
in a cellularized environment, and hence can not rely solely on transcription factor
gradients (Peel et al., 2005). This leads us to two equally interesting questions: (1) How
much of the molecular cascade underlying Drosophila segmentation is involved in
patterning segments in sequentially segmenting Arthropods? (2) What cell-cell
communication pathway(s) are necessary for proper sequential segment patterning?
Candidate approaches based on the Drosophila model have attempted to ascertain the
level of conservation in the segmentation patterning mechanisms of sequentially
segmenting arthropods. The following sections summarize what has been learned about
the conservation of the Drosophila pathway in sequentially segmenting arthropods.
Comparing Segmentation in Sequentially Segmenting Arthropods to Drosophila –
Maternal Coordinate Genes
One of the first maternal signals that establish the anterior of the embryo is bicoid.
Although bicoid genes have been isolated in several lower dipteran species (Sommer and
Tautz, 1991; Stauber et al., 2000), attempts at isolating this gene from more distantly
22
related species have been unsuccessful (Brown et al., 2001; Stauber et al., 2002). In fact,
it has been suggested that bicoid may be specific to Diptera (Stauber et al., 1999). In the
flour beetle Tribolium castaneum the gene products of hunchback, orthodenticle-1,
pangolin, and eagle are localized in the anterior of the embryo and there is functional
data that the first two are indeed required for anterior patterning (Bucher et al., 2005;
Schroder, 2003). Furthermore, knockdown of orhthodenticle-1 and hunchback in
Tribolium results in larvae missing the complete head, a phenotype reminiscent of bicoid
loss of function mutations in Drosophila (Schinko et al., 2008). As such, it appears that
these two genes, which serve as gap genes in the Drosophila segmentation hierarchy,
organize anterior patterning. This anterior patterning role for orthodenticle in non-
dipteran insects is supported by the finding that it is required for head formation in the
wasp Nasonia vitripennis (Lynch et al., 2006). The role of orthodenticle in patterning the
anterior of the embryo remains to be determined in non-insect arthropods. In fact, it has
been suggested that sequentially segmenting arthropods may not require anterior
signaling to properly pattern segments (Lall et al., 2003). However, it is just as likely that
the anterior signal is not one found in Drosophila. I propose several alternative
hypotheses to explain anterior coordination of the sequentially developing arthropod.
First, with the recent findings that some of the signaling components involved in
patterning vertebrate segments may be conserved in arthropods, it is possible that the
anterior signal is retinoic acid. Second, it is possible the anterior of the unsegmented
region in the sequentially segmenting arthropod embryo, the growth zone, is specified by
the posterior-most segment. Third, the anterior can be specified by the absence of a
23
signal rather than the presence of one, possibly creating a permissive zone for segment
formation.
Establishment of the posterior pole seems to be more highly conserved among the
arthropods. Caudal expression is required for proper posterior signaling during early
Drosophila segmentation, and the requirement for this gene product in the posterior of
the embryo is even more pronounced in other arthropods studied to date. In Drosophila,
caudal is required for the formation of only a few posterior segments (Macdonald and
Struhl, 1986). When caudal is knocked down in Tribolium and the cricket Gryllus
bimaculatus there is a loss of the posterior portion of the embryo and although the
phenotypes range in severity, the most extreme RNAi phenotype is a loss of all but the
anterior-most head segments (Copf et al., 2004; Shinmyo et al., 2005). As expected,
caudal is expressed in the posterior regions of both Tribolium and Gryllus embryos
(Schulz et al., 1998; Shinmyo et al., 2005). Similarly, knock down of caudal in the
crustacean Artemia franciscana resulted in the loss of posterior segments, in the most
severe cases including some of the posterior trunk segments (Copf et al., 2004).
Presumably, caudal is also necessary for the posterior patterning of the grasshopper
Schistocerca gregaria (insect) (Dearden and Akam, 2001), the barnacle Sacculina carcini
(crustacean) (Rabet et al., 2001), and the centipede Strigamia maritima (myriapod)
(Chipman et al., 2004) since it is expressed in the posterior region of the developing
embryo in both these species; however there is no functional data to support that
conclusion. Another signal required for the proper posterior patterning in Drosophila is
24
nanos. Unlike caudal, nanos expression has not been characterized in many other
arthropods, and its involvement in posterior patterning has not been functionally tested.
However, nanos is expressed in the posterior of the embryo in the grasshopper
Schistocerca americana (Lall et al., 2003), and three different species of mosquitoes
(Calvo et al., 2005).
Comparing Segmentation in Sequentially Segmenting Arthropods to Drosophila – Gap
Genes
Drosopihla gap genes are under direct regulation by the maternal coordinate genes. Their
expression spans several segments of the embryo, and loss of gap gene function results in
a loss of several neighboring segments creating a ‘gap’ along the anterior-posterior axis
of the embryo. Hunchback, Kruppel, and giant orthologs have been found in Tribolium
(Bucher and Klingler, 2004; Cerny et al., 2005; Marques-Souza et al., 2008; Schroder,
2003; Sommer and Tautz, 1993; Wolff et al., 1995), and as the Drosophila model would
predict, they are expressed in domains that span several segments. However, their loss of
function does not result in a gap in the segment pattern as in flies. Instead, it results in a
loss of posterior segments and homeotic transformation of some of the remaining
segments. These findings are parallel to what has been found in Gryllus (Mito et al.,
2006; Mito et al., 2005), Schistocerca (Patel et al., 2001) and Oncopeltus (Liu and
Kaufman, 2004a; Liu and Kaufman, 2004b). Thus, gap genes serve a different function in
sequentially segmenting arthropods than they do in Drosophila. In the fly, gap genes act
upstream of signals that initiate the patterning of each segment, where as in sequentially
25
segmenting arthropods, these genes appear to play a role in patterning the identity of
already patterned segments. It appears that their spatial expression domains made them
an ideal candidate for being co-opted for segment patterning in Drosophila.
In Drosophila, gap genes directly regulate the primary pair rule genes hairy, runt and
even-skipped. Kruppel loss of function results in a loss of even-skipped expression both
in the domain of Kruppel expression, and in more posterior segments in both Tribolium
(Cerny et al., 2005) and Gryllus (Mito et al., 2006), indicating that its regulation must be
transduced to neighboring posterior segments. Furthermore, even-skipped has been
shown to have gap gene function in both Oncopeltus (Liu and Kaufman, 2005) and
Gryllus (Mito et al., 2007), was also unexpected based on the Drosophila model and
suggests that perhaps even-skipped lost its gap gene role in Drosophila development.
There have been very few investigations of gap genes in arthropods other than insects. In
the myriapod (Strigamia) the expression of both hunchback and Kruppel has been looked
at, but only in relation to neural development (Chipman and Stollewerk, 2006). In the
crustacean (Artemia), expression studies suggest that hunchback function is restricted to
neurogenesis and mesodermal patterning, but has no role in the patterning of segments
(Kontarakis et al., 2006). All in all, gap gene function in Drosophila segmentation
appears to be highly derived and limited to simultaneously segmenting arthropods. By
contrast the involvement of Hox genes in segmentation of sequentially developing
arthropods appears to be more conserved.
26
Comparing Segmentation in Sequentially Segmenting Arthropods to Drosophila – Pair-
rule Genes
Based on the Drosophila model, one would predict that pair-rule genes would function in
alternating segment-wide domains, mirroring their role in the fly genetic hierarchy.
Interestingly, Drosophila pair-rule genes can be subdivided into two categories (Pankratz,
1993). The primary pair-rule genes, even-skipped, hairy, and runt, respond directly to
information from the gap genes and regulate secondary pair-rule genes, including fushi-
tarazu, odd-skipped, paired, odd-paired, and sloppy-paired, which then play a more
direct role in regulating segment polarity genes. Expression of these genes in Tribolium
suggests that pair-rule gene function is similar in short germ developers (Choe et al.,
2006; Patel et al., 1994; Sommer and Tautz, 1993). However, RNAi experiments have
led to a pair-rule function model that is different from that of Drosophila. Choe and
colleagues (2006) have found that even-skipped, runt and odd-skipped act as primary
pair-rule genes, while the functions of paired and sloppy-paired are secondary.
However, only the loss of even-skipped, odd-paired, and sloppy-paired leads to a loss of
alternating segments. Unlike in Drosophila, Tribolium primary pair-rules act in a
cascade, whereby even-skipped is required to activate runt, which is then required to
activate odd-skipped. Odd-skipped in turn inhibits even-skipped expression in the
neighboring segment. This lack of simultaneous signaling likely reflects the absence of a
syncytial environment and therefore the absence of simultaneous signaling by
transcription factors in neighboring nuclei.
27
The analysis of the function of pair-rule gene in other insects is much less complete,. The
expression patterns of paired orthologs have been described in Oncopeltus, Gryllus,
Schistocerca gregaria, and the honey bee Apis mellifera (Davis et al., 2001; Mito et al.,
2007; Osborne and Dearden, 2005). In all of these species, paired is expressed in every
segment of the developing animal and not in alternating segments as it is in Drosophila.
Even-skipped expression is also segmentally iterated in Oncopeltus where it has been
shown to have a pair-rule function, but it is expressed in partially single segment and
partially double-segment iterations in Gryllus (Liu and Kaufman, 2005; Mito et al.,
2007). Interestingly, fushi tarazu is expressed in stripes in the firebrat Thermobia
domestica, suggesting it plays a role in segment patterning in this organism, but there are
not stripes of fushi tarazu expression in Schistocerca gregaria (Dawes et al., 1994;
Hughes et al., 2004). In Artemia, the only crustacean where pair-rule gene expression has
been investigated, even-skipped is expressed in a broad posterior domain only (Patel et
al., 1992). There are segmentally iterated stripes of paired expression in the anterior
portion of the Artemia larvae, and there is a dynamic expression pattern in the posterior
region suggesting the presence of a segmentation clock similar to that regulating
vertebrate segmentation (Davis et al., 2005).
The double segmentally iterated pattern of Strigamia odd-paired suggests it has a pair-
rule role in segmentation in this myriapod (Chipman et al., 2004). In the centipede
Lithobius atkinsoni, even skipped is expressed in a broad posterior domain and only a few
28
stripes between the newly formed posterior segments, similarly to Artemia suggesting it
plays a role in segmentation (Hughes and Kaufman, 2002b). Similarly, Lithobius fushi
tarazu expression suggests it too is involved in segment patterning of this centipede, even
though it does not have a ‘pair-rule’ function as it does in Drosophila (Hughes and
Kaufman, 2002c).
The majority of pair-rule information for the chelicerates comes from expression data in
Cupiennius, where both primary and secondary pair rule genes are expressed in a stripe in
every segment (Damen et al., 2005; Damen et al., 2000; Schoppmeier and Damen,
2005a). Since some pair-rule genes are expressed more posterior in the growth zone than
others, it is likely that a hierarchy of regulation among pair-rules is conserved between
spiders, flies and beetles (Damen et al., 2005). Furthermore, in the spider mite
Tetranychus urticae paired is expressed in alternating segments similar to Drosophila
(Dearden et al., 2002).
The little data available for pair-rule genes in arthropods suggests that they are for the
most part involved in segment patterning in not only Drosophila, but the short germ
developers as well. However, the spotty sampling of expression patterns and even more
importantly, analysis of function of these genes are not sufficient to determine the exact
role these genes play, and their interactions not only with other pair-rules, but also with
gap genes and ultimately segment polarity genes. Apart from work done in Tribolium,
very little can be concluded from the expression data in other sequentially segmenting
29
arthropods, but it is apparent that pair-rule expression, and possibly function, is not as
highly conserved among this group as that of the segment polarity genes. Therefore, as
we shift our focus to the differences in segmentation mechanisms among arthropods as
opposed to the similarities, pair-rule genes are a prime target for those studies.
Comparing Segmentation in Sequentially Segmenting Arthropods to Drosophila –
Segment Polarity Genes
Unlike pair-rule genes, the expression of segment polarity genes is highly conserved
among all arthropods looked at to date, including insects, crustaceans, myriapods and
chelicerates (Damen, 2002; Damen et al., 2005; Dearden and Akam, 2001; Duman-
Scheel et al., 2002; Hughes and Kaufman, 2002d; Janssen et al., 2008; Janssen and
Damen, 2006; Manzanares et al., 1993; Nagy and Carroll, 1994; Nulsen and Nagy, 1999;
Patel et al., 1989; Peterson et al., 1998; Scholtz et al., 1994; Simonnet et al., 2004). Their
consistent expression in stripes in either the anterior (i.e.: wingless) or posterior (i.e.:
engrailed) of each embryonic segment suggests that just like in Drosophila segment
polarity genes specify anterior-posterior polarity in each segments. This is not surprising,
considering that in Drosophila segment polarity genes are active after cellularization of
syncytial nuclei has taken place. This suggests that the changes necessary to evolve the
Drosophila patterning system lie upstream of the segment polarity genes.
Comparing Segmentation in Sequentially Segmenting Arthropods to Drosophila –
Conclusions
30
With the notable exception of bicoid, the genes involved in the Drosophila segmentation
patterning network are also involved in the segmentation of sequentially segmenting
arthropods, even if their function is not conserved. This is remarkable, given that in
Drosophila these transcription factors can diffuse between closely neighboring nuclei in
the syncitium, whereas most arthropods, including crustaceans pattern their segments in a
cellularized environment. As such, in sequentially segmenting arthropods the signaling
between transcription factors patterning segmentation has to be transduced, for example
by receptor-ligand interactions. Indeed, the fact that segments form sequentially from
anterior to posterior in a cellularized environment makes short germ segmentation a
prime candidate for requiring signaling pathways that parallel those governing vertebrate
segmentation. This is especially true since the Notch/Delta signaling pathway mediates a
myriad of functions that involve cell-cell communication in all metazoanss (Lai, 2004).
An approach to determining the regulatory networks governing segmentation in
sequentially segmenting arthropods is to look at cell-cell signaling pathways. The most
obvious signaling pathway candidate is the Notch pathway, since its involvement in
vertebrate segmentation is well described, and the components of this signaling pathway
were the first molecular oscillators identified. Other signaling pathways that may be
transducing information in neighboring cells during segmentation are Wnt and FGF, both
of which are necessary for proper patterning of vertebrate segments. Finding that these
pathways coordinate segmentation in sequentially segmenting arthropods may support
the theory that the closest common relative to arthropds and vertebrates was segmented.
31
Cell-Cell Communication Pathway(s) in Sequentially Developing Arthropods:
Delta/Notch Signaling
Studies of components of the Notch signaling pathway in insects found no evidence that
they were involved in segment patterning in insects even though hairy and fringe are
expressed in segmentally iterated stripes in Tribolium and Schistocerca gregaria
respectively (Aranda et al., 2008; Dearden and Akam, 2000; Eckert et al., 2004; Sommer
and Tautz, 1993; Tautz, 2004). However, a very recent study in the cockroach
Periplaneta americana, where Notch signaling was knocked out with maternal RNAi,
found that Notch is required for the proper development of thoracic segments by
establishing both segmental borders and segment primorida, as well as being involved in
segment growth (Pueyo et al., 2008).
Furthermore, studies of the roles of Delta/Notch signaling in Cupiennius development
uncovered that this pathway is also involved in segment patterning in the spider
(Stollewerk, 2002; Stollewerk et al., 2003). Notch is ubiquitously expressed in the
posterior growth zone in the developing spider, and Delta, hairy, Suppressor of Hairless
and Presenillin are expressed in stripes that are added from anterior to posterior along the
growth zone and appear before there is morphological evidence of the corresponding
segment (Schoppmeier and Damen, 2005b; Stollewerk et al., 2003). Furthermore,
interruption of Delta/Notch signaling in spider by maternal RNAi of Notch leads to both
a loss of organization of hairy stripe expression in the growth zone and defects in the
32
size, shape and width of segments (Stollewerk et al, 2003; Schoppmeier and Damen,
2005b). In the spider Achaearanea tepidariorum, loss of Delta, Notch, or Suppressor of
Hairless activity by maternal RNAi also results in segmentation defects, the most severe
of which is an almost complete loss of segments, showing that Delta/Notch signaling is
involved in segment patterning in this spider species as well (Oda et al., 2007).
Mutations of Notch signaling components in spider show that the pathway is involved in
both patterning segments, and growth of the posterior growth zone, as mutants resulted in
a decrease in the size of the growth zone (Oda et al., 2007; Stollewerk et al., 2003).
Several studies report that myriapods too may use the Delta/Notch signaling pathway in
segment patterning. In the centipede Lithobius forficatus and the millipede Glomeris
marginata, Delta is expressed in stripes that are added in the posterior growth zone
during the course of development as it is in the spider (Damen et al., 2005; Kadner and
Stollewerk, 2004). Similarly, Delta and Notch expression in the centipede Sttrigamia
maritime also suggest a role for this pathway in segmentation (Chipman and Akam,
2008). To date, there is no functional data in myriapods to support Delta/Notch signaling
involvement in segmentation.
Within the crustacean, the only data which supports Delta/Notch signaling in
segmentation is from work done in Paryhale hawainesis. Notch is expressed in a broad
domain and Delta is expressed in stripes in the posterior growth zone of the animal.
Furthermore, treatment with the γ-secretase inhibitor DAPT which has been shown to
33
inhibit Notch cleavage, and hence disrupt Delta/Notch signaling in other organisms,
results in both a loss of segments and a decrease in the size of the growth zone in
Paryhale (O'Day, 2006).
Considering the fact that in short germ arthropods segments are added consecutively from
anterior to posterior in a cellularized environment, it is not surprising to find that a
receptor/ligand cell-cell communication pathway is involved in patterning these
segments. Several studies in arthropods show that Delta/Notch signaling components are
expressed in a diffuse band of expression in the posterior region of the growth zone.
Furthermore, stripes of expression of these signaling molecules, such as hairy and Delta,
originate from this posterior expression domain, and migrate anteriorly, eventually
forming a band of expression in each segment (Chipman and Akam, 2008; Pueyo et al.,
2008; Schoppmeier and Damen, 2005b; Stollewerk et al., 2003). Although this
expression pattern is reminiscent of the oscillations of these genes in the vertebrate PSM,
it is unlikely that these messages actually oscillate within individual cells in sequentially
segmenting arthropods. Instead, these expression patterns suggest that the expression of
these genes originates in cells in the posterior of the growth zone, and then these cells
move anteriorly to form segments. Perhaps instead of oscillatory expression patterns
along the anterior-posterior axis of the growth zone, these signaling molecules are turned
on or off as cells exit the posterior ‘determining zone’ of the growth zone.
34
Cell-Cell Communication Pathway(s) in Sequentially Developing Arthropods: Wnt and
FGF Signaling
Vertebrate segmentation also requires Wnt and FGF signaling. There are no reports of
investigation into the potential involvement of FGF in arthropod segmentation. Recently,
Wnt8 has been shown to control the formation of the posterior growth zone in
Achaearanea (McGregor et al., 2008). Loss of Wnt8 function results by RNAi results in
the absence of not only the posterior segments, but the posterior of the embryo.
Furthermore, loss of Wnt8 function in the spider affects the expression of Delta, hairy
and caudal in the posterior of the embryo. As such, Wnt8 not only establishes the growth
zone, but may play a permissive role for segmentation in the growth zone. This data is
supported by functional studies in Oncopeltus, and Tribolium (Angelini and Kaufman,
2005; Bolognesi et al., 2008). In both cases, wingless RNAi results in a complete loss of
several posterior segments. This is quite different from what is seen in Drosophila,
where wingless is a segment polarity gene, and its loss does not halt segmentation results
in the loss of a portion of each segment (Bejsovec and Martinez Arias, 1991).
Furthermore, wingless expression data the crustacean Triops longicaudatus, the myriapod
Lithobius, and the chelicerate Cupiennius, where wingless is not only expressed in a
segmentally iterated pattern, but also in a ring at the posterior of the growth zone,
supports this model (Damen, 2002; Hughes and Kaufman, 2002b; Nulsen and Nagy,
1999). These data suggest that future studies of arthropod segmentation shift from
looking for segmentation candidates from the Drosophila model to those in vertebrates.
35
Cell-Cell Communication Pathway(s) in Sequentially Developing Arthropods:
Conclusions
The use of the Delta/Notch and Wnt signaling in segmentation in both arthropods and
vertebrates suggests homology of segmentation between these two phyla, and that their
last common ancestor may have been segmented. The fact that both phyla use the Notch
signaling pathway to pattern segments may not necessarily be surprising, considering the
many processes that rely on this pathway. Since both Delta/Notch signaling is upstream
to many regulatory components in segment patterning may support the theory that there
is a common origin for the involvement of this pathway in segmentation, and suggest that
the common ancestor to vertebrates and arthropods shared this regulatory network
(Damen, 2007). However, the abundance of Delta/Notch signaling in development
makes the possibility that this pathway was co-opted to mediate cell-cell communication
in the segmentation process on two or more separate evolutionary occasions impossible
to completely disregard. The fact that both arthropods and vertebrates also use the Wnt
signaling pathway does add support to the theory of a common segmented ancestor. This
is especially true in light of recent findings that Wnt may play a permissive role in the
patterning of segments in both phyla (Aulehla et al., 2008; Dunty et al., 2008; McGregor
et al., 2008). The conclusion that the common ancestor to arthropods and chordates was
segmented would imply that segmentation is conserved among all three segmented phyla.
As such, this theory would be strengthened if Annelids shared common mechanism of
segmentation with arthropods and vertebrates.
36
Annelid Segmentation
Annelid segmentation stems from five teloblast cells on either side of the embryo. Soon
after their birth, teloblast cells undergo repeated asymmetrical divisions to produce a
bandlet of smaller daughter cells which are together referred to as primary blast cells.
Four of these five bands of cells join to form an ectodermal germ band and the remaining
forms a mesodermal germ band. As the pair of germ bands elongate, they gradually
curve and eventually meet to form the germinal plate, which is the site of segmentation.
Segments develop from anterior to posterior. Each segment is comprised of progeny
from one daughter cell of each of the teloblasts through a series of divisions of the blast
cells. As such, it is likely that molecular programs that govern the patterning of segments
are linked to the cell cycle (Rivera et al., 2005). This is a very different mechanism for
segmentation than in vertebrates, where cells in the PSM undergo very little division
during the segmentation process (see Pourquie, 2001 for review). However, the growth
zone of the sequentially segmenting arthropod continues to elongate in the posterior as
segments are added in the posterior, and segments may be prepatterned in the
proliferative posterior region (McGregor et al, 2008).
Unfortunately, our understanding of the molecular basis of segmentation in annelids is
very limited. In the marine annelid Platynereis dumerilii, expression data suggests that
caudal and even-skipped are involved in the posterior addition of segments as both are
expressed in the posterior growth zone (de Rosa et al., 2005). On the other hand, several
37
studies conclude that hunchback is not likely involved in the patterning of segments in
leech or Tubifex (Iwasa et al., 2000; Shimizu and Savage, 2002). Expression studies
suggest that hairy plays a role in segmentation in the leech Helobdella robusta (Song et
al., 2004). Hairy expression peaks when teloblast cells are generating blast cells.
Moreover, the timing of hairy expression of both transcript and protein show variation at
different stages of the cell cycle, as protein levels are high during interphase, and
transcripts are high during mitosis (Song et al, 2004). Similarly, Notch is also transcribed
in both the teloblasts and blast cells suggesting that it too plays a role in Helobdella
segmentation (Rivera et al, 2005). Inhibition of both hairy and Notch in Helobdella does
result in the perturbation of segmentation, as was predicted by the expression studies
(Rivera and Weisblat, 2008). Furthermore, expression studies of Notch, Delta, and
several hairy homologues in the polychete annelid Capitella sp. I suggest that the Notch
signaling pathway may be involved in prepatterning segments in the posterior
unsegmented growth zone in a pattern reminiscent of Wnt activity in the segmentation of
the spider as discussed above (Thamm and Seaver, 2008). This finding of the
involvement of Notch signaling in the patterning or annelid segments is further supports
the argument that the three segmented phyla shared a common segmented ancestor.
However, this conclusion has to be made cautiously since the role of Notch in the
formation of segments varies so much between the groups, and the since the Notch
signaling pathway is so prevalent in mediating cell-cell signaling throughout
development, it could easily have been co-opted to the process of segmentation in more
than one evolutionary event.
38
Concluding Remarks and Rationale for Study
There is still much to be learned about segmentation in the three segmented phyla in
order to differentiate between the three hypotheses regarding the origin of the
segmentation process (outlined in Figure 1.1A). Determining whether arthropods,
annelids and chordates shared a common segmented ancestor would be easier if we could
compare the segmentation program of the most basal living species in each of these
phyla. The arthropod group is comprised of hexapods, crustaceans, myriapods and
chelicerates, of which insects (a class of hexapods) are the most derived. Yet our
understanding of arthropod segmentation rests predominantly on information in insects.
Within the arthropods, insects are most closely related to crustaceans, from which they
likely derived (Averof, 1995). As such, crustaceans are more basal and their mode of
segmentation is more likely to accurately reflect that of the last common ancestor to all
arthropods. Moreover, comparing crustacean segmentation programs to those of insects
can help eliminate features of insect segmentation that are derived, and hence should not
be used to compare arthropod segmentation to segmentation in chordates and annelids.
Unfortunately, very little is known about segmentation in crustaceans. As such, I set out
to learn about the role of Notch signaling in the crustacean Artemia franciscana. The
Artemia larva is free swimming when it hatches out of its cyst, the first three of five head
segments are already formed, but the trunk is an unsegmented mass of undifferentiated
39
cells that grows by cell division spread throughout its field (Freeman, 1986; Freeman et
al., 1992) (Figure 1.2). As the larva develops the trunk segments are patterned
sequentially from anterior to posterior such that anterior segments are more developed
than their posterior counterparts (Anderson, 1967) (Figure 1.2). The first evidence of
segmentation is the rearrangement of nuclei in the posterior growth zone into several
distinct domains (Freeman et al, 1992). In the most anterior portion of the unsegmented
trunk, the nuclei are arranged into rows spanning the width of the trunk, which is the
earliest morphological marker for segment development. Further posterior, the nuclei are
arranged in columns along the anterior-posterior axis of the trunk. The posterior tip is
comprised of unorganized nuclei. The adult Artemia has 11 thoracic/trunk segments, 2
genital segments and 6 post-genital segments.
I first characterized the timing of segmentation in Artemia. I found that segments are not
added at regular intervals (Chapter 2). However, morphological analysis provided
evidence that all trunk segments are added sequentially in this crustacean, and there are
no morphological differences in the segments that can account for the increasing length
of time between segment additions (Appendix A). I then inhibited Notch signaling in
Artemia embryos by treating them with a γ-secretase inhibitor (see Figure 1.3), and found
that Notch plays a role in segmentation but not growth, as treated larvae have fewer
segments than controls but their body length is not affected by the drug (Chapter 2).
40
Figure 1.1: Hypotheses for the evolution of segmentation. Trees show relationship between Arthropods, Annelids and Chordates (Anderson, 2004). These three clades may share a common segmented ancestor (red). Alternatively, Arthropods and Annelids may share a common segmented ancestor, and Chordate segmentation evolved independently (blue). Lastly, segmentation may have evolved independently three times, leading to segmentation in these three phyla (green).
41
42
Figure 1.2: (A) The Artemia embryo hatches without trunk segments. Trunk segments are then added one at a time from anterior to posterior. The cartoon shows the nuclear organization of nuclei along the anterior-posterior axis of the trunk. Just posterior to the last segment the nuclei are arranged in rows across the width of the trunk indicating the patterning of the next segment. Below that, nuclei are arranged in columns. In the posterior tip, nuclei are not organized. (B) The adult brine shrimp has 11 trunk segments, 2 genital segments, and 5 post-genital segments. (C) DAPI stained larva with all 11 trunk segments, both genital segments, and the first post-genital segment. Since segments continue to develop after they are initially established, anterior segments have are more developed than anterior segments, as is seen in the progress of limb development on each segment.
43
44
Figure 1.3: Summary of the Delta/Notch signaling pathway. In short, Delta and Notch are both cell surface receptors, so their interaction requires cell-cell contact. Upon activation of Notch by Delta, the Notch intracellular domain (NID) is cleaved. This cleavage allows the NID to dissociate from the cell membrane and migrate to the nucleus, where it is involved in the regulation of gene expression. One of the key components in the cleavage of the Notch receptor is γ-secretase, which can be inhibited pharmacologically by DAPT. The inhibition of proteolytic cleavage of the Notch receptor inactivates the pathway, even in the presence of Delta (Adapted from Radtke et al, 2005).
45
46
CHAPTER 2: GAMMA SECRETASE INHIBITION ARRESTS SEGMENTATION IN THE BRANCHIOPOD CRUSTACEANS THAMNOCEPHALUS PLATYURUS AND
ARTEMIA FRANCISCANA
* The data in this chapter was gathered as a collaborative effort with Drs. Terri Williams and Tom Hegna at Yale University
Abstract
The presence of repeated body segments is the defining feature of arthropods,
however, the morphological process by which segments are added is itself evolving.
Although some species form their segments simultaneously without any measurable
growth, most arthropods add segments sequentially from a region of differential growth
in the posterior of the growing embryo or larva. We do not know whether the molecular
mechanisms underlying sequential segment addition are highly disparate or highly
conserved. Notch signaling is involved in segmentation in sequentially segmenting
arthropods, as inferred from both the expression of proteins required for Notch signaling
and the genetic or pharmacological disruption of Notch signaling. In this study, we
demonstrate that blocking N signaling by blocking γ-secretase activity causes a specific,
repeatable effect on segmentation in anostracan crustaceans. We observe a slowing of
the rate of segment addition. Slowing is dose-dependent and higher doses of DAPT cause
slower segment addition. Despite this marked effect on the rate of segment addition,
other aspects of segmentation are unaffected. Segment size and boundaries between
segments appear normal, EN stripes are normal in size and alignment, and overall growth
is unaffected. Our findings are consistent with a growing body of evidence that N plays a
role in sequential segmentation in arthropods. At the same time, our observations
47
contribute to an emerging picture that loss of function N phenotypes differ significantly
among the arthropods sampled to date.
Introduction
Nearly all of the millions of arthropod species develop their segments in the same
fashion: they add them one by one from the posterior in a region commonly called the
“growth zone”. The exceptions are the higher insects, including the intensively studied
arthropod model system, Drosophila. Instead of sequential segment addition, these
species produce their segments nearly simultaneously in an acellular syncytium (see Bate,
1993; Campos-Ortega, 1985 for review). Segmentation from a posterior growth zone is
also common to chordates and annelids. We do not know whether sequential segment
addition, either within or between phyla, involves a variety of highly disparate
mechanisms or, conversely, reflects a highly conserved set of mechanisms (Damen,
2007).
The best understood mechanisms of sequential segmentation come from vertebrate
models. Vertebrate segmentation occurs in a bilaterally symmetrical subpopulation of
paraxial mesodermal cells lying on either side of the neural tube that forms during
gastrulation. The presomitic mesoderm (PSM) extends posterior during segmentation by
the addition of cells from the posterior tip of the embryo (Kanki and Ho, 1997). Segments
48
form at regular time intervals and the periodicity of segment formation is species specific.
Segmental prepattern arises from a “segmentation clock” that functions to translate
temporal information of cyclic patterns of gene expression into positional information
(see Dequeant and Pourquie, 2008 for review).
The first genes recognized to oscillate with a periodicity similar to somite formation were
downstream targets of the Notch signaling pathway (Forsberg et al., 1998; Palmeirim et
al., 1997). The expression of these genes begins in the posterior of the PSM then travels
anteriorly in a wave: more anterior cells express the genes while more posterior cells turn
them off. Expression is eventually isolated to a small band of cells in the region where
the new somite will form. Patterning genes, most notably the Hox genes, then specify the
morphological identity of the somites based on their position along the PSM (Krumlauf,
1994). There is evidence for coordination between the Hox patterning pathway and
Notch signaling during somitogenesis, and likely begins before somitogenesis as Hox
genes are first expressed during garstrulation (Cordes et al., 2004; Dubrulle et al., 2001;
Peres et al., 2006). If the core components of Notch signaling are disturbed during
vertebrate embryogenesis, gene expression in the presomitic mesoderm is disrupted,
irregularly sized somites with disrupted segment borders form and symmetry between
somite pairs across the midline of the embryo is lost (Holley and Takeda, 2002; Pourquie,
2003; Rida et al., 2004). Despite the striking correlation between the oscillatory
expression of genes in the Notch signaling pathway and the pace of segment formation,
Notch does not appear to be the pacemaker of the molecular oscillations (Holley et al.,
49
2002; Jouve et al., 2000; Morales et al., 2002). In the absence of Notch signaling,
oscillations of gene expression are maintained. However, the synchrony of the
oscillations between neighboring cells is lost, suggesting that the segmentation clock does
not stop in individual cells but is desynchronized along the length of the PSM (Jiang et
al., 2000). In these mutants, the anterior segments are normal, but as the oscillations
become progressively desynchronized segment borders become increasingly affected,
and eventually segmentation fails caudally (Jiang et al., 2000; Lewis, 2003; Riedel-Kruse
et al., 2007). The identity of the molecular pacemaker remains unknown (Ozbudak and
Pourquie, 2008).
It is unclear whether anything analogous to a segmentation clock exists in arthropods.
Even the most basic data that might support this hypothesis, i.e., addition of segments in
regular time intervals and with species-specific periodicities, are lacking for most
arthropods. One approach for identifying a segmentation clock is to examine Notch
signaling in sequentially segmenting arthropods. There is evidence that Notch signaling
is required to form segments in the chelicerates, (Oda et al., 2007; Schoppmeier and
Damen, 2005b; Stollewerk et al., 2003) in the myriapods Lithobius forficatus (centipede)
and Glomeris marginata (millipede) (Janssen, 2004; Kadner and Stollewerk, 2004) and
basal insects (Pueyo et al., 2008). In these species, segmentally repeated patterns of
transcripts of Notch signaling components, including Notch, Delta, hairy, Suppressor of
Hairless, and Presenillin have been observed. RNAi knockdown of Notch yields a range
of phenotypes including defects in the size, shape and width of segments. Notch disrupts
50
segment addition and leaves gaps in engrailed expression. In the most extreme
phenotype, segmentation is halted. These results have established that Notch signaling is
required for proper segmentation in sequentially segmenting arthropods and likely
represents a component of the basal patterning mechanism although they do not as yet
provide evidence for a segmentation clock. As in vertebrates, once segments are
patterned, Hox genes play a major role in establishing the identity of each segment along
the anterior-posterior axis (Cook et al., 2001; Hughes and Kaufman, 2002d). However,
these genes are activated before the onset of segment patterning and the appearance of
segmentation, suggesting that their patterning role is coordinated with the patterning of
segments (Averof and Akam, 1995)
In the Notch pathway, both ligand and receptor are membrane associated and signaling is
triggered by cell interactions. The binding of Notch to its ligand initiates a γ-secretase-
mediated proteolysis of the intracellular domain of the receptor. The cleaved intracellular
domain translocates into the nucleus, where it associates with other transcription factors
(Artavanis-Tsakonas et al., 1999; Bray, 1998; Kadesch, 2004). Application of the γ-
secretase inhibitor, N-S-phenyl-glycine-t-butyl ester (DAPT) to Drosophila and zebrafish
embryos is known to phenocopy Notch mutations (Geling et al., 2002; Micchelli et al.,
2003). Cleavage by the γ-secretase is inhibited by DAPT, the intracellular domain of
Notch does not move to the nucleus, and thus, the Notch pathway is blocked.
51
To clarify a role for Notch signaling in crustaceans, we examined segmentation in two
species of branchiopod crustaceans Artemia franciscana and Thamnocephalus platyurus.
Based on the phylogenetic analysis, anostracans are basal among Pancrustacea (Dumont,
2002; Negrea, 1999) and have a developmental mode that is likely basal for all
arthropods (Davis and Patel, 2002; Stern, 1990; Tautz, 1994). We first established
normal timing of segmentation along the growth zone and find species specific variation
in segment addition. We then used a pharmacological approach to disrupt Notch function
in crustaceans by inhibiting Notch signaling with DAPT. We find that blocking γ-
secretase activity disrupts segment formation. In both species, DAPT treatment results in
a decreased rate of segment addition and a decrease in the number of total segments,
suggesting that the Notch/Delta pathway is involved in segmentation.
Methods
Animal Rearing and Fixing
Artemia franciscana cysts (San Francisco Bay Brand) were hatched and grown in filtered
artificial sea water (FASW) (Instant Ocean) at 21°C or 27°C in a temperature controlled
incubator at a density of 5 larvae/mL. Each day, the water was changed and then
supplemented with 0.002% of a super-saturated yeast and algae solution made by
grinding a 1.41 oz algae pellet (#21307, Hikari USA) in a mortar and pestle with equal
volume of active dry yeast (Fleischmann's, ACH Food Companies) dissolved in 10 mL of
52
FASW. The food solution was stored at 4°C for up to 1 week and before use the mixture
was stirred and larger particulates were allowed to settle for 5 min. Higher density
Artemia cultures were grown at 100 larvae/mL at 21°C. Animals were fixed on a shaker
for 30 min at RT in 3.7% formaldehyde in FASW and stored in 100% MeOH at -20°C
prior to immunodetection. Thamnocephalus platyurus Packard, 1877 cysts (a gift from
the late D. Belk) were hatched and grown in artificial pond water at room temperature
(21°C). The animals subsisted on yolk for the duration of the study. Animals were fixed
for 30 minutes in 9% formaldehyde in PBS with 50mM EGTA.
Drug Treatment
A stock solution of 10mM N-S-phenyl-glycine-t-butyl ester (DAPT, Calbiochem/ EMD
Biosciences) in DMSO was prepared. It was stored at -20°C for no more than 2 weeks
since we found that the potency of the drug decreases over time. In all treatments larvae
were continuously exposed to DAPT. For Artemia, cysts were hatched in 3 mL FASW
containing either 50µM or 100µM of DAPT and 1% DMSO. At the higher concentration,
not all the drug dissolved into the FASW. Larvae were collected from a pool of cysts
within 1 hour of hatching and raised at a concentration of 100 larvae/mL. The larvae were
placed in fresh FASW and DAPT supplemented with 0.002% yeast and algae solution
every 24 hours for the duration of the experiment. For T. platyurus larvae were removed
from their hatching tank immediately after hatching and raised individually in 1mL of
pond water supplemented with either 50µM or 100µM of DAPT and 1% DMSO. To
53
prevent the formation of large DAPT crystals the DAPT/pond water solution was agitated
immediately after addition of DAPT and one hour thereafter. All control animals were
treated with 1% DMSO and otherwise fed and treated like experimentals.
Immunolocalization
Antibody staining was performed according to established protocols (Panganiban, 1995;
Patel et al., 1994). Engrailed antibody (En4F11) (Patel et al., 1989) was used at a 1:10
dilution for Artemia and a 1:5 dilution for Thamnocephalus. Ubx/AbdA antibody
(FP6.87) (Kelsh et al., 1994) was used at a 1:5 dilution. Alkaline phosphatase (AbCam)
and Cy3 (Jackson) conjugated anti-mouse secondaries were used at 1:250 (Artemia) and
1:200 (Thamnocephalus). Alkaline phosphatase secondaries were detected with a color
reaction (NBT/BCIP) according to manufacturer directions (Roche). Animals were
visualized using a compound microscope (Artemia: Zeiss Axioplan, München, Germany;
Thamnocephalus; Nikon Ellipse 600). Artemia color staining was photographed with a
digital color camera (Go Series, QImaging) using Q Capture Pro 5.0 software, while
fluorescence was photographed with a digital black and white camera (Zeiss AxioCam)
using AxioVision software. Thamnocephalus animals were photographed with SPOT
Insight camera (Diagnostic Instruments) using Spot Advanced software.
Results
Timing of segment addition in Artemia and Thamnocephalus
54
Both species emerge from the egg with three anterior head appendages present. Segments
develop progressively from an undifferentiated trunk to form the adult (Fig. 1). Although
quite similar in their progressive addition of segments, Artemia and Thamnocephalus
differ in some aspects of segmentation, notably, the number of stripes of Engrailed (EN)
protein present at hatching and the rate of segment addition. Thamnocephalus hatch with
between 2 and 3 EN stripes and as new segments become visible there are two EN stripes
posterior to the last segment. The first segment becomes visibly distinct about an hour
after hatching and then segments are added at a rate of about one segment every 40
minutes (Fig. 2A). Within about 7 hours the 11 trunk segments are present.
Thamnocephalus is like another branchiopod, Triops longicaudatus, in having a relatively
rapid and steady addition of segments (Ko and Williams, in prep). By contrast, Artemia
hatch without any detectable EN stripes and they add segments more slowly. The first
five EN stripes are added at an interval of six hours/segment, the following six EN stripes
at intervals of roughly 24 hours/segment (there is a progressive lengthening of the
interval from 22 to 27 hours, Fig. 2B). The rate of segment addition in Artemia is the
same at room temperature (21° C) and at higher temperatures, (27°C, Fig. 2B). In
addition to the slower overall rate of segmentation in Artemia, they also have a delay of
10 hours between hatching and the expression of the first thoracic EN stripe.
DAPT disrupts segmentation in a dose dependent manner
55
When larvae of either species are raised in the presence of DAPT, segmentation slows in
a dose dependent manner (Fig. 3). Artemia larvae hatched and raised in 50µM
continuous DAPT develop fewer segments compared to control animals (Fig. 3 A-D).
Artemia larvae hatched and raised in 100µM of DAPT have fewer segments than both
control and 50µM treated animals (Fig. 3D). Larvae treated with the higher DAPT
concentration only develop the first two to three thoracic segments in the first 96 hours of
development, during which time untreated embryos develop five segments. Of note, the
first two segments always form, even under high concentrations of DAPT. The segments
in γ-secretase treated larvae were not visibly different from the controls in size and shape,
and the borders between segments were not affected. Similarly, the gnathal EN stripes
were unaffected by the drug treatment (Fig. 3A-C).
The results in Thamnocephalus parallel those found in Artemia. By 7.5 hours after
hatching, Thamnocephalus control larvae have on average 5.6 segments, larvae raised in
50 µM DAPT have 4 segments and those raised in 100 µM DAPT have 3.6 segments
(Fig. 3E.)
To evaluate the timecourse of DAPT effects, we fixed treated Artemia larvae at
intermediate time points between hatching and 96hrs. At the earliest timepoint we
measured, when control larvae have just over two segments, we find a small but
56
significant difference in the rate of segment addition. This difference between the average
number of segments added diverges over time.
DAPT treatments do not effect overall growth
The overall larval growth was not affected by exposure to DAPT. The body length did
not differ between 96 hour untreated control animals (1658 µm ± 74.1 µm; n=7) and 100
µM DAPT treated larvae (1568 µm ± 72.4 µm; n=7; Fig. 4A). They were both
significantly larger than the 32 hour control animals (1058 µm ± 37.0 µm; n=7, Fig. 4A)
that have approximately the same number of segments as the DAPT treated animals.
Thus, it is unlikely that the reduced number of segments in γ-secretase treated animals is
due to a generic developmental delay.
In Thamnocephalus, DAPT treated animals have a slightly shorter body length than in the
controls (Fig. 4B) but are much longer than younger larvae with a comparable number of
segments. Also, Thamnocephalus body length is the same, despite differences in segment
number, under the two different concentrations of DAPT treatment which are raised for
the same amount of time. Thus, it is unlikely that a decrease in the number of trunk
segments in DAPT treated animals can be explained by the slight difference in body
length between treated larvae and controls.
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Posterior HOX gene expression is arrested in DAPT treated animals
The monoclonal antibody that detects an epitope shared by the abdominal-A and
Ultrabithorax proteins (Ubx/abdA; Kelsh et al, 1994) is first expressed in the Artemia
larvae in a broad domain that corresponds to the unsegmented growth zone (Fig. 5A;
Averof and Akam, 1995). Stripes consisting of single cells expressing Ubx/abdA are first
detectable at the anterior end of the growth zone, prior to any evidence of segmental
morphology, or nuclear organization that precedes segment formation. In older larvae,
Ubx/abdA expression is detected throughout the visible segments, and in segmentally
iterated stripes in the growth zone, just posterior to the last morphologically discernible
segment. A broad domain of expression is also detectible in the posterior of the growth
zone. In larvae treated with 100µM DAPT, Ubx/abdA protein is expressed in the two or
three thoracic segments that develop normally. However, stripes of expression posterior
to the last visible segment are lost (Fig. 5B). The single cell wide stripes of Ubx/abdA
expression in the region of newly forming segments are absent. Expression in the broad
domain of Ubx/abdA in the posterior unsegmented region is detected in only a few
nuclei. Thus Notch signaling appears to be required to activate and/or maintain
Ubx/abdA expression in the posterior growth zone.
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Discussion
DAPT slows the rate of segmentation in anostracan crustaceans
Addition of DAPT, a known γ-secretase inhibitor, causes a specific, repeatable
effect on anostracan segmentation: the rate of segmentation is slowed. Slowing is dose-
dependent, with fewer segments produced at higher doses of DAPT. Although higher
concentrations of DAPT results in slower rates of segmentation, segment addition was
never completely halted. Increasingly higher doses simply resulted in larval death.
Despite this marked effect on the rate of segment addition, other aspects of segmentation
are unaffected. Segment size and boundaries between segments appear normal, and EN
stripes are normal in size and alignment. In addition, overall growth of the larvae is
unaffected by DAPT treatment.
Although segments from the third thoracic segment posterior could be measurably
slowed by exposure to DAPT, we were unable to discern whether the first two thoracic
segments are insensitive to N signaling. At our first measured timepoint the difference
between treatments is small but significant: it varies from 2.6 (110uM DAPT) to 3.2
(control) segments. In subsequent timepoints, the differences in segment number are
greater although the rate appears to remain more or less constant after 24h. This
nonlinearity in response to treatment could reflect a subset of anterior segments that are
59
not N sensitive – a phenomena found in a variety of other animals (Stollewerk et al,
2003; Pueyo et al, 2008).
Overall, our findings are consistent with a growing body of evidence that N plays
a role in sequential segmentation in arthropods. At the same time, our observations
contribute to the emerging picture that loss of function N phenotypes differ significantly
among the arthropods sampled to date.
Blocking Notch activity in sequentially segmenting arthropods has variable effects.
The major effect of disrupting N or N pathway genes in sequentially segmenting
arthropods is that segmentation either slows or halts. However, the way this occurs is
distinct in different taxa as indicated by a variety of phenomena (Table 1). For example,
effects on segmentation can reflect disruption of boundary formation between segments.
In the most extreme cases, (Pueyo et al, 2008) blocking N signaling sometimes produces
embryos with different numbers of EN stripes on the right and left side of the germband.
Other times, boundary effects are less severe and include incomplete EN stripes (Pueyo et
al, 2008) or crooked and irregular EN stripes (Stollewerk et al, 2003). In some cases
(anostracans crustaceans), EN stripes are normal. Thus, depending on the species tested,
segment boundaries show a wide range of effects, from no effect, to irregular boundaries,
to right-left mismatch.
Another contrast among arthropods that is particularly striking is the axial
position of segment disruption. As mentioned above, all segments behind the head in
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anostracans may be N-sensitive whereas, in cockroach for example, segmentation appears
N-insensitive until the fourth abdominal segment. In cases where maternal RNAi
injections are used, N-signaling is blocked well prior to segment formation and there are
clearly instances of N-insensitive segments. In our case, we are unable to expose animals
prior to their emergence from the cyst. Nonetheless, they quickly show N-sensitive
segment addition and clearly do not have a large number of N-insensitive anterior
segments.
Parallel to these differences in where segmentation is affected, species vary as to
whether blocking N signaling disrupts normal growth. At one extreme, both
branchiopods and spiders show normal growth even when segmentation is affected. At
the other extreme, cockroaches and isopods show extreme growth defects in the N-
sensitive segments. In both cases, treated animals appear like growth chimeras with
normal anterior segments and highly reduced posterior segments. In sum, in no case is
there an overall diminishment of growth when N signaling is blocked. In some species
growth appears wholly unaffected; in other species growth defects are limited to the N-
sensitive segments.
In sum, a general evolutionary trend toward patterning fewer segments in a N
dependent manner within the Pan-Crustacea is evident, although the sample set is
arguably too small to lend strong support to this trend. However, beyond this observation,
the discrete effects of inhibiting N signaling in arthropods are not easily generalized and
the similarities that emerge are not easy to map phylogenetically.
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Does the data support homology of segmentation between arthropods and vertebrates?
Finding a function for N in arthropods has revitalized the argument for the homology of
segmentation between vertebrates and arthropods. To evaluate whether the evidence
supports a homologous segmentation process across these two phyla, we compare N
function in segmentation between vertebrates and arthropods.
Mesodermal vs. ectodermal segmentation
Any comparison of vertebrate and arthropod segmentation has to take in account
the cellular mechanisms by which segments form. The vertebrate process to which
arthropod segmentation is compared is somitogenesis or the segmentation of the
mesoderm. The tissue level behavior that N controls is a transition from mesenchymal
cells to epithelial cells during which mesenchymal cells in the PSM come together and
form an epithelial sac that forms the precursor to the segment. Failure to accomplish this
mesenchymal to epithelial transition halts segmentation and often gives rise to
incomplete regions of epithelialization (Burgess et al., 1996). In comparison to the
vertebrate mode of segmentation, analyses of arthropod segmentation are based primarily
on cell behaviors within an already formed epithelia, the ectodermal epithelium of the
growth zone. Comparisons between arthropods and vertebrates are further complicated
because the growth zones in the arthropod species sampled vary both in spatial
dimensions, timing of segment formation, and mode of assembly.
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Linearity of segmentation rates
Vertebrate segments are added at linear species exhibit species-specific rates.
These rates correlate with the periodicity of the gene expression cycle in the PSM in each
species. Do arthropods have species-specific rates of segment addition? Finding that
arthropod species share with vertebrates species specific, linear time courses of segment
addition would lend correlative support for a “clock” within arthropods. (Note that the
oscillatory mechanism that underlies vertebrate segmentation could still be present in
arthropods even in the absence of a linear periodicity). Although segmentation has been
described for a number of arthropods ((Damen, 2004; Davis et al., 2005; Manzanares et
al., 1996), there are typically no time data. Within the two anostracans we studied, there
are differences in the timing of normal segment addition. In Thamnocephalus, and the
notostracan Triops (Ko and Williams, in prep.), the temporal addition of segments is
approximately linear. Segmentation begins immediately upon hatching and proceeds at a
relatively constant rate. By contrast, although the morphological sequence of segment
addition in Artemia is linear, the temporal sequence is not. There is an initial delay after
hatching followed by a relatively linear rate of segment addition in the first 5 segments
then a rate about four times slower in subsequent segments (a rate that slows
progressively as more segments are added). This difference in Artemia between the
morphological versus temporal sequence in segment addition points to the relevance of
temporal data for studies of segmentation. The observation that the morphological
addition of segments is linear has led to the assumption that the temporal addition of
segments is also linear, which may not always be the case.
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The Clock-Wavefront model
The molecular basis of segmentation of the vertebrate presomitic mesoderm
(PSM) is characterized by synchronized waves of gene expression that pass from
posterior to anterior cells in a periodic manner. At the same time, the growth and
maturation of the determination front gives rise to a wave of somite formation from the
anterior to posterior. The interaction of these two phenomena is modeled by a
combination of a wave front passing anterior to posterior and a clock operating in the
PSM (see (Dubrulle and Pourquie, 2004a) for review). N pathway genes synchronize
oscillations of gene expression in cells in the PSM and establish boundaries between
adjacent somites (Barrantes et al., 1999; Jiang et al., 2000; Saga, 2007; Ozbudak and
Lewis, 2008). If N pathway genes are knocked out, segmentation proceeds normally until
7-9 segments form and then it becomes unsynchronized, and the boundaries between
segments are affected (Conlon et al, 1995; Takke and Campos-Ortega, 1999). As in
vertebrates, loss of N signaling in arthropods does not affect the anterior most segments.
However, in arthropods, loss of N signaling results in a loss of segments, not just the loss
of synchrony of segmentation as in vertebrates. Therefore in arthropods N signaling is
either required for the initiation or maintenance of segmentation process. Despite these
differences, arthropod data does not discount the presence of a clock segmentation
mechanism similar to that in vertebrates. Although oscillating expression of signaling
molecules in arthropods has not yet been described, the regular oscillations of Hairy, a
downstream target of Notch signaling in vertebrate somitogenesis, were originally found
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through a rigorous comparison of the timing of gene expression patterns between
dissected lateral halves of embryos allowed to develop in culture for different times
(Palmeirim et al, 1997). This type of analysis has not yet been performed on any
arthropod species.
Furthermore, genes of the Wnt and FGF signaling pathways also oscillate in the
PSM and have been shown to function in the wave front (Aulehla et al, 2003; Dequeant
et al, 2006). As the wave front passes posterior to anterior, it signals to determine which
region of the elongating embryo will form a somite. Cells are able to form somites only
if they are in the correct phase of the clock. There are no reports of investigation into the
potential involvement of FGF in arthropod segmentation. Recently, Wnt8 has been
shown to control the formation of the posterior growth zone in Achaearanea (McGregor
et al., 2008). Loss of Wnt8 function results by RNAi results in the absence of not only
the posterior segments, but the posterior of the embryo. Furthermore, loss of Wnt8
function in the spider affects the expression of Delta, hairy and caudal in the posterior of
the embryo. As such, Wnt8 not only establishes the growth zone, but may play a
permissive role for segmentation in the growth zone. This data is supported by functional
studies in Oncopeltus, and Tribolium (Angelini and Kaufman, 2005; Bolognesi et al.,
2008). In both cases, wingless RNAi results in a complete loss of several posterior
segments. Moreover, wingless expression data the crustacean Triops longicaudatus, the
myriapod Lithobius, and the chelicerate Cupiennius, where wingless is not only expressed
in a segmentally iterated pattern, but also in a ring at the posterior of the growth zone,
65
supports this model (Damen, 2002; Hughes and Kaufman, 2002b; Nulsen and Nagy,
1999). Taken together, these data suggest that the mechanisms of arthropod
segmentation share many similarities to vertebrate somitogenesis.
Are the phenotypic effects of disrupting N signaling similar?
In parallel with vertebrates, numerous segments form normally in arthropod
species when N signaling is disrupted and segmentation can be halted. The affected
segments in both phyla show irregular boundaries. Nonetheless, irregular boundaries
formed in arthropods seem different in kind from the irregular boundaries formed in
vertebrates when N signaling is disrupted since segments can no longer be recognized
and counted.
In both arthropods and vertebrates, the initial patterning of segments precedes
assignment of positional identity by the Hox genes. However, Hox gene expression
precedes segment patterning. Interestingly, vertebrate studies have shown some link
between Hox gene activity and somitogenesis. For example, ectopic FGF8 activity alters
the axial somite position, and also results in a parallel shift in Hox expression (Dubrulle
et al, 2001). Furthermore, some alterations in Notch signaling affect the position of Hox
gene expression and may result in homeotic transformations (Zakarny et al, 2001; Cordes
et al; 2004). Similarly to vertebrates, expression of Ubx/abdA precedes segmentation in
Artemia (Averof and Akam, 1995), as is evident in the posterior growth zone in our
control animals. The posterior domain of Ubx/Abda expression is not lost in DAPT
66
treated larvae, but is lost in the region of the growth zone where segments fail to form,
suggesting that Notch signaling is necessary for transition of a ubiquitous expression of
Ubx/Abda in the growth zone, to a segmentally iterated pattern of expression. As such,
the anterior two to three segments that form with drug treatment, presumably because
they are patterned before the larva hatches out of the cyst and hence before it is exposed
to DAPT, Ubx/abdA expression is comparable to untreated larvae. Although the Hox
genes are not thought to play a role in generating the segments themselves in any
arthropod (Averof and Akam, 1993; Abzhanov and Kaufman, 2000; Galant and Carroll,
2002), our data supports some coordination between Notch signaling and Hox expression
during arthropod segmentation.
Conclusions
There are some interesting differences in the effects of Notch on segmentation in
arthropods. First of all, whereas we did not detect any effects on segment borders on
segments that formed with DAPT treatment in Artemia and Thamnocephalus, segment
borders are affected when Notch is knocked out in spiders (Stollewerk et al, 2003). It
also appears that segment borders are affected by Notch inhibition in cockroach (Pueyo et
al, 2008) and Parhyale (O’Day, 2006). However, differences in how these segment
borders are affected by Notch inhibition in these animals can not be considered
independently of effects on growth, since some of the border effects may be due to
decreases in growth. We do not see effects on growth in Artemia and Thamnocephalus.
However, DAPT treatment affected growth in Parhyale (O’Day, 2006), just as Notch
67
RNAi affects growth in spiders (Stollewerk et al, 2003; and Oda et al, 2007) and the
cockroach (Pueyo et al, 2008). Together, these data suggest that Notch involvement in
growth and segmentation may be ancestral to the arthropods. Furthermore, the
involvement of Notch signaling on growth may have been lost at some point in the
evolutionary history of branchiopods, after they diverged from malacostracans, while its
involvement in segmentation remained conserved.
Second, the number of segments that remain unaffected when Notch is inhibited is
variable. In spiders and crustaceans, only the first few thoracic segments are not affected
by loss of Notch signaling (Stollewerk et al, 2003; Oda et al, 2007). However, in the
cockroach maternal Notch RNAi does not affect the formation of segments T1-A3, and
only segments posterior to A3 are affected by the loss of Notch signaling (Pueyo et al,
2008). Cockroach thoracic segments are therefore likely not under the control of Notch
signaling. This suggests that the last common ancestor to insects and crustaceans most
likely used Notch signaling to pattern its thoracic and abdominal segments and that this
was slowly lost in insects over time. This is consistent with the fact that Notch is used to
pattern abdominal segments in a basal insect such as the cockroach, but the Notch
signaling does not seem to be used at all in segmentation in more derived insects like
Tribolium and Drosophila (Eckert et al, 2004; Sommer and Tautz, 1993; Tautz; 2004).
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Figure 2.1: (A) The Artemia larva hatches without trunk segments. Segments are then added one at a time from anterior to posterior. Since segments continue to develop after they are initially established, anterior segments have progressed farther in development than their posterior counterparts. (B) The adult brine shrimp has 11 thoracic segments, 2 genital segments, and 5 post-genital segments.
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70
Figure 2.2: (A) Timing of the addition of Artemia segments, as measured by the appearance of EN stripes. There is no EN expression in Artemia for the first 10 hours of development. Temperature ( ) had no effect on the rate of segmentation while high density ( ) slowed segment addition. The insert in (A) shows the timing of addition of Thamnocephalus segments, demonstrating the similarity in the rate of segment addition of the anterior Artemia segments to the rate of Thamnocephalus segmentation. (B) Timing of addition of Thamnocephalus segments. Larvae hatch with at least two EN stripes and add segments at a fairly linear rate. Morphological segments were scored based on the symmetry and regularity of the posterior boundary of the segment, and the posterior-most full segment was the last segment counted.
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Figure 2.3: DAPT affects segmentation in Artemia in a dose dependent manner. (A-C) Animals after 96 hrs of continuous DAPT treatment grown in (A) sea water + DMSO: (B) 50 µM DAPT, and (C) 100 µM DAPT. Both control and drug treated animals were grown at high density. The gnathal segments and corresponding EN stripes are labeled ‘gn’, while thoracic segments are labeled T1-T6. The data for three separate replicates was combined for the final results. Addition of segments slows with DAPT treatment in a concentration dependent manner in both (D) Artemia and (E) Thamnocephalus.
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Figure 2.4: DAPT treatment does not affect growth in (A) Artemia, and causes only a small decrease in body size in (B) Thamnocephalus.
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Figure 2.5: Ubx/abdA expression in the posterior growth zone of DAPT treated larvae. (A) Untreated 96 hour animals. (B) 100 µM DAPT treated animals. Numbers designate the thoracic segments.
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Table 2.1: The comparison of the effects of the loss of Notch signaling on segment patterning in vertebrate and arthropod species.
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CHAPTER 3: SUMMARY OF CONCLUSIONS AND FUTURE DIRECTIONS
Since γ-secretase inhibits the formation of segments in Artemia and Thamnocephalus, it
is likely that Notch signaling plays a role in segmentation in these two branchiopod
crustaceans. Experiments that inhibit Notch signaling during segmentation, such as
RNAi, can verify this result. In light of this finding, it would be interesting to see the
expression of Notch signaling targets, such as hairy. As previously discussed, hairy is
part of a molecular clock in chordates, and its expression oscillates along the PSM in time
with the addition of segments (Paleirim et al, 1997; Bessho et al, 2003; Holley et al,
2000; Oates and Ho, 2002). If such a molecular clock exists in Artemia, a dynamic
expression pattern of Notch pathway components is expected.
Growth of the Artemia larva was not affected by γ-secretase inhibitor treatment, as the
treated larvae continued to grow at the same rate as the controls. These data suggest that
even though growth is necessary for the formation of segments, the two processes are
regulated independently. Interestingly, window experiments suggest that in the presence
of growth but absence of segmentation, cells remain in a permissive state, allowing
segmentation to resume when γ-secretase mediated signaling is restored. The window
treatment experiments were problematic, because complete elimination of the drug post
treatment could not be confirmed. However, this approach can be used to establish the
timing between segment determination and segment formation by treating the larvae with
the drug at progressively later times and assaying the affected segments.
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In sequentially segmented arthropods, segments are added one at a time from anterior to
posterior as the larva develops. The timing of the addition of segments has not been
assayed previously, and was assumed to be linear. In the case of Thamnacephalus, this
assumption is correct, as segments are added at regular intervals. However, Artemia
segments are not added at regular intervals. It appears that the addition of the first 5
thoracic segments is linear, but the subsequent segments are added at much longer
intervals. It is possible that this slowing of segmentation in Artemia is due to nutritional
deficiencies during the incubation period. However, this is unlikely due to depleting food
sources as food was refreshed daily and there is no corresponding daily spike in the rate
of segment addition in response to fresh food. It is also a possibility that there is a
naturally occurring supplement that Artemia larvae need as they continue to develop,
which is not as abundant or even absent in lab media. Growing Artemia larvae in sea
water from sources where these larvae naturally occur would address this issue.
Therefore, to gain an accurate understanding of the rates of segmentation in Artemia, and
confirm that segmentation is indeed not linear, it would be best to grow larvae in mesh
enclosures in their natural habitat.
Interestingly, small changes in temperature do not affect the rate of Artemia
segmentation, which is consistent with previous findings that the growth of these larvae is
quite resilient to shifts in temperature and salinity (Vanhaecke, 1984; Wear, 1986). This
is not surprising considering that Artemia is an ectothermic organism, and hence relies on
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environmental conditions for optimal developmental temperatures. Hence, such an
organism would have to be adapted to withstand shifts in temperatures to ensure its
survival. By contrast, endothermic larvae develop in very consistent temperature
conditions, are not adapted to withstand temperature shifts, and hence are extremely
susceptible to temperature variation during development (Feast, 1998).
The finding that Notch may be involved in the patterning of segments in branchiopod
crustaceans, as well as similar findings in other arthropods, suggest that the common
ancestor to arthropods was not only segmented, but also likely utilized Notch signaling to
pattern segments. The fact that Notch signaling is also at the heart of vertebrate
segmentation strengthens the argument that the common bilaterian ancestor was
segmented. In addition to offering more evidence to the debate surrounding the evolution
of segments in the three segmented phyla, comparative experiments can also be useful to
the overall understanding of segmentation mechanisms in all segmented species.
However the debate regarding whether the Urbilaterian was in fact segmented is far from
over. At this time, there are certainly more differences between the molecular
mechanisms of segmentation between the segmented phyla than there are similarities.
For example, evidence for the involvement of Wnt and FGG signaling in arthropod
segmentation is lacking, even though these two pathways play such an integral role in
vertebrate segmentation. If these two pathways are in fact not involved in the
segmentation of most arthropods, then it is likely that more differences than similarities
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will be found between segmentation mechanisms of these two phyla. If this is the case,
then it is likely that the Notch signaling pathway was co-opted to pattern segments in
arthropods independently of its use to pattern chordate segments.
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APPENDIX A: MORPHOLOGY OF SEGMENTATION
Trunk Segments form sequentially from Anterior to Posterior
Based on these detailed descriptions of cell rearrangements during the early stages of
segment patterning (Freeman, 1986; Freeman et al, 1992) I was able to determine that the
anterior-most trunk segments are added sequentially. Artemia larvae hatch out of the
dehydrated cyst with three out of five head segments and no morphologically visible
trunk segments and no EN expression corresponding to the trunk segments (Figure
A1.1A). The anterior nuclei in the trunk form columns along the anterior-posterior axis
of the trunk, which precedes the first evidence of segmentation (Freeman, 1992). As the
larva continues to grow and develop, the first morphological evidence of segmentation is
the arrangement of nuclei in rows across the trunk, which is followed by expression of
EN in the forming segment. At 4 hours after hatching, the cells in the anterior-most
region of the trunk begin to show the first morphological evidence of segment patterning
– the nuclei arrange into rows across the trunk (Figure A1.1B). 4 hours later, at 8 hours
after hatching, there are more rows of cells, indicating the first evidence of progressive
segmental patterning from anterior to posterior (Figure A1.1C). This suggests that
anterior segments are not patterned simultaneously, but instead are patterned sequentially,
with a short time interval between them. By 12 hours after hatching, the first trunk EN
stripe is clearly formed, and there is low level of EN expression where the second stripe
is forming (Figure A1.1D,E). The fact that EN does not come on at the same time in the
first two segments further implies that these two segments are indeed patterned
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consecutively and not simultaneously. The first two trunk segments show EN expression
at 18 hours after hatching (Figure A1.1F,G). These segments continue to develop as
more posterior trunk segments are added. As such, the anterior segments are more
developed than their posterior counterparts and are first to develop appendages (Figure
A.1.1C). Interestingly, the two posterior-most gnathal EN stripes appear more or less at
the same time as the first trunk stripe. That is, I have never seen a larva that has the
gnathal EN stripes, but not the first trunk stripe. This suggests that these three segments
are patterned at the same time. However, due to the overlap in tissues in the head region,
the nuclear morphology is difficult to resolve. In sum, our findings show that all Artemia
trunk segments are patterned sequentially from anterior to posterior. The EN stripes for
the first two segments come on in quick succession, but EN comes on in the first segment
before it does in the second. A close morphological analysis, more specifically nuclear
arrangements in the anterior trunk in the hours just after the larva hatches out of its cyst
confirms this observation. Nuclei are arranged in rows that form from anterior to
posterior over time, as opposed to being arranged simultaneously.
According to Freeman (1986), the first morphological evidence of an emerging segment
in Artemia is the reorganization of cells from columns that run along the anterior-
posterior axis of the posterior growth zone to rows of cells that are perpendicular to that
axis. These cells then are grouped together, and the ectoderm constricts around them in
the region where the limb buds will eventually form (Freeman, 1989). Previous studies
have shown that EN expression is initiated before the ectodermal constriction clearly
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delineates the morphological borders between segments (Manzanares et al, 1993).
However, we have shown that nuclear organization into rows precedes EN expression,
suggesting that EN is a downstream component of the regulatory network that patterns
Artemia segments. This is not surprising considering that EN has been shown to give
posterior identity to already patterned Drosophila embryonic segments, a role that seems
to have been conserved among all arthropods (Patel et al, 1989; Scholtz et al, 1994;
(Dearden and Akam, 2001; Hughes and Kaufman, 2002a)Janssen et al, 2004). The
appearance of ectodermal constrictions after EN expression is initiated suggests that EN
is involved in maintaining, and perhaps even delineating, the border between the
posterior of a newly formed segment, and the anterior of the next segment.
Segment Morphology Does Not Account for the Difference in the Time of Addition of
Trunk Segments
The first five segments are patterned on average every 7 hours. Segment 6 takes 18 hours
to develop, which is more than twice the time. There is not size difference between
segments 5 and 6 to account for such a drastic difference in the timing of their patterning
(Figure A1.2). Furthermore, nuclear organization proceeds as normal during the addition
of the 6th segment. Nuclei are organized in rows down to just posterior of the last stripe
of EN expression. Posterior to that, nuclei are organized in columns, which is
characteristic of unsegmented posterior growth zone just posterior of the last segment
(Freeman, 1992). In sum, there is no morphological evidence to account for the increase
in the time each segment is patterned segment 6 and all the remaining trunk segments.
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Figure A1.1: (A) Newly hatched larva has no evidence of trunk segments. Nuclei are not yet organized into rows across the trunk even in the most anterior region of the trunk. Instead, columns of nuclei instead extend to the anterior of the trunk (arrow). The bracket designates the trunk region pictured in the subsequent panels. (B) At 4 hours, anterior-most nuclei begin to organize into rows (arrow), but there is still no EN expression. (C) By 8 hours several more rows of nuclei suggesting initiation of segment patterning are apparent (arrows). (A’-C’) Duplicate images of A-C with overlaying colored dots to demarcate nuclei and show patterns. The purple dots overlay nuclei arranged in rows of cells as segment patterning begins, and the blue dots show columns of nuclei in the more posterior unsegmented region. (D) A 12 hour larva has its first trunk EN stripe and the second stripe begins to form. There is also EN expression in the gnathal region. Rows of nuclei span the anterior region of the trunk, until just posterior of the first trunk stripe (open arrow). Posterior to EN expression, nuclei are organized in columns (closed arrow). The bright field photo of this animal in panel (E) shows a clearer view of EN expression. (G) Bright field photo showing the EN expression in the 18 hour larva pictured in (F). At 18 hours the initial patterning of the first two trunk segments is complete. There are only 2 rows of nuclei in the trunk, just posterior to the posterior-most EN stripe (open arrow). Posterior to that, nuclei are organized in columns (closed arrow). All embryos were stained with EN and the nuclei were marked with DAPI.
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89
Figure A1.2: There is no morphological difference between segments five and six to account for the increased length of time that it takes to form segment six. (A) Larva with 5 EN stripes and early EN expression in the 6th segment (arrow). (B) Higher magnified view of DAPI stained trunk of the larva in panel A. (C) Larva with 6 EN stripes – the 5th and 6th segments, and corresponding EN stripes do not vary in size or cellular organization. (D) DAPI staining of trunk of larva in panel C showing nuclear organization.
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APPENDIX B: MORPHOLOGY OF DAPT MUTANTS
DAPT treatment disrupts cellular organization within the growth zone
In untreated Artemia larvae, the nuclear organization of the growth zone posterior
to the last segment can be split into three distinct domains (Figure A2.1A) (Freeman,
1986). The posterior-most nuclei do not exhibit any organization, more anterior nuclei
are organized into columns and nuclei just posterior to the last morphologically
detectable segment are organized into rows (Figure A2.1A). Thus segmentation is pre-
figured by the organization of the cells. The onset of EN expression also occurs prior to
any visible manifestation of segmentation and is detected first in the cells that have
organized into rows. I looked at the organization of nuclei in the posterior growth zone
to determine whether DAPT treatment affects the formation of segments, and not just EN
expression. DAPT treated larvae not only have fewer EN stripes, but also do not show
the organization of cells into rows, which suggests that no segments are patterned
posterior to the last EN stripe that forms (Figure A2.1B). There are only a few rows of
cells past the posterior most EN stripe and in the remainder of the posterior trunk the cells
remain disorganized. Therefore, the decrease in EN stripes in DAPT treated larvae
corresponds to a loss of segments as well as the cellular organization that precedes
segmentation in the posterior growth zone, and not just a decrease in EN expression.
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Figure A2.1: Cellular organization in the posterior of DAPT treated larvae. (A) control and (B) 100 µM DAPT treated larvae. Open arrows indicate columns of cells in the unsegmented posterior growth zone. Filled arrows indicate rows of cells posterior to the last distinguishable segmented region. (C and D) Duplicate images of A and B with overlaying colored dots to demarcate nuclei and show patterns. The purple dots overlay nuclei arranged in rows of cells as segment patterning begins posterior to the last patterned segment and the blue dots show columns of nuclei in the more posterior unsegmented region. The dashed lines mark borders of already patterned segments in the anterior of the growth zone.
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94
APPENDIX C: PARTIAL SEQUENCE OF ARTEMIA NOTCH
Notch Cloning
Artemia Notch was cloned from a pool of Artemia embryos ranging in age from newly
hatched to 96 hours after hatching using degenerate primers (5’
CCNGGNMYRCCGAAGTGGGGAGCTACG;
GGTGCTGCTGGACCACTTCKCNAAYMGNGA) designed by the COnsensus-
DEgenerate Hybrid Oligonucleotide Primers (CODEHOP) program (Rose et al., 2003).
95
Figure A3.1: Partial Artemia Notch amino acid sequence the same portion of the sequence of the protein in other invertebrates.
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APPENDIX D: WINDOW TREATMENTS OF ARTEMIA LARVAE WITH DAPT
Since continuous DAPT treatment results in a severe slowing down and in some cases
arrest of segmentation, I did several window treatments to determine whether
segmentation would resume when the drug was removed.
In these series of experiments, larvae were exposed to a 100µM DAPT treatment as
described in Chapter 2. However, after 12 hours of treatment, the drug was washed out
with 3 quick rinses and 3 twenty minute incubations with FASW at 21°C. After washing,
the larvae were incubated in FASW supplemented with algae and yeast and grown at
21°C as described in Chapter 2. At each sampling, larvae were fixed and stained for EN
expression as previously described in Chapter 2. EN stripes were used as markers of
segments along the growth zone. The data shown is a pool of three separate experiments
since the results were consistent between trials.
The Artemia larvae treated with DAPT for 12 hours had fewer segments than the controls
(Figure A4.1). However, these larvae had fewer segments than those treated
continuously with DAPT. This indicates that the short window treatment with DAPT had
an overall effect on the timing of segmentation. One limitation of this experiment is that
the effectiveness of DAPT removal from the larvae can not be assayed. As such, it is
unclear whether the results are indeed due solely to a short exposure to a high
concentration of DAPT, or both a short exposure and a subsequent longer exposure to
low DAPT levels of drug that remained inside the cells after washing. This problem can
98
be addressed in the future with a western blot analysis of the abundance of the Notch
intracellular domain in the treatment larvae after washing compared to the control larvae,
showing the relative levels of Notch activity, once an antibody to this region of the
Artemia protein becomes available.
Assuming that all traces of the drug were washed away at the end of the window
treatment, segmentation continues from where it left off once Notch signaling resumes.
This is an interesting finding considering that DAPT does not affect growth and cell
division. As such, in DAPT treated larvae, cells continue to divide, but their patterning
is delayed requiring them to remain undifferentiated for a longer period of time than in
untreated larvae. It is clear, therefore, that the maximum time that a cell in the growth
zone can remain undifferentiated was not challenged by this experiment. As such, this
experiment suggests that the permissive region for segmentation would span more of the
growth zone in the DAPT treated larvae than in the controls. If this was not the case, and
the cells only had a set period of time to differentiate and form segments, then a gap
between segments or missing segments along the growth zone would be expected. In this
scenario, segmentation would be halted due to the presence of a high concentration of
DAPT, and then resume at the end of the treatment at the segment that was to be
patterned at that time point, resulting in skipped segments. No such segment gaps were
observed.
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Figure A4.1: 12 hour DAPT treatment window at 12-24 hours after hatching (by the red bar), affects the number of segments (as counted by the presence of EN stripes) at 96 hours after hatching. Even though the DAPT treated larvae had the same number of segments as controls at the end of treatment (24 hour time point), they had significantly fewer segments at 72 and 96 hours as compared to the controls. (n = sample size; # = number of EN stripes; s.d. = standard deviation).
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