Molecular and Morphological Characterizationof Segmentation in Artemia franciscana

Item Type text; Electronic Dissertation

Authors Blachuta, Beata J

Publisher The University of Arizona.

Rights Copyright © is held by the author. Digital access to this materialis made possible by the University Libraries, University of Arizona.Further transmission, reproduction or presentation (such aspublic display or performance) of protected items is prohibitedexcept with permission of the author.

Download date 19/05/2018 17:21:15

Link to Item http://hdl.handle.net/10150/194771

1

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

2

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

3

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

4

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.

5

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

6

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

7

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.

8

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

9

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

10

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

11

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

12

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

13

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

14

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.,

15

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

16

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

17

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

18

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

19

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

20

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

21

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.

57

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.

58

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

60

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.

61

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.

62

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.

63

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

64

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).

68

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.

69

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.

71

72

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.

73

74

Figure 2.4: DAPT treatment does not affect growth in (A) Artemia, and causes only a small decrease in body size in (B) Thamnocephalus.

75

76

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.

77

78

Table 2.1: The comparison of the effects of the loss of Notch signaling on segment patterning in vertebrate and arthropod species.

79

80

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.

81

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

82

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

83

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.

84

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

85

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

86

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.

87

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.

88

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.

90

91

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.

92

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.

93

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.

96

97

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.

99

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).

100

101

WORKS CITED Adoutte, A., Balavoine, G., Lartillot, N., and de Rosa, R. (1999). Animal evolution. The end of the intermediate taxa? Trends Genet 15, 104-108.

Anderson, D. T. (1967). Larval development and segment formation in the branchiopod crustaceans Limnadia stanleyana (Concostraca) and Artemia salina (Anostraca). Aust J Zool 15, 47-91.

Anderson, F. E., A. J. Cordoba, and M. Thollesson (2004). Bilaterian phylogeny based on analyses of a region of the sodium-potassium ATPase alpha-subunit gene. Journal of Molecular Evolution 58, 252-268.

Angelini, D. R., and Kaufman, T. C. (2005). Functional analyses in the milkweed bug Oncopeltus fasciatus (Hemiptera) support a role for Wnt signaling in body segmentation but not appendage development. Dev Biol 283, 409-423.

Aranda, M., Marques-Souza, H., Bayer, T., and Tautz, D. (2008). The role of the segmentation gene hairy in Tribolium. Dev Genes Evol 218, 465-477.

Artavanis-Tsakonas, S., Rand, M. D., and Lake, R. J. (1999). Notch signaling: cell fate control and signal integration in development. Science 284, 770-776.

Aulehla, A., Wehrle, C., Brand-Saberi, B., Kemler, R., Gossler, A., Kanzler, B., and Herrmann, B. G. (2003). Wnt3a plays a major role in the segmentation clock controlling somitogenesis. Dev Cell 4, 395-406.

Aulehla, A., Wiegraebe, W., Baubet, V., Wahl, M. B., Deng, C., Taketo, M., Lewandoski, M., and Pourquie, O. (2008). A beta-catenin gradient links the clock and wavefront systems in mouse embryo segmentation. Nat Cell Biol 10, 186-193.

Averof, M. a. A., M (1995). Insect-crustacean relationships: Insights from comparative developmental and molecular studies. Philos Trans R Soc Lond B Biol Sci 347, 293-303.

Baker, N. E. (1987). Molecular cloning of sequences from wingless, a segment polarity gene in Drosophila: the spatial distribution of a transcript in embryos. Embo J 6, 1765-1773.

Balavoine, G., Adoutte, A. (1998). One or three Cambrian radiations? Science 280, 397-398.

Barrantes, I. B., Elia, A. J., Wunsch, K., Hrabe de Angelis, M. H., Mak, T. W., Rossant, J., Conlon, R. A., Gossler, A., and de la Pompa, J. L. (1999). Interaction between Notch

102

signalling and Lunatic fringe during somite boundary formation in the mouse. Curr Biol 9, 470-480.

Bate, M., Martinez-Arias, A. M. (1993). The development of Drosophila melanogaster (Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press).

Bejsovec, A., and Martinez Arias, A. (1991). Roles of wingless in patterning the larval epidermis of Drosophila. Development 113, 471-485.

Bessho, Y., Hirata, H., Masamizu, Y., and Kageyama, R. (2003). Periodic repression by the bHLH factor Hes7 is an essential mechanism for the somite segmentation clock. Genes Dev 17, 1451-1456.

Bolognesi, R., Farzana, L., Fischer, T. D., and Brown, S. J. (2008). Multiple Wnt genes are required for segmentation in the short-germ embryo of Tribolium castaneum. Curr Biol 18, 1624-1629.

Bray, S. (1998). A Notch affair. Cell 93, 499-503.

Brown, S., Fellers, J., Shippy, T., Denell, R., Stauber, M., and Schmidt-Ott, U. (2001). A strategy for mapping bicoid on the phylogenetic tree. Curr Biol 11, R43-44.

Brusca, R. C., Brusca, G.J. (1990). Invertebrates (Sunderland, Massachusetts: Sinauer Associates).

Bucher, G., Farzana, L., Brown, S. J., and Klingler, M. (2005). Anterior localization of maternal mRNAs in a short germ insect lacking bicoid. Evol Dev 7, 142-149.

Bucher, G., and Klingler, M. (2004). Divergent segmentation mechanism in the short germ insect Tribolium revealed by giant expression and function. Development 131, 1729-1740.

Burgess, R., Rawls, A., Brown, D., Bradley, A., and Olson, E. N. (1996). Requirement of the paraxis gene for somite formation and musculoskeletal patterning. Nature 384, 570-573.

Calvo, E., Walter, M., Adelman, Z. N., Jimenez, A., Onal, S., Marinotti, O., and James, A. A. (2005). Nanos (nos) genes of the vector mosquitoes, Anopheles gambiae, Anopheles stephensi and Aedes aegypti. Insect Biochem Mol Biol 35, 789-798.

Cambray, N., and Wilson, V. (2007). Two distinct sources for a population of maturing axial progenitors. Development 134, 2829-2840.

Campos-Ortega, J. A. a. H., V. (1985). The embryonic development of Drosophila melanogaster (New York, NY: Springer-Verlag).

103

Carroll, S. B., and Scott, M. P. (1986). Zygotically active genes that affect the spatial expression of the fushi tarazu segmentation gene during early Drosophila embryogenesis. Cell 45, 113-126.

Cerny, A. C., Bucher, G., Schroder, R., and Klingler, M. (2005). Breakdown of abdominal patterning in the Tribolium Kruppel mutant jaws. Development 132, 5353-5363.

Chipman, A. D., and Akam, M. (2008). The segmentation cascade in the centipede Strigamia maritima: involvement of the Notch pathway and pair-rule gene homologues. Dev Biol 319, 160-169.

Chipman, A. D., Arthur, W., and Akam, M. (2004). A double segment periodicity underlies segment generation in centipede development. Curr Biol 14, 1250-1255.

Chipman, A. D., and Stollewerk, A. (2006). Specification of neural precursor identity in the geophilomorph centipede Strigamia maritima. Dev Biol 290, 337-350.

Choe, C. P., Miller, S. C., and Brown, S. J. (2006). A pair-rule gene circuit defines segments sequentially in the short-germ insect Tribolium castaneum. Proc Natl Acad Sci U S A 103, 6560-6564.

Clark, R. B. (1981). Locomotion and the phylogeny of Metazoa. Bolletino di Zoologia 48, 11-28.

Conlon, R. A., Reaume, A. G., and Rossant, J. (1995). Notch1 is required for the coordinate segmentation of somites. Development 121, 1533-1545.

Cook, C. E., Smith, M. L., Telford, M. J., Bastianello, A., and Akam, M. (2001). Hox genes and the phylogeny of the arthropods. Curr Biol 11, 759-763.

Cooke, J., and Zeeman, E. C. (1976). A clock and wavefront model for control of the number of repeated structures during animal morphogenesis. J Theor Biol 58, 455-476.

Copf, T., Schroder, R., and Averof, M. (2004). Ancestral role of caudal genes in axis elongation and segmentation. Proc Natl Acad Sci U S A 101, 17711-17715.

Cordes, R., Schuster-Gossler, K., Serth, K., and Gossler, A. (2004). Specification of vertebral identity is coupled to Notch signalling and the segmentation clock. Development 131, 1221-1233.

Dale, J. K., Malapert, P., Chal, J., Vilhais-Neto, G., Maroto, M., Johnson, T., Jayasinghe, S., Trainor, P., Herrmann, B., and Pourquie, O. (2006). Oscillations of the snail genes in the presomitic mesoderm coordinate segmental patterning and morphogenesis in vertebrate somitogenesis. Dev Cell 10, 355-366.

104

Damen, W. G. (2002). Parasegmental organization of the spider embryo implies that the parasegment is an evolutionary conserved entity in arthropod embryogenesis. Development 129, 1239-1250.

Damen, W. G. (2004). Arthropod segmentation: why centipedes are odd. Curr Biol 14, R557-559.

Damen, W. G. (2007). Evolutionary conservation and divergence of the segmentation process in arthropods. Dev Dyn 236, 1379-1391.

Damen, W. G., Janssen, R., and Prpic, N. M. (2005). Pair rule gene orthologs in spider segmentation. Evol Dev 7, 618-628.

Damen, W. G., Weller, M., and Tautz, D. (2000). Expression patterns of hairy, even-skipped, and runt in the spider Cupiennius salei imply that these genes were segmentation genes in a basal arthropod. Proc Natl Acad Sci U S A 97, 4515-4519.

Davis, G. K., D'Alessio, J. A., and Patel, N. H. (2005). Pax3/7 genes reveal conservation and divergence in the arthropod segmentation hierarchy. Dev Biol 285, 169-184.

Davis, G. K., Jaramillo, C. A., and Patel, N. H. (2001). Pax group III genes and the evolution of insect pair-rule patterning. Development 128, 3445-3458.

Davis, G. K., and Patel, N. H. (2002). Short, long, and beyond: molecular and embryological approaches to insect segmentation. Annu Rev Entomol 47, 669-699.

Dawes, R., Dawson, I., Falciani, F., Tear, G., and Akam, M. (1994). Dax, a locust Hox gene related to fushi-tarazu but showing no pair-rule expression. Development 120, 1561-1572.

De Robertis, E. M., and Sasai, Y. (1996). A common plan for dorsoventral patterning in Bilateria. Nature 380, 37-40.

de Rosa, R., Grenier, J. K., Andreeva, T., Cook, C. E., Adoutte, A., Akam, M., Carroll, S. B., and Balavoine, G. (1999). Hox genes in brachiopods and priapulids and protostome evolution. Nature 399, 772-776.

de Rosa, R., Prud'homme, B., and Balavoine, G. (2005). Caudal and even-skipped in the annelid Platynereis dumerilii and the ancestry of posterior growth. Evol Dev 7, 574-587.

Dearden, P., and Akam, M. (2000). A role for Fringe in segment morphogenesis but not segment formation in the grasshopper, Schistocerca gregaria. Dev Genes Evol 210, 329-336.

105

Dearden, P. K., and Akam, M. (2001). Early embryo patterning in the grasshopper, Schistocerca gregaria: wingless, decapentaplegic and caudal expression. Development 128, 3435-3444.

Dearden, P. K., Donly, C., and Grbic, M. (2002). Expression of pair-rule gene homologues in a chelicerate: early patterning of the two-spotted spider mite Tetranychus urticae. Development 129, 5461-5472.

Delfini, M. C., Dubrulle, J., Malapert, P., Chal, J., and Pourquie, O. (2005). Control of the segmentation process by graded MAPK/ERK activation in the chick embryo. Proc Natl Acad Sci U S A 102, 11343-11348.

Dequeant, M. L., Glynn, E., Gaudenz, K., Wahl, M., Chen, J., Mushegian, A., and Pourquie, O. (2006). A complex oscillating network of signaling genes underlies the mouse segmentation clock. Science 314, 1595-1598.

Dequeant, M. L., and Pourquie, O. (2008). Segmental patterning of the vertebrate embryonic axis. Nat Rev Genet 9, 370-382.

Deutsch, J. S. (2004). Segments and parasegments in arthropods: a functional perspective. Bioessays 26, 1117-1125.

DiNardo, S., Kuner, J. M., Theis, J., and O'Farrell, P. H. (1985). Development of embryonic pattern in D. melanogaster as revealed by accumulation of the nuclear engrailed protein. Cell 43, 59-69.

DiNardo, S., and O'Farrell, P. H. (1987). Establishment and refinement of segmental pattern in the Drosophila embryo: spatial control of engrailed expression by pair-rule genes. Genes Dev 1, 1212-1225.

Driever, W., Ma, J., Nusslein-Volhard, C., and Ptashne, M. (1989). Rescue of bicoid mutant Drosophila embryos by bicoid fusion proteins containing heterologous activating sequences. Nature 342, 149-154.

Driever, W., and Nusslein-Volhard, C. (1989). The bicoid protein is a positive regulator of hunchback transcription in the early Drosophila embryo. Nature 337, 138-143.

Dubrulle, J., McGrew, M. J., and Pourquie, O. (2001). FGF signaling controls somite boundary position and regulates segmentation clock control of spatiotemporal Hox gene activation. Cell 106, 219-232.

Dubrulle, J., and Pourquie, O. (2004a). Coupling segmentation to axis formation. Development 131, 5783-5793.

106

Dubrulle, J., and Pourquie, O. (2004b). fgf8 mRNA decay establishes a gradient that couples axial elongation to patterning in the vertebrate embryo. Nature 427, 419-422.

Duman-Scheel, M., Pirkl, N., and Patel, N. H. (2002). Analysis of the expression pattern of Mysidium columbiae wingless provides evidence for conserved mesodermal and retinal patterning processes among insects and crustaceans. Dev Genes Evol 212, 114-123.

Dumont, H. J. a. N., SV (2002). Introduction to the Class Branchiopoda. (Leiden: Backhuys Publishers).

Dunty, W. C., Jr., Biris, K. K., Chalamalasetty, R. B., Taketo, M. M., Lewandoski, M., and Yamaguchi, T. P. (2008). Wnt3a/beta-catenin signaling controls posterior body development by coordinating mesoderm formation and segmentation. Development 135, 85-94.

Eckert, C., Aranda, M., Wolff, C., and Tautz, D. (2004). Separable stripe enhancer elements for the pair-rule gene hairy in the beetle Tribolium. EMBO Rep 5, 638-642.

Eernisse, D. J., Albert, J. S., Anderson, F. E. (1992). Annelida and Arthropoda are not sister taxa: A phylogenetic analysis of spiralian metazoan morphology. . Systematic Biology 41.

Eldon, E. D., and Pirrotta, V. (1991). Interactions of the Drosophila gap gene giant with maternal and zygotic pattern-forming genes. Development 111, 367-378.

Feast, M., Noble, R. C., Speake, B. K., and Ferguson, W. J. (1998). The effect of temporary reductions in incubation temperature on growth characteristics and lipid utilisation in the chick embryo. Journal of Anatomy 193, 383-390.

Forsberg, H., Crozet, F., and Brown, N. A. (1998). Waves of mouse Lunatic fringe expression, in four-hour cycles at two-hour intervals, precede somite boundary formation. Curr Biol 8, 1027-1030.

Freeman, J. A. (1986). Epidermal cell proliferation during thoracic development in larvae of Artemia. j crust biol 6, 37-48.

Freeman, J. A. (1989). Segment morphogenesis in Artemia larvae. (New York: Plenum Press).

Freeman, J. A., Cheshire, L. B., and Macrae, T. H. (1992). Epithelial morphogenesis in developing artemia the role of cell replication cell shape change and the cytoskeleton, Vol 152).

107

Frigerio, G., Burri, M., Bopp, D., Baumgartner, S., and Noll, M. (1986). Structure of the segmentation gene paired and the Drosophila PRD gene set as part of a gene network. Cell 47, 735-746.

Geling, A., Steiner, H., Willem, M., Bally-Cuif, L., and Haass, C. (2002). A gamma-secretase inhibitor blocks Notch signaling in vivo and causes a severe neurogenic phenotype in zebrafish. EMBO Rep 3, 688-694.

Gergen, J. P., and Butler, B. A. (1988). Isolation of the Drosophila segmentation gene runt and analysis of its expression during embryogenesis. Genes Dev 2, 1179-1193.

Goldbeter, A., Gonze, D., and Pourquie, O. (2007). Sharp developmental thresholds defined through bistability by antagonistic gradients of retinoic acid and FGF signaling. Dev Dyn 236, 1495-1508.

Gomez, C., Ozbudak, E. M., Wunderlich, J., Baumann, D., Lewis, J., and Pourquie, O. (2008). Control of segment number in vertebrate embryos. Nature 454, 335-339.

Goto, T., Macdonald, P., and Maniatis, T. (1989). Early and late periodic patterns of even skipped expression are controlled by distinct regulatory elements that respond to different spatial cues. Cell 57, 413-422.

Grobben, K. (1908). Die systematiche Einteilung des Tierreisches. Verh Zool Bot Ges Wien 58, 491-511.

Hader, T., La Rosee, A., Ziebold, U., Busch, M., Taubert, H., Jackle, H., and Rivera-Pomar, R. (1998). Activation of posterior pair-rule stripe expression in response to maternal caudal and zygotic knirps activities. Mech Dev 71, 177-186.

Hafen, E., Kuroiwa, A., and Gehring, W. J. (1984). Spatial distribution of transcripts from the segmentation gene fushi tarazu during Drosophila embryonic development. Cell 37, 833-841.

Harding, K., Hoey, T., Warrior, R., and Levine, M. (1989). Autoregulatory and gap gene response elements of the even-skipped promoter of Drosophila. Embo J 8, 1205-1212.

Holley, S. A., Geisler, R., and Nusslein-Volhard, C. (2000). Control of her1 expression during zebrafish somitogenesis by a delta-dependent oscillator and an independent wave-front activity. Genes Dev 14, 1678-1690.

Holley, S. A., Julich, D., Rauch, G. J., Geisler, R., and Nusslein-Volhard, C. (2002). her1 and the notch pathway function within the oscillator mechanism that regulates zebrafish somitogenesis. Development 129, 1175-1183.

108

Holley, S. A., and Takeda, H. (2002). Catching a wave: the oscillator and wavefront that create the zebrafish somite. Semin Cell Dev Biol 13, 481-488.

Horikawa, K., Ishimatsu, K., Yoshimoto, E., Kondo, S., and Takeda, H. (2006). Noise-resistant and synchronized oscillation of the segmentation clock. Nature 441, 719-723.

Howard, K., Ingham, P., and Rushlow, C. (1988). Region-specific alleles of the Drosophila segmentation gene hairy. Genes Dev 2, 1037-1046.

Hrabé de Angelis, M., McIntyre, J. and Gossler, A. (1997). Maintenance of somite borders in mice requires the Delta homologue Dll1. Nature 386, 717–721.

Hughes, C. L., and Kaufman, T. C. (2002a). Exploring myriapod segmentation: The expression patterns of even-skipped, engrailed, and wingless in a centipede. Developmental Biology 247, 47-61.

Hughes, C. L., and Kaufman, T. C. (2002b). Exploring myriapod segmentation: the expression patterns of even-skipped, engrailed, and wingless in a centipede. Dev Biol 247, 47-61.

Hughes, C. L., and Kaufman, T. C. (2002c). Exploring the myriapod body plan: expression patterns of the ten Hox genes in a centipede. Development 129, 1225-1238.

Hughes, C. L., and Kaufman, T. C. (2002d). Hox genes and the evolution of the arthropod body plan. Evol Dev 4, 459-499.

Hughes, C. L., Liu, P. Z., and Kaufman, T. C. (2004). Expression patterns of the rogue Hox genes Hox3/zen and fushi tarazu in the apterygote insect Thermobia domestica. Evol Dev 6, 393-401.

Huppert, S. S., Ilagan, M. X., De Strooper, B., and Kopan, R. (2005). Analysis of Notch function in presomitic mesoderm suggests a gamma-secretase-independent role for presenilins in somite differentiation. Dev Cell 8, 677-688.

Ishikawa, A., Kitajima, S., Takahashi, Y., Kokubo, H., Kanno, J., Inoue, T., and Saga, Y. (2004). Mouse Nkd1, a Wnt antagonist, exhibits oscillatory gene expression in the PSM under the control of Notch signaling. Mech Dev 121, 1443-1453.

Iwasa, J. H., Suver, D. W., and Savage, R. M. (2000). The leech hunchback protein is expressed in the epithelium and CNS but not in the segmental precursor lineages. Dev Genes Evol 210, 277-288.

Janssen, R., Budd, G. E., Damen, W. G., and Prpic, N. M. (2008). Evidence for Wg-independent tergite boundary formation in the millipede Glomeris marginata. Dev Genes Evol 218, 361-370.

109

Janssen, R., and Damen, W. G. (2006). The ten Hox genes of the millipede Glomeris marginata. Dev Genes Evol 216, 451-465.

Janssen, R., Prpic, N-M, Damen, WGM. (2004). Gene expression sugest decoupled dorsol and ventral segmentation in the milipede Glomeris marginata. Dev Biol 268, 89-104.

Jiang, Y. J., Aerne, B. L., Smithers, L., Haddon, C., Ish-Horowicz, D., and Lewis, J. (2000). Notch signalling and the synchronization of the somite segmentation clock. Nature 408, 475-479.

Jouve, C., Palmeirim, I., Henrique, D., Beckers, J., Gossler, A., Ish-Horowicz, D., and Pourquie, O. (2000). Notch signalling is required for cyclic expression of the hairy-like gene HES1 in the presomitic mesoderm. Development 127, 1421-1429.

Kadesch, T. (2004). Notch signaling: the demise of elegant simplicity. Curr Opin Genet Dev 14, 506-512.

Kadner, D., and Stollewerk, A. (2004). Neurogenesis in the chilopod Lithobius forficatus suggests more similarities to chelicerates than to insects. Dev Genes Evol 214, 367-379.

Kanki, J. P., and Ho, R. K. (1997). The development of the posterior body in zebrafish. Development 124, 881-893.

Kelsh, R., Weinzierl, R. O., White, R. A., and Akam, M. (1994). Homeotic gene expression in the locust Schistocerca: an antibody that detects conserved epitopes in Ultrabithorax and abdominal-A proteins. Dev Genet 15, 19-31.

Kilchherr, F., Baumgartner, S., Bopp, S., Frei, E., Noll, M (1986). Isolation of the paired gene of Drosophila and its spatial expression during early embryogenesis. Nature 321, 493-499.

Kimmel, C. B. (1996). Was Urbilateria segmented? Trends Genet 12, 329-331.

Knipple, D. C., Seifert, E., Rosenberg, U. B., Preiss, A., and Jackle, H. (1985). Spatial and temporal patterns of Kruppel gene expression in early Drosophila embryos. Nature 317, 40-44.

Kontarakis, Z., Copf, T., and Averof, M. (2006). Expression of hunchback during trunk segmentation in the branchiopod crustacean Artemia franciscana. Dev Genes Evol 216, 89-93.

Kornberg, T., Siden, I., O'Farrell, P., and Simon, M. (1985). The engrailed locus of Drosophila: in situ localization of transcripts reveals compartment-specific expression. Cell 40, 45-53.

110

Krumlauf, R. (1994). Hox genes in vertebrate development. Cell 78, 191-201.

Lai, E. C. (2004). Notch signaling: control of cell communication and cell fate. Development 131, 965-973.

Lall, S., Ludwig, M. Z., and Patel, N. H. (2003). Nanos plays a conserved role in axial patterning outside of the Diptera. Curr Biol 13, 224-229.

Lehmann, R., Nusslein-Volhard, C. (1984). Hunchback, a gene required for segmentation of an anterior and posterior region of the embryo. Developmental Biology 119, 402-417.

Lewis, J. (2003). Autoinhibition with transcriptional delay: a simple mechanism for the zebrafish somitogenesis oscillator. Curr Biol 13, 1398-1408.

Liaw, G. J., and Lengyel, J. A. (1993). Control of tailless expression by bicoid, dorsal and synergistically interacting terminal system regulatory elements. Mech Dev 40, 47-61.

Liu, P. Z., and Kaufman, T. C. (2004a). hunchback is required for suppression of abdominal identity, and for proper germband growth and segmentation in the intermediate germband insect Oncopeltus fasciatus. Development 131, 1515-1527.

Liu, P. Z., and Kaufman, T. C. (2004b). Kruppel is a gap gene in the intermediate germband insect Oncopeltus fasciatus and is required for development of both blastoderm and germband-derived segments. Development 131, 4567-4579.

Liu, P. Z., and Kaufman, T. C. (2005). even-skipped is not a pair-rule gene but has segmental and gap-like functions in Oncopeltus fasciatus, an intermediate germband insect. Development 132, 2081-2092.

Lynch, J. A., Brent, A. E., Leaf, D. S., Pultz, M. A., and Desplan, C. (2006). Localized maternal orthodenticle patterns anterior and posterior in the long germ wasp Nasonia. Nature 439, 728-732.

Macdonald, P. M., and Struhl, G. (1986). A molecular gradient in early Drosophila embryos and its role in specifying the body pattern. Nature 324, 537-545.

Manzanares, M., Marco, R., and Garesse, R. (1993). Genomic organization and developmental pattern of expression of the engrailed gene from the brine shrimp Artemia. Development (Cambridge) 118, 1209-1219.

Manzanares, M., Williams, T. A., Marco, R., and Garesse, R. (1996). Segmentation in the crustacean Artemia: Engrailed staining studied with an antibody raised against the Artemia protein. Roux's Archives of Developmental Biology 205, 424-431.

111

Maroto, M., Dale, J. K., Dequeant, M. L., Petit, A. C., and Pourquie, O. (2005). Synchronised cycling gene oscillations in presomitic mesoderm cells require cell-cell contact. Int J Dev Biol 49, 309-315.

Marques-Souza, H., Aranda, M., and Tautz, D. (2008). Delimiting the conserved features of hunchback function for the trunk organization of insects. Development 135, 881-888.

Maruhashi, M., Van De Putte, T., Huylebroeck, D., Kondoh, H., and Higashi, Y. (2005). Involvement of SIP1 in positioning of somite boundaries in the mouse embryo. Dev Dyn 234, 332-338.

McGregor, A. P., Pechmann, M., Schwager, E. E., Feitosa, N. M., Kruck, S., Aranda, M., and Damen, W. G. (2008). Wnt8 is required for growth-zone establishment and development of opisthosomal segments in a spider. Curr Biol 18, 1619-1623.

McGrew, M. J., Dale, J. K., Fraboulet, S., and Pourquie, O. (1998). The lunatic fringe gene is a target of the molecular clock linked to somite segmentation in avian embryos. Curr Biol 8, 979-982.

Micchelli, C. A., Esler, W. P., Kimberly, W. T., Jack, C., Berezovska, O., Kornilova, A., Hyman, B. T., Perrimon, N., and Wolfe, M. S. (2003). Gamma-secretase/presenilin inhibitors for Alzheimer's disease phenocopy Notch mutations in Drosophila. Faseb J 17, 79-81.

Mito, T., Kobayashi, C., Sarashina, I., Zhang, H., Shinahara, W., Miyawaki, K., Shinmyo, Y., Ohuchi, H., and Noji, S. (2007). even-skipped has gap-like, pair-rule-like, and segmental functions in the cricket Gryllus bimaculatus, a basal, intermediate germ insect (Orthoptera). Dev Biol 303, 202-213.

Mito, T., Okamoto, H., Shinahara, W., Shinmyo, Y., Miyawaki, K., Ohuchi, H., and Noji, S. (2006). Kruppel acts as a gap gene regulating expression of hunchback and even-skipped in the intermediate germ cricket Gryllus bimaculatus. Dev Biol 294, 471-481.

Mito, T., Sarashina, I., Zhang, H., Iwahashi, A., Okamoto, H., Miyawaki, K., Shinmyo, Y., Ohuchi, H., and Noji, S. (2005). Non-canonical functions of hunchback in segment patterning of the intermediate germ cricket Gryllus bimaculatus. Development 132, 2069-2079.

Morales, A. V., Yasuda, Y., and Ish-Horowicz, D. (2002). Periodic Lunatic fringe expression is controlled during segmentation by a cyclic transcriptional enhancer responsive to notch signaling. Dev Cell 3, 63-74.

Nagy, L. M., and Carroll, S. (1994). Conservation of wingless patterning functions in the short-germ embryos of Tribolium castaneum. Nature 367, 460-463.

112

Nauber, U., Pankratz, M. J., Kienlin, A., Seifert, E., Klemm, U., and Jackle, H. (1988). Abdominal segmentation of the Drosophila embryo requires a hormone receptor-like protein encoded by the gap gene knirps. Nature 336, 489-492.

Negrea, S., Botnariuc, N. and Dumont, H.J. (1999). Phylogeny, evolution and classification of the Branchiopoda (Crustacea). Hydrobiology 412, 191-212.

Nielsen, C. (2001). Animal Evolution: Interrelationships of Living Phyla Oxford Univesity Press, Oxford, UK).

Niwa, Y., Masamizu, Y., Liu, T., Nakayama, R., Deng, C. X., and Kageyama, R. (2007). The initiation and propagation of Hes7 oscillation are cooperatively regulated by Fgf and notch signaling in the somite segmentation clock. Dev Cell 13, 298-304.

Nulsen, C., and Nagy, L. M. (1999). The role of wingless in the development of multibranched crustacean limbs. Dev Genes Evol 209, 340-348.

Nusslein-Volhard, C., Lohs-Schardin, M., Sander, K., and Cremer, C. (1980). A dorso-ventral shift of embryonic primordia in a new maternal-effect mutant of Drosophila. Nature 283, 474-476.

Nusslein-Volhard, C., and Wieschaus, E. (1980). Mutations affecting segment number and polarity in Drosophila. Nature 287, 795-801.

O'Day, K. E. (2006) Notch signaling and segmentation in Paryhale hawainesis, University of California, Berkeley, San Francisco, CA.

Oates, A. C., and Ho, R. K. (2002). Hairy/E(spl)-related (Her) genes are central components of the segmentation oscillator and display redundancy with the Delta/Notch signaling pathway in the formation of anterior segmental boundaries in the zebrafish. Development 129, 2929-2946.

Oates, A. C., Mueller, C., and Ho, R. K. (2005). Cooperative function of deltaC and her7 in anterior segment formation. Dev Biol 280, 133-149.

Oda, H., Nishimura, O., Hirao, Y., Tarui, H., Agata, K., and Akiyama-Oda, Y. (2007). Progressive activation of Delta-Notch signaling from around the blastopore is required to set up a functional caudal lobe in the spider Achaearanea tepidariorum. Development 134, 2195-2205.

Osborne, P. W., and Dearden, P. K. (2005). Expression of Pax group III genes in the honeybee (Apis mellifera). Dev Genes Evol 215, 499-508.

Ozbudak, E. M., and Lewis, J. (2008). Notch signalling synchronizes the zebrafish segmentation clock but is not needed to create somite boundaries. PLoS Genet 4, e15.

113

Ozbudak, E. M., and Pourquie, O. (2008). The vertebrate segmentation clock: the tip of the iceberg. Curr Opin Genet Dev 18, 317-323.

Palmeirim, I., Henrique, D., Ish-Horowicz, D., and Pourquie, O. (1997). Avian hairy gene expression identifies a molecular clock linked to vertebrate segmentation and somitogenesis. Cell 91, 639-648.

Panganiban, G., Sebring, A., Nagy, L., and Carroll, S. (1995). The Development of Crustacean Limbs and the Evolution of Arthropods. Science 270, 1363-1366.

Pankratz, M. J., Jackle, J. (1993). Blastoderm segmentation., Vol 1 (New York, New York: Cold Spring Harbor Lab Press).

Patel, N. H., Ball, E. E., and Goodman, C. S. (1992). Changing role of even-skipped during the evolution of insect pattern formation. Nature 357, 339-342.

Patel, N. H., Condron, B. G., and Zinn, K. (1994). Pair-rule expression patterns of even-skipped are found in both short- and long-germ beetles. Nature 367, 429-434.

Patel, N. H., Hayward, D. C., Lall, S., Pirkl, N. R., DiPietro, D., and Ball, E. E. (2001). Grasshopper hunchback expression reveals conserved and novel aspects of axis formation and segmentation. Development 128, 3459-3472.

Patel, N. H., Martin-Blanco, E., Coleman, K. G., Poole, S. J., Ellis, M. C., Kornberg, T. B., and Goodman, C. S. (1989). Expression of engrailed proteins in arthropods, annelids, and chordates. Cell 58, 955-968.

Peel, A. D., Chipman, A. D., and Akam, M. (2005). Arthropod segmentation: beyond the Drosophila paradigm. Nat Rev Genet 6, 905-916.

Peres, J. N., McNulty, C. L., and Durston, A. J. (2006). Interaction between X-Delta-2 and Hox genes regulates segmentation and patterning of the anteroposterior axis. Mech Dev 123, 321-333.

Peterson, M. D., Popadic, A., and Kaufman, T. C. (1998). The expression of two engrailed-related genes in an apterygote insect and a phylogenetic analysis of insect engrailed-related genes. Dev Genes Evol 208, 547-557.

Pourquie, O. (2001). Vertebrate somitogenesis. Annu Rev Cell Dev Biol 17, 311-350.

Pourquie, O. (2003). Vertebrate somitogenesis: a novel paradigm for animal segmentation? Int J Dev Biol 47, 597-603.

Prince, V. E., Holley, S. A., Bally-Cuif, L., Prabhakaran, B., Oates, A. C., Ho, R. K., and Vogt, T. F. (2001). Zebrafish lunatic fringe demarcates segmental boundaries. Mech Dev 105, 175-180.

114

Pueyo, J. I., Lanfear, R., and Couso, J. P. (2008). Ancestral Notch-mediated segmentation revealed in the cockroach Periplaneta americana. Proc Natl Acad Sci U S A 105, 16614-16619.

Quiring, R., Walldorf, U., Kloter, U., and Gehring, W. J. (1994). Homology of the eyeless gene of Drosophila to the Small eye gene in mice and Aniridia in humans. Science 265, 785-789.

Rabet, N., Gibert, J. M., Queinnec, E., Deutsch, J. S., and Mouchel-Vielh, E. (2001). The caudal gene of the barnacle Sacculina carcini is not expressed in its vestigial abdomen. Dev Genes Evol 211, 172-178.

Richardson, M. K., Allen, S. P., Wright, G. M., Raynaud, A., and Hanken, J. (1998). Somite number and vertebrate evolution. Development 125, 151-160.

Rida, P. C., Le Minh, N., and Jiang, Y. J. (2004). A Notch feeling of somite segmentation and beyond. Dev Biol 265, 2-22.

Riedel-Kruse, I. H., Muller, C., and Oates, A. C. (2007). Synchrony dynamics during initiation, failure, and rescue of the segmentation clock. Science 317, 1911-1915.

Rivera, A. S., Gonsalves, F. C., Song, M. H., Norris, B. J., and Weisblat, D. A. (2005). Characterization of Notch-class gene expression in segmentation stem cells and segment founder cells in Helobdella robusta (Lophotrochozoa; Annelida; Clitellata; Hirudinida; Glossiphoniidae). Evol Dev 7, 588-599.

Rivera, A. S., and Weisblat, D. A. (2008). And Lophotrochozoa makes three: Notch/Hes signaling in annelid segmentation. Dev Genes Evol.

Rose, T. M., Henikoff, J. G., and Henikoff, S. (2003). CODEHOP (COnsensus-DEgenerate Hybrid Oligonucleotide Primer) PCR primer design. Nucleic Acids Res 31, 3763-3766.

Saga, Y. (2007). Segmental border is defined by the key transcription factor Mesp2, by means of the suppression of Notch activity. Dev Dyn 236, 1450-1455.

Salazar-Ciudad, I., Sole, R. V., and Newman, S. A. (2001). Phenotypic and dynamical transitions in model genetic networks. II. Application to the evolution of segmentation mechanisms. Evol Dev 3, 95-103.

Satoh, W., Gotoh, T., Tsunematsu, Y., Aizawa, S., and Shimono, A. (2006). Sfrp1 and Sfrp2 regulate anteroposterior axis elongation and somite segmentation during mouse embryogenesis. Development 133, 989-999.

115

Sawada, A., Shinya, M., Jiang, Y. J., Kawakami, A., Kuroiwa, A., and Takeda, H. (2001). Fgf/MAPK signalling is a crucial positional cue in somite boundary formation. Development 128, 4873-4880.

Schinko, J. B., Kreuzer, N., Offen, N., Posnien, N., Wimmer, E. A., and Bucher, G. (2008). Divergent functions of orthodenticle, empty spiracles and buttonhead in early head patterning of the beetle Tribolium castaneum (Coleoptera). Dev Biol 317, 600-613.

Scholtz, G. (2002). The Articulata hypothesis – or what is a segment? Organisms Diversity & Evol 2, 197-215.

Scholtz, G., Patel, N. H., and Dohle, W. (1994). Serially homologous engrailed stripes are generated via different cell lineages in the germ band of amphipod crustaceans (Malacostraca, Peracarida). Int J Dev Biol 38, 471-478.

Schoppmeier, M., and Damen, W. G. (2005a). Expression of Pax group III genes suggests a single-segmental periodicity for opisthosomal segment patterning in the spider Cupiennius salei. Evol Dev 7, 160-169.

Schoppmeier, M., and Damen, W. G. (2005b). Suppressor of Hairless and Presenilin phenotypes imply involvement of canonical Notch-signalling in segmentation of the spider Cupiennius salei. Dev Biol 280, 211-224.

Schroder, R. (2003). The genes orthodenticle and hunchback substitute for bicoid in the beetle Tribolium. Nature 422, 621-625.

Schulz, C., Schroder, R., Hausdorf, B., Wolff, C., and Tautz, D. (1998). A caudal homologue in the short germ band beetle Tribolium shows similarities to both, the Drosophila and the vertebrate caudal expression patterns. Dev Genes Evol 208, 283-289.

Seaver, E. C. (2003). Segmentation: mono- or polyphyletic? Int J Dev Biol 47, 583-595.

Shifley, E. T., Vanhorn, K. M., Perez-Balaguer, A., Franklin, J. D., Weinstein, M., and Cole, S. E. (2008). Oscillatory lunatic fringe activity is crucial for segmentation of the anterior but not posterior skeleton. Development 135, 899-908.

Shimizu, T., and Savage, R. M. (2002). Expression of hunchback protein in a subset of ectodermal teloblasts of the oligochaete annelid Tubifex. Dev Genes Evol 212, 520-525.

Shinmyo, Y., Mito, T., Matsushita, T., Sarashina, I., Miyawaki, K., Ohuchi, H., and Noji, S. (2005). caudal is required for gnathal and thoracic patterning and for posterior elongation in the intermediate-germband cricket Gryllus bimaculatus. Mech Dev 122, 231-239.

116

Simonnet, F., Deutsch, J., and Queinnec, E. (2004). hedgehog is a segment polarity gene in a crustacean and a chelicerate. Dev Genes Evol 214, 537-545.

Sommer, R., and Tautz, D. (1991). Segmentation gene expression in the housefly Musca domestica. Development 113, 419-430.

Sommer, R. J., and Tautz, D. (1993). Involvement of an orthologue of the Drosophila pair-rule gene hairy in segment formation of the short germ-band embryo of Tribolium (Coleoptera). Nature 361, 448-450.

Song, M. H., Huang, F. Z., Gonsalves, F. C., and Weisblat, D. A. (2004). Cell cycle-dependent expression of a hairy and Enhancer of split (hes) homolog during cleavage and segmentation in leech embryos. Dev Biol 269, 183-195.

Stauber, M., Jackle, H., and Schmidt-Ott, U. (1999). The anterior determinant bicoid of Drosophila is a derived Hox class 3 gene. Proc Natl Acad Sci U S A 96, 3786-3789.

Stauber, M., Prell, A., and Schmidt-Ott, U. (2002). A single Hox3 gene with composite bicoid and zerknullt expression characteristics in non-Cyclorrhaphan flies. Proc Natl Acad Sci U S A 99, 274-279.

Stauber, M., Taubert, H., and Schmidt-Ott, U. (2000). Function of bicoid and hunchback homologs in the basal cyclorrhaphan fly Megaselia (Phoridae). Proc Natl Acad Sci U S A 97, 10844-10849.

Stern, C. D. (1990). Two distinct mechanisms for segmentation? Semin Dev Biol 1, 109-116.

Stollewerk, A. (2002). Recruitment of cell groups through Delta/Notch signalling during spider neurogenesis. Development 129, 5339-5348.

Stollewerk, A., Schoppmeier, M., and Damen, W. G. (2003). Involvement of Notch and Delta genes in spider segmentation. Nature 423, 863-865.

Takahashi, Y., Koizumi, K., Takagi, A., Kitajima, S., Inoue, T., Koseki, H., and Saga, Y. (2000). Mesp2 initiates somite segmentation through the Notch signalling pathway. Nat Genet 25, 390-396.

Takke, C., and Campos-Ortega, J. A. (1999). her1, a zebrafish pair-rule like gene, acts downstream of notch signalling to control somite development. Development 126, 3005-3014.

Tautz, D. (1994). Evolutionary analysis of genes involved in early embryonic pattern formation in Drosophila. Exs 69, 525-536.

Tautz, D. (2004). Segmentation. Dev Cell 7, 301-312.

117

Valentine, J. W. (1994). Late Precambrian bilaterians: grades and clades. Proc Natl Acad Sci U S A 91, 6751-6757.

Valentine, J. W., and Collins, A. G. (2000). The significance of moulting in Ecdysozoan evolution. Evol Dev 2, 152-156.

Vanhaecke, P., Siddall, S.E., Sorgeloos, P. (1984). International study on Artemia. XXXII. Combined effects of temperature and salinity on the survival of Artemia of various geographical origin. Journal of Experimental Marine Biology and Ecology 80, 259-275.

Wang, C., and Lehmann, R. (1991). Nanos is the localized posterior determinant in Drosophila. Cell 66, 637-647.

Wear, R. G., Haslett, S. J. (1986). Effects of temperature and salinity on the biology of Artemia franciscana Kellogg from lake Grassmere, New Zealand. 1. Growth and mortality. . Journal of Experimental Marine Biology and Ecology 98, 153-166.

Wieschaus, E., Nusslein-Volhard, C., and Kluding, H. (1984). Kruppel, a gene whose activity is required early in the zygotic genome for normal embryonic segmentation. Dev Biol 104, 172-186.

Wilmer, P. (1990). Body Divisions - Metamerism and Segmentation, In, I. R. P. i. A. Evolution, ed. (Cambridge University Press, Cambridge, UK), pp. 39-45.

Wolff, C., Sommer, R., Schroder, R., Glaser, G., and Tautz, D. (1995). Conserved and divergent expression aspects of the Drosophila segmentation gene hunchback in the short germ band embryo of the flour beetle Tribolium. Development 121, 4227-4236.