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2017 Studies on development in Euphilomedes ostracods: Embryology, nervous system development, and the genetics of sexually dimorphic eye development Kristina Koyama University of the Pacific, [email protected]
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STUDIES ON DEVELOPMENT IN EUPHILOMEDES OSTRACODS: EMBRYOLOGY, NERVOUS SYSTEM DEVELOPMENT, AND THE GENETICS OF SEXUALLY DIMORPHIC EYE DEVELOPMENT
by
Kristina H. Koyama
A Thesis Submitted to the
Office of Research and Graduate Studies
In Partial Fulfillment of the Requirements for the Degree of
MASTER OF SCIENCE
College of the Pacific Biological Sciences
University of the Pacific Stockton, California
2017 2
STUDIES ON DEVELOPMENT IN EUPHILOMEDES OSTRACODS: EMBRYOLOGY, NERVOUS SYSTEM DEVELOPMENT, AND THE GENETICS OF SEXUALLY DIMORPHIC EYE DEVELOPMENT
by
Kristina H. Koyama
Approved by
Thesis Advisor: Ajna Rivera, Ph.D.
Committee Member: Douglas Weiser, Ph.D.
Committee Member: Tara Thiemann, Ph.D.
Department Chair: Craig Vierra, Ph.D.
Dean of Graduate School: Thomas H. Naehr, Ph.D.
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DEDICATION
For everyone who gave me their support, encouragement, and patience during this crazy journey. It wouldn’t have been possible without you (no contribution too small)!
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ACKNOWLEDGEMENTS
I would like to thank Dr. Ajna Rivera for her mentorship and support during the writing of this thesis, and for giving the opportunity to explore and learn. I would also like to thank my other two committee members, Dr. Thiemann and Dr. Weiser, for their support and guidance. I also would like to thank my lab mates, past and present, for your lab help, comradery, and friendship. It’s been a pleasure working with you all. Finally, I would like to thank the staff, faculty, and graduate students of the Biology department, for your support. You are all wonderful people to work with, and truly inspiring. Thank you all for believing in me.
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Studies on development in Euphilomedes ostracods: Embryology, nervous system development, and the genetics of sexually dimorphic eye development
Abstract
By Kristina H. Koyama
University of the Pacific 2015
Model organism studies have been fundamental in understanding evolutionary and developmental biology. However, non-model organisms present opportunities to study unique characteristics and as comparisons to model organisms, leading us toward broader and more relevant perspectives on diversity. The Euphilomedes genus of ostracods is an example of a non-model group with potential for evolutionary and developmental studies.
Ostracoda is an ancient, basally branching lineage of Crustaceans with a complete and prodigious fossil record. Despite the group’s promise for evolutionary studies, much remains unknown about the basic biology of this clade. There are a limited number of embryogenesis studies in Ostracoda; here, I study development in Euphilomedes.
In Part 1, I study the main events in Euphilomedes’ embryology, focusing on cleavage and cell migration. I describe the general embryology of Euphilomedes, and devise a visual staging scheme for their development. Using fluorescent nuclear staining and microscopy, I visualize nuclei in cleavage throughout development of nuclear 6 divisions and migrations during development. The meroblastic cleavage observed in
Euphilomedes resembles that of another Myodocopid ostracod, Vargula hilgendorfii.
Finally, immunostaining for acetylated-alpha tubulin and phalloidin staining are used to visualize the general anatomy of the embryonic brain. This provides new protocols for visualizing the nervous system, enabling more detailed nervous system studies in the future.
In Part 2, I explore differential gene expression patterns in the developing eyes of juvenile Euphilomedes. Euphilomedes have sexually dimorphic eye types – males have lateral compound eyes, while females instead have eye rudiments. Previous studies in E. carcharodonta show that genes in the retinal determination and phototransduction gene networks have differential expression in males and females during eye development. In this thesis, we attempt to compare these patterns to expression in a sister species, E. morini.
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TABLE OF CONTENTS
LIST OF TABLES ...... 10
LIST OF FIGURES ...... 11
LIST OF ABBREVIATIONS ...... 12
CHAPTER
1. General introduction ...... 14
2. Introduction to Part 1: Embryogenesis and Embryonic Nervous System in Euphilomedes carcharodonta ...... 18
Euphilomedes ostracods as representatives of Myodocopid ostracods ....18
Embryogenesis in Myodocopid ostracods ...... 20
Euphilomedes as a model for the superfamilly Sarsielloidea ...... 21
Euphilomedes ostracods as representatives in Crustacea ...... 21
Neurophylogeny in Crustacea and Arthropoda...... 23
The nervous system in Ostracoda ...... 24
Practical techniques for embryology studies in Euphilomedes ...... 25
3. Methods...... 27
Collection of E. carcharodonta ...... 27
Immunohistochemistry, phalloidin, and DAPI staining ...... 27
Fluorescent microscopy and microphotography ...... 29
Color staging and photography ...... 30
Estimation of brooding duration ...... 30
Improvement of artificial rearing ...... 31
4. Results ...... 32
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General embryology...... 32
Early nuclear divisions and cell migrations ...... 34
Limb bud formation and late development ...... 37
Improvement of artificial rearing technique ...... 38
Nervous system visualization ...... 39
Brain structure in Euphilomedes embryos ...... 41
5. Discussion ...... 43
6. Introduction to Part 2: Differential gene expression patterns in the sexually dimorphic eyes of Euphilomedes ostracods ...... 46
Why study sexual dimorphism? ...... 47
Euphilomedes ostracods as models for arthropod eye evolution ...... 49
Ommatidial development ...... 52
Phototransduction ...... 54
Morphological differences from similar genetic backgrounds ...... 55
Can the same morphology be determined by different gene expression patterns? ...... 56
Studying differential gene expression in Euphilomedes ...... 57
7. Methods...... 60
Collection of Euphilomedes morini ...... 60
Dissection ...... 60
RNA isolation and cDNA synthesis ...... 61
Generation of qPCR standard curves ...... 61
8. Results ...... 64
9. Discussion ...... 68
9
REFERENCES ...... 69
10
LIST OF TABLES
Table Page
1. Expression data for TOY ...... 64
2. Expression data for ELAV ...... 65
3. Expression data for PLC ...... 66
4. Expression data for DAC ...... 66
5. Expression data for SIA ...... 66
6. Expression data for SO ...... 67
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LIST OF FIGURES
Figure Page
1. Cleavage patterns in Ostracoda ...... 19
2. Cleavage patterns in Crustacea ...... 22
3. Preparation of embryos and agarose-pad mounting ...... 28
4. Normal and malformed embryo morphology ...... 30
5. Embryos of the ostracod E. carcharodonta ...... 32
6. Early embryogenesis visualized with nuclear staining ...... 34
7. Asynchronous and synchronous mitoses ...... 35
8. Presumptive fates of limb buds in early embryo ...... 36
9. Carapace development in late embryos ...... 37
10. Average percent survival for artificially reared embryos ...... 39
11. Embryonic brain labeled with ac-α tubulin immunostaining and phalloidin.40
12. Central nervous system in Euphilomedes ...... 41
13. Comparative cleavage and migration patterns in Parhyale, Vargula, and Euphilomedes ...... 43
14. Sexually dimorphic eyes of the Euphilomedes ostracods ...... 50
15. Comparison of compound eye development in arthropods ...... 52
16. Signaling events in the retinal determination gene network (RDGN) ...... 53
17. Phototransduction cascade in Drosophila ...... 54
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LIST OF ABBREVIATIONS
ac Acetylated au Nauplius eye ca Carapace cDNA Complementary deoxyribonucleic acid Chp Chaoptic Dac Dachshund DAG Diacyl glycerol DAPI 4',6-diamidino-2-phenylindole DC Deutocerebrum DNA Deoxyribonucleic acid DNAse Deoxyribonuclease Dpp Decapentaplegic EGFR Epidermal growth factor receptor Ey Eyeless Eya Eyes absent fa first antenna fl fifth limb fu furcal claw GDP Guanosine diphosphate gnt Tegumentary nerve ganglion Gq guanine nucleotide binding protein q polypeptide GRN Gene regulatory network GTP Guanosine-5'-triphosphate Hh Hedgehog IP3 inositol triphosphate LPC latereal protocerebrum MN mandible MPC medial protocerebrum mRNA messenger ribonucleic acid MX maxilla NA1 Nerve of the first antennae Pax Paired box protein pb polar body PBS Phosphate-buffered saline PC Protocerebrum PCR Polymerase chain reaction PFA Paraformaldehyde PIP2 phosphatidylinositol biphosphate 13
PUFA Polyunsaturated fatty acids qPCR Real-time quantitative polymerase chain reaction RDGN Retinal determination gene network Rh1 Rhodopsin Rh-phalloidin Rhodamine-conjugated phalloidin RNA Ribonucleic acid sa second antenna So Sine oculis Svn Shaven TC tritocerebrum Toy Twin of eyeless TRP Transient receptor potential channel TRPL TRP-like
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Chapter 1: General introduction
From where does diversity originate? This is one of the fundamental questions in biology. Two fields which stem from this question are evolutionary and developmental biology. Much of the knowledge amassed around developmental processes and genetics has resulted from studies in a relatively small number of model organisms. The popularity and laboratory practicality of these animals has made them not only relatively easy to study, but has resulted in reliable laboratory procedures to do so. Due to ever- increasing standardization and optimization, there is a consistent stream of studies performed in these animals with an otherwise unattainable level of detail. However, studying a limited number of representatives of the animal kingdom inevitably leaves room for error in assumptions and false conclusions (1). Model organisms have been, in many ways, selected for uncommon features which make them ideal for efficient and consistent studies (2). Most model organisms (C. elegans, mice, zebrafish, frogs, chickens) were initially chosen for study because of their high level of domestication: fast development, short lifespans, a lab-sustainable diet, and consistent phenotypes. This has enabled a great deal of standardization and detail in studies, and laid much of the groundwork for developmental biology. Studies in these animals led to the discovery of consistent developmental schedules orchestrated directly by tightly regulated genetic programs. However, the intense focus on consistent phenotypes and direct genetic effects has also largely ignored other large questions in development. For example, developmental plasticity and other external effects on development are often ignored in 15 genetic studies on development, but their importance is undeniably relevant to real-world environments, and finally is gaining recognition in the context of climate change (3).
Intraspecific variation is largely ignored in model organisms, and is often deliberately removed, for example, through the establishment of reliable “wild type” lineages (4).
However, it is now evident that understanding this variation is imperative to our understanding of real-world variations (4). The value of model organism studies should not be diminished or ignored, but these studies will not suffice on their own as accurate depictions of development or evolution. Therefore, it is crucial to study a broader diversity of animals to establish an equally broad view of developmental processes and the genetics that drive them.
Accordingly, modern evolutionary biologists are coming to incorporate a wider range of animals in their analyses. Next-generation sequencing techniques and modern computing power make it possible and affordable to include an increasing number of organisms into genomic, transcriptomic, and proteomic studies (5). This allows for studies which look at diversity in a broader manner, with incorporation of many more representatives into studies. This technology is necessary for advancement of evolutionary knowledge, but is even more so powerful when combined with other methodologies, such as morphological studies. Amid a technological wave, morphological studies remain imperative to understanding evolution. The use of morphology and character traits help to alleviate biases of purely molecular studies (6).
Furthermore, morphological studies also give context to genetic data, and demonstrate functional changes over time. 16
One relatively diverse, yet understudied animal group is Subphylum Crustacea.
Arthropods in general are one of the most diverse and successful groups on the planet, and the Crustaceans themselves exhibit a wide amount of morphological diversity amongst this taxon. Even within Crustacea, most studies are limited to the order
Malacostraca (e.g. amphipods and decapods). While Malacostracans are diverse themselves, studies on Malacostraca alone again can lead to false conclusions about the origins and general biology of crustaceans on a broader scale.
One group which is understudied, yet promising, is the crustacean order
Ostracoda. Ostracods are an ancient and diverse group of crustaceans. As one of the earliest groups to branch off of the crustacean tree, ostracods are in a unique evolutionary position for study (7). In recent bioinformatic studies, once obscure and understudied groups such as Ostracoda are now being included and help to reflect a broader and more inclusive approach to phylogenetics (7). Furthermore, Ostracoda also has a uniquely rich and complete fossil record which can be used in conjunction with molecular and morphological studies to answer evolutionary questions. Applications of ostracod studies are not limited to evolutionary and developmental questions. Basic biology in Ostracoda is a mostly unexplored, but promising topic; ostracods are frequently used as bioindicators. A more nuanced understanding of their biology could make them even more powerful as tools to monitor changing aquatic environments.
This thesis study aims to take a closer look at several aspects of development in
Euphilomedes, a genus of marine ostracods. The first topic is embryonic development.
Embryology is a relatively obscure topic within Ostracoda. Furthermore, most studies done prior to the 21st century were done in Podocopid ostracods, one of the two extant 17 lineages of ostracods. More recent evidence from a study on a Myodocopid ostracod, V. hilgendorfii, suggests that these two ostracod lineages may have fundamental differences in their development (8). Part 1 of this thesis will discuss embryonic development in detail, providing a much-needed representation of ostracod development in Myodocopa.
Perhaps even more rare is studies on the anatomy of embryonic nervous systems in Ostracoda. Only a single study in the last century has described the anatomy of the embryonic nervous system, and again, this study was done in a Podocopid ostracod. This study will visualize the embryonic nervous system architecture in Part 1, and will be the first representation of the nervous system in a Myodocopid ostracod.
Beyond embryonic development, Euphilomedes ostracods have other unique features which make them interesting models for studying the evolution of complex organs. Ostracods of this genus exhibit radical sexual dimorphism in their eye types.
Males have compound lateral eyes, while females have only simple eyespots. Combined with XX/XO sex determination, this makes them idea for studying the genetic mechanisms behind complex organ development and evolution (9). Their sexually dimorphic eye development will be the main topic of Part 2 of this thesis.
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Introduction to Part 1: Embryogenesis and Embryonic Nervous System in
Euphilomedes carcharodonta
Ostracoda is an incredibly large crustacean group, including some 65,000 extant and extinct described species (10). Though the group is incredibly large, it is relatively understudied, and much remains unknown about the biology of these animals.
Euphilomedes ostracods as representatives of Myodocopid ostracods.
Euphilomedes is a genus of benthic marine ostracods belonging to the Subclass
Myodocopa, and Superfamily Sarsielloidea. Sarsielloidea is a superfamily of great interest due to its unique features, including bioluminescence and sexually dimorphic compound lateral eyes (9,11–13). The Euphilomedes genus itself has sexually dimorphic eyes, and this feature has been the topic of several studies (9,14–16). This thesis study will focus on the species Euphilomedes carcharodonta, a benthic species found in the shallow subtital off the coast of California. Its adult and several juvenile stages have been previously described (9,17). Myodocopid ostracods, including Euphilomedes, undergo direct development – their juvenile instars are free-swimming and have a body plan which generally resembles that of the adult. Euphilomedes females brood their eggs as described in other Myodocopid ostracods, protected within the carapace in a dorsal area called the marsupium (18). Embryos complete embryogenesis and are released from the marsupium when they are ready to hatch as first juvenile instars. Juveniles have 5 instars before their adult stage (19). Euphilomedes’ eggs are surrounded by a tough, 19
transparent chorion, and while the composition of the membrane has not yet been
investigated, other crustaceans with lengthy embryonic development have chitin-based
chorions (20). More evidence of a chitinous composition comes from laboratory
observations. The chorions of Euphilomedes are extremely tough, technically impossible
to remove manually from living embryos, and are not naturally permeable to anything but
small molecules, bearing resemblance to the chitinous chorions of other crustaceans (20).
Treatment of fixed embryos with chitinase degrades the embryo, including the chorion
(A. Sajuthi, personal communication, 2015).
The Euphilomedes genus is a promising system for studying the development and
phylogenetic relationships of ostracods. Specifically, Euphilomedes could be an ideal
developmental representative for Myodocopida. This study will focus on the embryology
of E. carcharodonta, with emphases on early nuclear divisions, general morphological
changes, and the structure of the nervous system. Juvenile development is has been
described in several Myodocopid species, and is the topic of most developmental studies
in Ostracoda (e.g. 9–15). However, embryological studies are severely lacking in
Figure 1. Cleavage patterns in Ostracoda. Summary of cleavage patterns from studied ostracod species. Cleavage patterns differ between Podocopa and Myodocopa. Podocopa has been described to have holoblastic and equal cleavages, while cleavages in Myodocopa do not show evidence of a visible cleavage plane or furrow. Cyprideis littoralis, Weygoldt 1960. Heterocypris incongruens, Müller-Calé 1913. Vargula hilgendorfii, Wakayama et al. 2007. 20
Ostracoda. Only a handful of ostracod species have been studied in any detailed embryological manner, and these few representatives are vastly limited in their ability to represent the diversity of a large and ancient group such as Ostracoda (8,28,29).
Embryogenesis in Myodocopid ostracods. Vargula hilgendorfii is the only
Myodocopid ostracod whose embryonic development has been previously studied in detail (30). V. hilgendorfii’s early development has been investigated using nuclear staining and scanning electron microscopy (8). In V. hilgendorfii, nuclei are initially embedded in the yolk and undergo early divisions without evidence of a cleavage furrow
(8). The nuclei migrate and form a cell aggregation on one side of the embryo, in a stage the authors call ‘pseudo-cleavage,’ due to its resemblance to two blastomeres (8).
However, this is not a true cleavage: instead, the two spheroid masses actually consist of a yolk sphere and a cell aggregation (8). The cell aggregation grows and spreads to cover the yolk and form a blastula (8). Following the formation of the blastula, limb buds form, followed by the development of the carapace (8). Finally, organogenesis occurs and the ostracod hatches from its chorion as a juvenile first instar.
Carapace development in Ostracoda is of interest – specifically, the developmental and evolutionary origins of the carapace hinge are a topic of debate (8). It has been hypothesized that the bivalved carapace in Ostracoda is derived from a one piece carapace – during development, this one piece carapace would split and form a hinge as growth proceeded (8). Evidence for this hypothesis comes from fossil species which have a one-piece carapace, such as Manawa (8). However, in Vargula, the carapace develops from two distinct buds, which join to form a one-piece carapace, then 21 once again split to form a hinge (8). This suggests that the one-piece carapace found in
Vargula may have different origins than from that found in fossils like Manawa (8).
As the lone Myodocopid representative, V. hilgendorfii has no comparative companion in this group – other embryological studies in ostracods were performed on ostracods from the other extant ostracod group, Podocopa (28,29). Comparison between
V. hilgendorfii and two Podocopid representatives provides evidence that embryogenesis likely differs between the two groups, a reasonable suggestion as the two groups have an ancient divergence (Figure 1) (8). However, reliable comparisons between the twogroups’ embryology require studies on more species from each group to draw solid conclusions about their development and their relationships.
Euphilomedes as a model for the superfamily Sarsielloidea. Euphilomedes also can be used as a model for ostracods of the superfamily Sarsielloidea. Sarsielloidea unlike other ostracod groups, does not have fossil representatives, underscoring the need for methods outside of paleontology in investigating its phylogenetic relationships (11).
Sarsiellids are also of great interest in regard to their basic biology – this group contains species investigated for behavioral, biochemical, and genetic studies (e.g. 6–8,24–28).
Euphilomedes ostracods as representatives in Crustacea. Cleavage patterns in
Crustacea vary widely within and between groups, making it difficult to determine a pleisomorphic cleavage pattern for Crustacea. Though some evidence points toward total cleavage as the ancestral state for Crustacea, many groups lack information on cleavage patterns, leading one to question this conclusion for the clade (Figure 2) (36). Some clades have been studied, such as Ostracoda, but only have a few representatives whose cleavage has been characterized (36). In the case of Ostracoda, this lack of representation 22
has led to potentially false conclusions regarding its ancestral cleavage pattern: the
Podocopid cleavage pattern had previously been assumed to be representative of the
general pattern for the entire group (36). Yet, with more recent studies on V. hilgendorfii,
it is increasingly evident that this may not be the case, and the ancestral cleavage pattern
for Ostracoda must be reconsidered (29). Elucidating the ancestral ostracod cleavage
pattern could have implications for Oligostraca, the group containing Ostracoda,
Mystacocarida, Branchiura, and Pentastomida. Previously, it was thought that
Oligostraca likely had an ancestor with total cleavage, but with superficial cleavage
patterns found in Vargula and recently in a Branchiuran, the ancestral state looks less
obvious (36,37). Thus, more studies on these groups must be done to draw reliable
conclusions about the evolution of their cleavage patterns. Consequently, a definitive
ancestral state of crustacean cleavage patterns remains elusive. Though ostracod
Figure 2. Cleavage patterns in Crustacea. Cleavage patterns vary amongst Crustacean taxa. Many lineages, (e.g. Malaostraca, Ostracoda) have several cleavage patterns represented within the taxon. Several groups are missing information regarding their cleavage patterns. Crustacean tree and taxon names based on Oakley et. al 2013. 23 cleavage may not change conclusions about the ancestral crustacean cleavage pattern, the presence of more data in understudied clades may help to strengthen the hypotheses already proposed.
Crustacean cleavage patterns are partly of interest to uncover relationships between arthropod groups and arthropod sister group, as well as the origin of the arthropods themselves. Within Arthropoda, there is strong support for the Mandibulata hypothesis, which places Pancrustacea (also called Tetraconata) as sister group to
Myriapoda (38). With this placement, it is likely total cleavage which is the plesiomorphic state of Arthropoda (36). Arthropod relationships remain a topic of investigation, as much remains to be known about the biology of this large and diverse group. Cleavage patterns are one tool to bring insight to these relationships, but are not the only such feature crucial to phylogenetic studies. Other features, such as the structure of the nervous system, can also be used in a similar fashion, and will be discussed in the following section.
Neurophylogeny in Crustacea and Arthropoda. Although modern phylogenetic research relies heavily on molecular data, the value of morphological and developmental studies is not lost. Molecular studies cannot always resolve relationships, and in these cases morphological studies are imperative. One such feature which is heavily studied in Arthropoda and its relatives is the nervous system. The nervous system has been an important feature of study in phylogenetics, so much so that this field has its own term, neurophylogeny (39). The structure and development of the nervous system has long been used to find similarities between groups and resolve evolutionary relationships. 24
In general, crustacean embryos have a shared architecture, despite divergence of adult nervous anatomy. The crustacean nervous system is segmented, and is comprised of a circumoral ring, also referred to as a brain, and a ventral nerve cord (40,41). The ventral nerve cord is made of segments called neuromeres – these embryonic segments will condense during development into ganglia (40,41).
The nervous system in Ostracoda. The adult ostracod nervous system has been described in Podocopids, including Cypridopsis, Cyclocypris, Candona, Doloria, amongst others (42). These studies were done using classical histological techniques – studies on ostracod nervous systems have not been performed using modern techniques, such as immunohistochemical staining, which have since come into common practice in arthropod research. Aside this small number of studies in the 19th and 20th centuries, investigation of ostracod nervous system structure have been largely ignored (29,43).
Embryonic nervous system studies are even more rare than adult studies. In the last century, the lone study on the developing ostracod nervous system was performed by
Weygoldt (1960) in the Podocopid, Cyprideis litoralis (29). Weygoldt’s study follows the development of the central nervous system, with an emphasis on the development of the nerve ganglia (20).
Though referenced in text frequently, the study is difficult to use as a general reference for the ostracod nervous system. The exclusion of Weygoldt’s study here stems from two primary sources: First, the study has never been translated into English.
Second, though referenced often, the study is not included in any substantial review.
Though this may simply be a symptom of general neglect for ostracod biology, regardless, it is difficult to interpret Weygoldt’s study in the context of modern nervous 25 system studies. A lack of modern ostracod nervous system studies serves to exacerbate this predicament.
Amidst the general neglect of ostracod neuroanatomy, the nervous system of
Myodocopid ostracods is an untouched topic in biology. As fundamental developmental patterns differ between the two ostracod orders – i.e. cleavage – it is plausible that other developmental patterns, such as neurogenesis – may have diverged as well. Furthermore, the anatomy of first instar Podocopids and Myodocopids differ in their number of appendages, suggesting that the overall structure of the nervous system must differ to accommodate the two body plans. Myodocopids also have species with compound eyes, while Podocopids do not, and this would suggest differences in the structure of brain anatomy and development (44). As seen before, assumptions about a group as ancient and diverse as Ostracoda cannot accurately be made with only a few representatives from a small group within the clade. Such has been a predicament of evolutionary biology since its birth.
Consequently, development of modern histochemical techniques in ostracods is imperative. This study represents the first use of immunohistochemical staining in
Ostracoda, and is the first step at taking the field of ostracod anatomy and embryology into an era of modern techniques.
Practical techniques for embryology studies in Euphilomedes. The ability to perform detailed studies on Euphilomedes requires basic knowledge about these ostracods’ life history and basic embryogenesis. Here, we estimate the total duration of brooding. This study will explore a more effective artificial rearing technique for embryos. We estimate the number of embryos per brood, and explore embryo mortality 26 during embryogenesis. Finally, we provide more accurate time-based context for embryonic changes. Ultimately, this basic knowledge should make studies in
Euphilomedes more efficient and enable better experimental design. 27
Chapter 3: Methods
Collection of E. carcharodonta. Euphilomedes carcharodonta were collected at
Pillar Point, California (+37°29′55.22″, −122° 29′44.14″). Sand was collected about 10 meters from shore during low tide using hand nets. Preliminary sorting was accomplished on-site using No. 5, No. 35, and No. 45 test sieves to remove excess sediment. Samples were stored in collected seawater in buckets containing cold packs during transport back to the University of the Pacific, about 2-3 hours. Once in the laboratory, animals in sediment were stored at 12°C with an aquarium bubbler until sorted. Sand was sorted using a dissection microscope and paintbrushes to identify E. morini within the sand. Animals were removed from the sand with transfer pipettes and stored in petri dishes containing artificial seawater at 4°C. Seawater was replaced weekly with additional artificial seawater.
Immunohistochemistry, phalloidin, and DAPI staining. Animals were fixed in
4% PFA overnight at 4°C. Following fixation, animals were washed in PBS 3 x 15 mins.
Animals were then preincubated in the following sequence: 0.1 M NH4Cl for 5 minutes;
0.2% Triton-X in PBS for 10 minutes; cold PBS for 5 minutes. Juvenile instar or adult animals then were treated directly with blocking solution. For embryos, the embryonic membrane was carefully punctured with an insect pin (size 000) or a microneedle prior to pre-incubation to increase antibody permeability (Figure 3). Animals were then blocked in 0.1% BSA in 0.3% Triton-X in PBS for 1 hour. During this time, the primary antibody 28
(Mouse anti acetylated alpha tubulin, Sigma) was diluted 1:20 in blocking solution (5µl in 100µL) in a separate tube. Once finished blocking, animals were then transferred into the primary antibody solution and stained overnight at 12°C on a shaker. The following day, the primary antibody solution was removed, and the animals were washed 3x30 mins in PBS. During washes, the secondary antibody was diluted 1:20 in blocking solution for 1 hr. Following washes, animals were transferred to the secondary antibody solution and incubated at 12°C overnight on a shaker. Prior to mounting, animals were incubated in a DAPI/30% glycerol overnight. Following overnight incubations, animals were incubated for at least 1 hour in 70% glycerol. Animals were mounted in 70% glycerol.
Animals stained in phalloidin were treated similarly to those stained with antibodies. Pre-incubations were carried out similarly, but the membrane was not
Figure 3. Preparation of embryos and agarose-pad mounting. The embryonic membrane was punctured prior to pre-incubations for immunostaining. (A) An insect pin or microneedle was used to puncture the embryo, either on the ventral side to avoid damaging the embryo, or on the dorsal side to avoid damage to the ventral nerve cord. (B) Two layers of lab tape are applied to the edges of a slide. (C) A drop of agarose solution is placed on the center of the slide. (D) A second slide is placed on top to flatten the agarose to an even thickness. (E) The second slide and tape are removed after the agarose sets, and the agarose pad is trimmed into a neat rectangle using a razor. (F) A series of wells are cut into the agar pad using a p200 micropipette tip. 29 punctured using insect pins. Following pre-incubations, animals were placed in a 1:20 dilution of Rhodamine-phalloidin in PBS overnight at 4° or 12°C. Animals were washed
3x15 minutes in PBS prior to incubation in DAPI/30% glycerol for at least 1 hour and up to overnight. Animals were incubated in 70% glycerol for 1 hour prior to mounting in
70% glycerol.
Mounting was performed using agar slides in a fashion modified from methods used to visualize C. elegans (45). Microscope slides were prepared with 2 layers of lab tape on each edge (Figure 3). A drop of 2% agarose solution was placed on the center of the slide, and the drop was flattened with a second microscope slide placed on top (Figure
3). The agarose cooled and set, then the top microscope slide and tape were removed
(Figure 3). Excess agarose was cut away with a razor (Figure 3). A p200 micropipette tip was used to punch holes of the appropriate size in the agarose pad (Figure 3). The agarose pad was covered in 70% glycerol, and embryos were placed into wells. The entire pad was then cover-slipped. Embryos could then be rotated in the wells by sliding the coverslip over the agarose pad, allowing repositioning of the same embryo under the microscope.
Fluorescent microscopy and microphotography. Embryos were visualized under a compound LED microscope (Zeiss Axio Scope.A1) and microphotographs were taken using a Canon EOS Digital Rebel XS camera. Microphotographs were taken at several depths of field, although a specific distance was not used for all specimens.
Images were taken in color. Images were compiled into stacks using ImageJ (Version
1.50i, National Institutes of Health, USA), and merged into composite images using
ImageJ’s Z-projection function, using the maximum intensity setting. Because images 30 were used for reference and not for quantification, DAPI images were converted to black and white 8-bit images for more contrast.
Color staging and photography. Embryos were mounted in a large drop of artificial seawater under a dissection microscope (Jenco). Top lighting was used to illuminate the embryos. A Canon Rebel camera was used for microphotography.
Estimation of brooding duration. 30 females with dark purple broods were placed in separate wells in 12-well culture dishes with artificial seawater. Females were checked regularly (every few days) for hatchings and color changes in embryos were noted. Because females may have oviposited earlier than they were found, the average duration of brooding is a simple estimate.
Figure 4. Normal and malformed embryo morphology. Embryos shown in A/B and in C/D are from single broods at the same age. Normal morphologies are shown in A and C, while malformed embryos are shown in B and D. Malformed embryos cease development and eventually degrade if left in dish. Microphotographs were taken under a compound microscope. 31
Improvement of artificial rearing. Three broods of roughly 20 embryos were split into 2 groups, approximately 9-10 embryos per group. Half of each brood was subjected to experimental “shaker” treatment, while the other half remained in control conditions. In both treatments, embryos were placed in 6-well tissue culture dishes with artificial seawater and were stored in a 12°C refrigerator. All embryos were rotated daily when number of survivors were counted. Dead embryos were identified by malformed morphology (Figure 4). After identification, dead embryos were removed from the dish.
Percent survival was calculated by determining the number of embryos alive compared to the initial size of the group. Percent survival for the three broods was averaged to obtain the average percent survival for the group.
32
Chapter 4: Results
General embryology. Fertilized eggs are approximately 300-350 µm in diameter lengthwise and 250 µm widthwise. Embryos do not undergo a significant change in size over the course of embryogenesis, and remain approximately the same size until hatching from the chorion. Embryos have characteristic appearances as they progress through
Figure 5. Embryos of the ostracod E. carcharodonta. Photographs of live embryos were taken under a dissection microscope with top lighting. (A) Early purple stage. (B) Early purple stage, limb buds visible as light-colored stripes on either side of the embryo. (C) Early orange stage. A thin, transparent band of embryonic tissue is visible wrapping around the orange-colored yolk. (D) Late pre-eyespot stage. The amount of transparent tissue increases, and begins to accumulate at the anterior end of the embryo. Carapace is visible. (E) Late pre-eyespot stage, appendages and carapace are more fully developed than in D. (F) Late eyespot stage, just prior to hatching. Carapace covers almost entirely covers the lateral sides of the embryo, red eyespots are visible. Between A-F, embryos shown are from different broods. Bar, 300 µm. 33 development. The color of the yolk is largely dependent on the lighting source: under a dissection microscope with lighting from above, yolk granules appear purple, but with bottom lighting from below, yolks appear orange-colored. Descriptions on color made are assumed to be under a dissection microscope with top lighting. Following extrusion into the marsupium, eggs were filled with purple yolk granules (Figure 5). As the embryos approach the end of embryogenesis, small red eyespots are visible and movement of the limbs can be observed within the chorion (Figure 5). Finally, the embryos use their furcal claw to tear through the chorion and hatch as first instar juveniles. Over the course of a day, embryos lose their chorion and the carapace expands to cover the soft body parts. The duration of embryogenesis is estimated to be around 70 to 75 days from oviposition to hatching, based on laboratory observations of brooding females. This lengthy timeline, paired with inefficient artificial rearing, has made it difficult to estimate the exact timing of development of E. carcharodonta. In lieu of a timepoint-type developmental study, embryos from various broods were collected and observed, and their general morphology and number of nuclei were used to determine the relative age of the embryo. 34
Early nuclear divisions and cell migrations. The earliest stage observed
contained a single nucleus, as well as three nuclei of a presumptive polar body (Figure 6).
Figure 6. Early embryogenesis visualized with nuclear staining. Embryos stained with DAPI. (A) Single-cell embryo, polar body is present on one face of embryo. (B) Single-cell embryo, nucleus on the opposite side of the embryo relative to polar body. (C) Two nuclei embryo. Polar body is still visible. Polar body visible. (D) Four nuclei, shown later than in C. (E) 8 nuclei. Polar body still visible. (F) 16 nuclei, nuclei have moved superficially. (G) Uncounted, number of nuclei. Polar body no longer evident. (H) Nuclei concentrated to either pole of the embryo. (I) Nuclei continue dividing in a linear pattern, creating a band that extends medially in (J). Nuclei migrate to lateral surfaces in (K). (L) Nuclei continue to divide, covering the embryo (L) and eventually creating an even layer around the yolk (M). The lateral sides of the embryo undulate inwards (N), then create buds (O), which elongate into limb buds (P). pb, polar body. Bar, 100 µm. 35
The polar body appears as a collection of 3 nuclei, and is found on the opposing side of the embryo. Early cell divisions occurred synchronously – in embryos where cells appeared to be dividing (i.e. with metaphase or anaphase appearance) all nuclei exhibited
Figure 7. Asynchronous and synchronous nuclear divisions. Nuclei are visualized with DAPI staining. Dividing cells were identified by their appearance of metaphase or anaphase morphologies. Early divisions occurred synchronously, while later divisions were asynchronous. (A) 16 cell stage with synchronous divisions. (B) Synchronous divisions. (C) Asynchronous divisions. (D) Detail of region shown in C. (E) Asynchronous divisions. (F) Detail of region in (E). Bar in A, B, C, E, 100 µm. Bar in D,F, 30µm. 36 visible signs of active division (Figure 7). Nuclei within the yolk sphere appeared to divide spirally for the first few divisions, according to their arrangement in the yolk sphere. No clear cleavage plane was visible during these early divisions. After approximately 16 nuclei had formed, the nuclei appeared to move superficially and remained on the surface of the embryo (Figure 6). Cells continued to divide synchronously until about 24 cells were visible. Now, nuclei appeared to migrate toward the poles of the embryo, forming rows of nuclei on either end (Figure 6). The nuclei then spread peripherally outward from both poles, creating linear rows of nuclei wrapping around the embryo, resembling a germ band (Figure 6). Eventually, the nuclei migrate toward the lateral surfaces, then continue to spread and cover the entire yolk evenly
(Figure 6). It is apparent that cell divisions in later stages are asynchronous, with only a few cells appearing actively mitotic at any given time (Figure 7). The lateral surfaces form undulations, which deepen, project outward, then form limb buds (Figure 6).
Figure 8. Presumptive fates of limb buds in early embryo. Visualized with DAPI. Embryo shown a purple embryo and is the same as in FigurP. The same embryo is shown in both panels, but limb buds are outlined in panel B. Abbreviations:fa, first antenna; sa, second antenna; mn, mandible; mx, maxilla; fl, fifth limb; fc, furcal claw. Bar 100 µm.
37
Limb bud formation and late development. The fates of the limb buds are based on findings in V. hilgendorfii, but as the first instar morphology of E. carcharodonta has not yet been investigated, they are labeled presumptively (Figure 8).
After limb buds began forming, the carapace began to develop, starting as a small bud on the dorsolateral surface of the embryo (Figure 9). The carapace continues to develop, eventually covering most of the lateral sides of the embryo (Figure 9). Carapace hinge
Figure 9. Carapace development in late embryos. Embryos were stained with DAPI. False coloration was applied to distinguish the carapace and limbs. (A) Dorsal view of embryo with developing carapace, with evidence of a hinge or furrow between the two valves. (B) Left lateral view of embryo from A. (C) Left lateral view of late pre-eyespot embryo with a larger carapace and limbs. (D) Left lateral view of late eyespot embryo with carapace and longer limbs. All images are composites of photos at several planes. Yellow regions, carapace. Blue regions, limbs. Red arrowhead, presumptive carapace hinge. Bar, 100 µm. 38 development is of interest for phylogenetic studies, as its origins may resolve some evolutionary questions concerning the hinged nature of the carapace. Evidence of a furrow or hinge is evident with nuclear staining, but due to the use of DAPI, the true morphology of the carapace cannot be determined definitively (Figure 9).
Improvement of artificial rearing technique. One logistic obstacle in studying
Euphilomedes development is the inefficient and relatively unsuccessful artificial rearing.
Embryos must be removed from the carapace to observe in any detail, but it is not practical to remove only a few embryos at a time without harming the brooding female.
However, once removed from the marsupium, early embryos have incredibly low survival rates, and an entire brood rarely survives the course of a week. This is barely a small fraction of the complete 65- to 75-day duration of brooding. Previously, embryos removed from females were simply stored in artificial seawater in a petri dish, in conditions similar to those used to store living juveniles and adults. However, due to low survivorship, other methods were investigated to improve the survivorship of embryos in artificial rearing conditions.
One method which was thought to aid in survival was rotating embryos. In
Vargula, failure to rotate embryos daily resulted in the embryos adhering to the dish and death (8). Though the exact cause of death was not determined, it is likely that rotation in the dish aids in oxygenation of the early embryos. Brooding behavior in ostracods is not well-studied, but other crustaceans do pass water around embryos and rotate embryos to improve oxygenation (46,47). In Euphilomedes, a daily rotation of the embryo did not suffice to aid in survival (Figure 10). Embryos rotated once daily did not survive longer 39
Figure 10. Average percent survival for artificially reared embryos. 3 broods of early purple embryos were each split into two groups, one to remain in control conditions, and one to be subjected to the shaker treatment. All embryos, including the control condition, were checked daily and rotated with a transfer pipette. Both treatments were subject to the same temperature conditions, at 12C. The shaker group was placed on a horizontal shaker table, while the control group was placed on a stationary shelf in the same refrigerator. Since brood-mates saw significantly different survival rates according to the rearing conditions, inter-brood variability is not the cause of survivorship differences.
than one week (Figure 10). However, when cultured on a flat shaker table, the embryos survived for a significantly longer period of time, with embryos surviving a maximum of
28 days (Figure 10). While the shaker table treatment was not sufficient to maintain embryos throughout the entire duration of embryogenesis, embryos did show physical signs of development during the course of treatment. This is a vast improvement over previous techniques, and would allow researchers to study artificially reared embryos over the course of several weeks instead of several days.
Nervous system visualization. The embryonic nervous system in Euphilomedes was visualized using two techniques. The first was immunofluorescent staining against 40 acetylated-alpha tubulin. Acetylated-alpha tubulin is abundant in developing axons in arthropods, and acetylated-alpha tubulin staining is a common technique for visualizing developing arthropod nervous systems (e.g. in 4–8). This is the first instance of immunohistochemical staining in an ostracod. Staining of embryos using antibodies required physical permeabilization of the chorion using a microneedle. No nervous system staining was observed in primary antibody omission control, although some background staining was observed in both experimental and control groups at and near the site of penetration with the needle.
The second method used to stain the embryonic nervous system was phalloidin staining. Phalloidin stains F-actin, which is also abundant in developing arthropod axons.
However, this stain is only useful for staining early embryos, as it also stains muscle
Figure 11. Embryonic brain in E. carcharodonta labeled with ac-α tubulin immunostaining and phalloidin. The embryonic brain in Euphilomedes has the three typical parts of a crustacean embryonic brain: the protocerebrum, deutocerebrum, and tritocerebrum. (A) anterior view of embryonic brain visualized with ac-α tubulin staining. (B) Ventral view of brain visualized with ac-α tubulin staining. (C) Tissue- squash of phalloidin stain. The three parts of the brain are very distinct in this preparation. Abbreviations: LPC, lateral protocerebrum; MPC, median protocerebrum; DC, deutocerebrum; TC, tritocerebrum. 41 tissue. As the embryos develop muscle in later stages, muscles staining obscures nervous system staining; thus, phalloidin is primarily useful for visualizing the nervous system in only very early embryos.
Figure 12. Central nervous system in Euphilomedes. Ganglia pairs posterior to the circumoral ring are numbered from anterior to posterior. The first two ganglia pairs likely give rise to mandibular and maxillary nerves. Differences in the number of ganglia are more due to the angle of the microphotographs rather than developmental stage, as all embryos shown are of a similar stage, pre-eyespot with limb buds. A, B, C, E are immunostained for ac-α tubulin and are composite images of several focal planes. D is a tissue-squash preparation of a phalloidin stain. Abbreviations: MD, mandibular; MX, maxillary. 42
Brain structure in Euphilomedes embryos. The typical crustacean brain has three parts: the protocerebrum, deutocerebrum, and tritocerebrum. Like other crustacean embryos, E. carcharodonta displays a typical brain structure (Figure 11). The brain and ventral nerve cord can be observed at a very early point in development, while the embryo appears purple and not much embryonic tissue is visible. A maximum of 9 ganglia pairs are visible in the observed specimens; however, due to staging issues, it is not known whether the number observed are simply due to staining and age variability
(Figure 12). A developmental timeline is not shown – rather, the images presented are representative of the embryonic nervous system (Figure 12). Further studies must be done to uncover the developmental process behind the nervous system’s development in
Euphilomedes.
43
Chapter 5: Discussion
This present study on E. carcharodonta is the first to be performed on a
Philomedid ostracod, and presents several new findings regarding ostracod development.
Euphilomedes embryonic development lasts 65-75 days, a much lengthier brood
than most ostracods whose development has been studied. The only other Philomedid
ostracod with a recorded brood time had an even lengthier brood time of 8 months, which
Figure 13. Comparative cleavage and migration patterns in Parhyale, Vargula, and Euphilomedes. Cleavage in the first 2 divisions differ between Parhyale and the two ostracods shown. (A-C) Parhyale demonstrates holoblastic cleavage, while (G-I) Vargula and (L-N) Euphilomedes have meroblastic cleavage. Later divisions and migrations result in cell aggregations, such as the (D) ventral view of soccerball stage and (E) ventral view of rosette stage in Parhyale, (J) cell aggregation in Vargula, (P) or polar aggregations in Euphilomedes. Germ-band structures form in (F) Parhyale (lateral view) and in (Q) Euphilomedes, but no data regarding germ bands is available for Vargula. 44 suggests that brooding in Euphilomedes is perhaps much longer than in other families
(53). Amongst species with embryogenic studies Euphilomedes is a clear outlier; other species such as Vargula developmental timelines which last anywhere from 16-30 days.
While lengthy brooding in Philomedidae does present somewhat of a logistical challenge, it does offer insight into the developmental timing of a slower-to-develop species.
Furthermore, with more practical artificial rearing techniques,
Cleavage patterns in Ostracoda, as discussed before, appear to vary between
Myodocopa and Podocopa. Cleavage in E. carcharodonta most closely resembles that of
V. hilgendorfii. No cleavage furrow was observed in E. carcharodonta, and the arrangement of the nuclei within the yolk also resembles that of V. hilgendorfii. As previously discussed, the ancestral state of ostracod cleavage was one assumed to be holoblastic and equal, based on previous studies on Podocopids alone (36). However, with new insight from E. carcharodonta and V. hilgendorfii, it is evident that a universal cleavage pattern is not applicable to Ostracoda.
Euphilomedes development differs from that seen in other crustaceans. Primarily,
Euphilomedes has two large cell aggregations following cleavage (Figure 13). Other crustaceans, such as Parhyale, do not (54). In Parhyale, cells have a germ disc which forms during the rosette and soccerball stages (Figure 13). This germ disc is the main cell aggregation at these stages, with only a small number of cells located in other areas of the embryo (54). Unlike Parhyale, Euphilomedes has two distinct cell aggregations
(Figure 13). Both aggregations have regular structure, similar to that seen in the germ disc of Parhyale, but they are both of similar size to each other and both have an organized appearance (Figure 13). The presence of two cell aggregations in 45
Euphilomedes also differs from that seen in Vargula (Figure 13). Vargula appears to have only a single large cell aggregation following cleavage (4). However, this stage of development was shown using SEM microphotography, so the location of the nuclei remains unknown. Studies in other ostracods, such as the Podocopid C. littoralis, also do not show the location of nuclei at this stage (29). The Podocopid H. incongruens may show a single cell aggregation similar to the rosette in Parhyale, but the quality of scans from this century old paper make it difficult to confirm (28).
46
Chapter 6: Introduction to Part 2: Differential gene expression patterns in the sexually dimorphic eyes of Euphilomedes ostracods
The development and evolution of complex eyes is a question that intrigues and excites biologists and the general public alike. How could such a complicated organ with highly integrated components evolve? In the last half century a body of theoretical work based on eye morphology has given us solid theories and explanations for some of the eye diversity we see in animals (55,56). Other more recent work has focused on the diversity of opsins, the light-activated proteins found in nearly all eyes (57,58).
Combined with developmental data from a few key model organisms, we can generate plausible explanations for eye evolution from existing genetic cassettes and tissues
(59,60).
At first glance, the complex genetic interactions, cell interactions, and tissue morphogenesis seems irreducibly complex. However, careful dissection of gene regulatory networks (GRNs) and comparative morphology shows us that co-option of existing GRNs to specify existing neuronal/eye anlagen can create new tissues and new pattern elements (60,61). Currently, developmental genetic research like this is primarily confined to a handful of model species. However, it is necessary to extend these findings to other species or we cannot begin to understand the rampant diversity of eyes that we see in nature. One way to look at how GRNs can be co-opted for use in new eye-types is to compare genetically identical or similar individuals with different types of eyes. This allows us to focus on issues of gene expression, rather than the quagmire of gene loss/gain. Our previous work examined the development of eyes in a sexually 47 dimorphic ostracod crustacean, Euphilomedes carcharodonta. We found that compound image-forming male eyes and rudimentary female eyes develop from the same tissue and use most of the same gene regulatory network components to develop (9). Only a small fraction of the genes examined showed differential expression in the two eye-types (62).
Here we examine whether the same genes are differentially expressed in a sister species,
Euphilomedes morini. This will give a rough measure as to how malleable GRNs are over short evolutionary time periods.
Why study sexual dimorphism? Sexually dimorphic species are particularly interesting systems in which to study the evolution of novel traits. In these systems, male and female members of a species have distinct phenotypes from one another (63). These dimorphisms can manifest in many ways, altering behaviors or morphologies between the sexes (64). Sexual dimorphisms are then further categorized into primary, secondary, and ecological sex traits which confer advantages in mating, mate choice and intrasexual competition, and adaptation to sex-specific ecological niches, respectively (63,65).
Most sexually dimorphic traits have been studied in the context of sexual selection. Secondary sex traits increase an individual’s mating success, but do not confer any other adaptive advantages. At times, secondary sex traits can be burdensome for individuals to display, affecting mortality rates and the amount of energy allocated to activities such as foraging (66,67). Both intrasexual and intersexual interactions drive the evolution of secondary sex traits. Intrasexual competition can lead to the evolution of weaponry or larger body size (68). Competitive success can increase an individual’s access to mates or resources, therefore increasing its reproductive success (68).
However, weaponry and large body size can also impose fitness costs, increasing 48 metabolism, decreasing endurance, and attracting parasites (66,69,70). Intersexual interactions, such as mating preference, can drive signaling characteristics which make individuals more attractive as mates (71,72). These signaling traits range from bright colorations to intricate mating displays, and the conspicuous nature of these displays often makes them costly to carry (73,74).
Though sexually dimorphic traits are often associated with sexual selection, not all of these traits are driven exclusively by sexual selection. Ecological pressures can also differentially affect either sex of a species. Resource partitioning drives the evolution of beak shape in some hummingbirds; in one particular species, males and females feed on two different species of flowers and have dimorphic beaks adapted for the shape of these flowers (75). Behavioral differences between the sexes can also change the environmental pressures placed on either sex. In Euphilomedes ostracods, males spend a significantly larger proportion of their lifetime in the water column, while females spend the majority of their lifetime buried in the sand (76). These differing environments, pelagic and sand, subject male ostracods to a higher risk of predation, a risk shown to play a role in the maintenance of sexually dimorphic eyes in this genus
(14).
Although the selection for sexually dimorphic traits has been studied for a wide variety of animals, the genetic mechanisms orchestrating their development are not yet fully understood. Naturally, theories have developed implicating sex chromosomes and, more specifically, sex-linked genes as likely regulators of sexual dimorphism development, because they differ between the sexes (77). However, it is now increasingly evident that autosomal genes play an even greater role, as sex-liked genes usually only 49 comprise a small fraction of the genome (78). Furthermore, sexual dimorphisms are also observed in species without sex chromosomes, or without sex-specific genes (78). In these cases, there cannot be any influence of sex-lined genes on dimorphism development, as no genes in the genome are unique to either sex. These situations demonstrate the importance of autosomal gene regulation in the development of sexual dimorphisms.
The insect doublesex gene in is a powerful example of an autosomal gene regulating development of sexually dimorphic characters. In insects, autosomal genes and alternate splicing direct sex determination pathways (79). Autosomal genes such as doublesex play a role in directing sexually dimorphic traits in Drosophila, as well as in
Onthophagus rhinocerous beetles (80). In these animals, doublesex directs the development of sexually dimorphic characters such as morphology and courtship behaviors in Drosophila, and horns in rhinoceros beetles (81,82).
Although developmental regulators such as doublesex can direct the formation of dimorphic characters, the exact number of genes implicated in differentiating phenotypes between sexes of other animals seems to be situationally dependent and diverse. In some species, the expression levels for numerous genes differ between the sexes (83,84).
Conversely, studies have also demonstrated that a single gene can direct the development of a bipotential tissue in a dimorphic fashion (82,85). Understanding both scenarios and the mechanisms involved can better identify the roles of autosomal genes in directing development of sexual dimorphisms.
Euphilomedes ostracods as a model for arthropod eye evolution.
Pancrustacea exhibits the widest diversity of eye types in the animal kingdom, and the 50 evolutionary origins of such designs is still a topic of debate (16). Myodocopida is the only family of Ostracods (Crustacea) which possesses compound eyes (86). These compound eyes are thought to originate from gene duplication events from median eye genes (16). Median eyes in Myodocopida are thought to be phylogenetically homologous to the median eyes of closely related lineages, but compound lateral eyes are hypothesized to have evolved in the common ancestor of Myodocopids (16).
Euphilomedes ostracods are small Myodocopid ostracods which exhibit sexually dimorphic eye types (19,76). Adult male Euphilomedes possess compound eyes, while females do not and instead have a rudimentary eye (Figure 14) (9,19,76). These sexually dimorphic eyes are quite distinct from one another. While adult males have ommatidial eyes consisting of about 20-33 ommatidia (14), adult female eyes are much smaller and
Figure 14. Sexually dimorphic eyes of the Euphilomedes ostracods. Lateral view of (A) adult female and (B) adult male eyes in E. morini. Right valve has been removed. Males have a large, dark pigmented ommatidial field present, while females have a very small rudimentary eye. (C) Illustrations of the relative eye sizes of juvenile and adult E. carcharodonta. The dark colored field in males contains ommatidia. Females do not have ommatidia during any juvenile stage. E. carcharodonta and E. morini both exhibit comparable degrees of sexual dimorphism. Scale bar: (A,B) 500 mm, (C) 30 mm, 60 mm for male adult eye. Figures from Rivera et al 2009.
51 lack ommatidial structure throughout their development (Figure 14) (9). Males’ eyes also are capable of forming images, and have been shown to play a role in evading predators
(14). On the other hand, female eyes do not possess the necessary components to form images and do not appear to be used for predator evasion, although their function and sensitivity to light has yet to be tested (14,62).
Euphilomedes ostracods have a series of developmental stages in their life cycle.
Following embryonic development, the ostracods molt and grow larger in a series of 5 instar stages before reaching a sexually mature adult stage (19). Male eyes form ommatidia beginning in juvenile instar stage III and conclude eye development at the adult stage (Figure 14) (9). The developmental timing of ommatidia in Euphilomedes makes the system accessible to study, as a variety of free-swimming juvenilles can be collected and studied at one time and without time-course type experiments.
Euphilomedes has XX/XO sex determination, which makes it an ideal system to study the developmental genetics of sexual dimorphism, as differences between male and female eye development will be due to expression differences and not due to the effects of sex-linked genes (14). The clear distinction between male and female eye morphology is also somewhat unique, as it represents the presence or absence of a complex organ, rather than a difference in growth rate or size, as is often studied in regards to sexual dimorphisms (9).
The study of ommatidial development in Euphilomedes has begun relatively recently, and there is no genomic data for this genus at this time. However, transcriptome
52
Figure 15. Comparison of compound eye development in Arthropods. (A) Schistocerca, (B) Tribolium, (C) Drosophila (D) Homarcus americanus and (E) Euphilomedes carcharodonta . Black arrows mark the morphogenic furrow or transitional region, and the grey fill marks patterned regions. (E) Development of a male compound eye in Euphilomedes. The black arrow shows the putative morphogenic front. The red arrow in (D) denotes the mitotic band present in the developing H. americanus eye. A, B, and C adapted and redrawn from Callaerts et al 2006. D based on Hafner and Tokarski. E based on Rivera and Oakley, 2009.
analysis of male juvenile E. carcharodonta eyes has identified several potential genes involved in ommatidial development and in phototransduction (14). These genes are members of highly conserved gene networks whose functions are well-characterized in
Drosophila, and thus are likely to play a role in directing the development of male ommatidia.
Ommatidial development. Despite the diversity of optical designs in arthropods, compound eyes in Arthropoda are all composed of subunits called ommatidia.
Ommatidia in arthropods are rhabdomeric (utilizing r- opsin) and can contain pigment cells, cone cells, and secreted lenses (87). Drosophila eyes contain 750-800 ommatidia
(88), while the simpler compound eyes of Euphilomedes contain about 25-33 ommatidia
(14). Though arthropod eyes vary in optical design and complexity, they all share similar waves of events in regards to proliferation and differentiation (Figure 15) (9,87,89).
In Drosophila, the eye is derived from the antennal-eye imaginal disc, a small collection of epithelial tissue which develops during embryogenesis (90). This imaginal disc is derived from a cluster of cells marked by the expression of Twin of eyeless (Toy) 53 and Eyeless (Ey) (88). Both Toy and Ey are Pax-6 homologs, each with a paired box
DNA binding domain (91,92). Pax-6 genes are widely conserved across the animal kingdom, functioning as master regulators of eye development in various eye types, including those of vertebrates (55). Pax-6 genes are also the upstream regulators in a highly conserved gene regulatory network, the retinal determination network (Figure 2).
Toy acts upstream of Ey, and Ey activates the downstream components of the network, including Sine Oculis (So), Eyes absent (Eya), and Dachshund (Dac) (93).
Following expression throughout the antennal-eye disc, Ey expression then moves exclusively to the posterior portion of the disc, which will subsequently develop into the eye (88). The beginning of pattern formation in the eye is marked by the appearance of the morphogenetic furrow (Figure 15), a wave of differentiation originating at the posterior margin of the antennal-eye disc (94). The morphogenetic furrow moves anteriorly; cells posterior to the furrow begin to differentiate into ommatidial precursors,
Figure 16. Signaling events in the retinal determination gene network (RDGN). Genes controlling cell proliferation are shown in green, and those controlling retinal differentiation in blue. Genes included in this thesis project study are outlined in darker blue. Adapted and redrawn from Tsachaki and Sprecher 2012.
54 and cells anterior to the furrow remain unpatterned (95). The progression of the morphogenetic furrow is driven by the expression of the genes Hedgehog (Hh) and
Decapentaplegic (Dpp) (96,97). Hh is a secreted protein which induces differentiation of eye disc cells into photoreceptors (97). To do so, Hh participates in an autoregulatory loop, activating its own expression in cells anterior to the morphogenetic furrow as is progresses (98). Dpp functions redundantly with Hh and its expression in the developing eye is activated by Hh (98). Additionally, Notch and EGFR signaling pathways are involved in determining the fate of the antennal-eye disc, as well as regulating Hh signaling (88). Posterior to the morphogenetic furrow, the photoreceptor (R) cells differentiate in an organized and sequential manner. R8 differentiates first followed by the pairwise addition of R2 and R5, R3 and R4, R1 and R6, and finally R7 (87,95). After
R7 differentiates, cone cells, primary, secondary, and tertiary pigment cells differentiate, and later develop into the mature ommatidium (87).
Figure 17. Phototransduction cascade in Drosophila. Rhodopsin is coupled to a Gq protein. A light-induced conformational change in rhodopsin activates Gq, which in turn activates a signaling cascade. Phospholipase C is activated by Gq. When active, PLC cleaves phosphotydylinositol biphosphate (PIP2) from the phospholipid bilayer into inositol triphosphate (IP3) and diacylglycerol (DAG). DAG activates PUFA, and together DAG and PUFA activate the opening of the TRPL and TRP channels. Figure adapeted from Tian et al 2012. 55
Phototransduction. Regardless of optic design, an eye performs phototransduction to convert light into an electrical signal which can be processed by the animal. In invertebrates, the signaling cascade is initiated by a conformational change in rhodopsin (99). Rhodopsin is composed of a seven-pass transmembrane protein, r-opsin, and a chromophore (99). Rhodopsin, like other seven-pass transmembrane receptors, is coupled to a Gq protein, which when stimulated activates a signaling cascade (Figure 17).
Once rhodopsin isomerizes, it is then called metarhodopsin; metarhodopsin then activates
Gq by exchanging the GDP bound to Gq with GTP, converting Gq to its active form (99).
Gq then activates phospholipase C, which catalyze the conversion of phosphatidylinositol biphosphate (PIP2) from the phospholipid bilayer to form inositol triphosphate (IP3) and diacyl glycerol (DAG) (99). DAG and downstream effector PUFA then activate the opening of TRPL and TRP, allowing Na+ and Ca2+ influx (99).
A timely deactivation of cation influx is crucial to avoid negative effects of prolonged cation influx and to prepare the signaling cascade for a new stimulus (99).
Arrestins displace Gqα to bind with rhodopsin, and Gqα terminates its own activity, with intrinsic GTPase activity converting the bound GTP to a GDP, returning the entire cascade back to an unactivated state (99).
Morphological differences arise from similar genetic backgrounds. One of the attractive features of studying Euphilomedes ostracods is the genetic similarity in males and females. As mentioned previously, Euphilomedes ostracods use XO/XX sex determination, and so males and females have the same genetic background. Therefore, we assume that morphological differences between the sexes must be due to differential 56 expression between the sexes during development, as there are no genes that are exclusive to either sex.
Another well-studied animal which uses XO/XX sex determination and has sexually dimorphic phenotypes is the nematode Caenorhabditis elegans. In C. elegans, hermaphroditic and male phenotypes are the result of differential gene expression of a shared set of genes. The sys-1 gene, for example, is required for the generation of distal tip cells in hermaphrodites, which are necessary for correct polarity and growth of the germ-cells, as well as the development of the somatic gonadal primordium (100). This gene is not necessary for these functions in males – the sys-1 mutant allele is not completely penetrant in males (100). Males do not show gonadal defects to the same degree as hermaphrodites – they can produce progeny, while mutant hermaphrodites are sterile (100).
Another instance where polyphenisms emerge from similar genetic backgrounds is in the queen-worker dimorphisms in hymenopteran eusocial insects, such as the honeybee Apis mellifera. This species uses an interesting system of sex determination, relying on heterozygosity at the Sex determination Locus (101). Heterozygosity at this locus to specifies the larger and reproductively active queens, while homozygosity or hemizygosity at this locus specifies smaller male bees (101). In combination with nutritional inequality, differential gene expression has shown to play a large role in determining caste roles during larval development in these animals (102,103). In larval queens, higher feeding is associated with an upregulation of metabolic genes allowing for larger body size, while in workers, a larger suite of novel genes are upregulated (104). 57
Evidence of widespread DNA methylation sites in A. mellifera provides a likely control mechanism for the differential gene expression during larval development (103).
Can the same morphology be determined by different gene expression patterns? Perhaps less intuitive is the concept of deriving the same end-point phenotype from different developmental processes. A prime example of this is germ band formation and segmentation in insects. Insects share similar segmented morphologies, yet several developmental methods for specification of segments exist (105). These are categorized as short, intermediate, and long germ band insects (105). In long band insects, such as
Drosophila, the entire body plan fate map is established simultaneously during the blastoderm stage (105). In intermediate and short band insects, only the anterior portions of the body are specified during this stage; these insects will undergo a second wave of specification which will pattern the posterior segments (105).
Though conservation of an endpoint phenotype, like segmentation pattern, in the presence of evolving developmental patterns is a perplexing conundrum, it nevertheless exists. The developmental flexibility of a phenotype, or rather, the fact that several developmental patterns can result in the same phenotype, is a primary interest of this comparative study on Euphilomedes eye development.
Studying differential gene expression in Euphilomedes. Understanding gene expression differences in the developing eyes of male and female Euphilomedes ostracods is key to understanding how they regulate development. Two species, E. morini (Santa Barbara, California) and E. carcharodonta (Half Moon Bay, California)
(19), are discussed in this study. Both species exhibit a comparable degree of sexual dimorphism, and ommatidial development happens along the same developmental 58 timeline in both (9). Previous analysis of the E. carcharodonta juvenile male eye transcriptome revealed orthologs of genes well-characterized in Drosphila eye development and phototransduction (14).
A study by Sajuthi et al. on E. carcharodonta focused these gene homologs in developing male and female eyes (62). mRNA expression analysis revealed expression differences between male and female eyes during development (62). Dachshund, shaven, and chaoptic homologs were shown to be upregulated in male eyes when compared to female expression levels (62). Dachshund is involved in early patterning of the eye as part of the retinal gene determination network (Figure 16). Shaven, a Pax2/5/8 homolog, is involved in specification of different ommatidial cells (106), while chaoptic is involved in directing the arrangement of photoreceptor cells within the ommatidium (107). In addition to developmental gene expression difference, phototransduction genes were also upregulated in adult males (62). This is not surprising as males are the only sex with ommatidia, and therefore possess more photoreceptive tissue than females. Despite these few developmental and phototransduction genes which showed expression level differences, the majority of genes surveyed did not show significant differences in male and female eyes (62). Sajuthi’s study focused on global expression throughout the eye for all genes studied, and it is important to note that many eye development genes are expressed in specific locations and patterns within the developing eye (e.g. eyeless, hedgehog). Therefore, some genes that showed overall similar expression levels in male and female eyes during this previous study may actually have differing expression patterns within the eye. 59
The expression differences found in E. carcharodonta provide trends to compare to and to dissect further. It is not known whether the genetic mechanism directing E. carcharodonta’s dimorphic eye development is conserved outside of this species, or to what degree it is genetically flexible. If the genetic pattern observed in E. carcharodonta is indeed conserved in other genus members, such as E. morini, this would suggest that
Euphilomedes’ dimorphic eyes develop according to a very specific genetic scheme.
However, if the mechanism is not conserved outside of the species, this would strongly suggest that eye development is genetically flexible in the Euphilomedes genus, and would demonstrate how different genetic programs can result in the same morphologies.
This study will compare male and female eye development and phototransduction gene expression in the developing eyes of Euphilomedes morini. This study, study like the previous study in E. carcharodonta, will quantify expression levels of both developmental and phototransduction genes by comparing mRNA expression levels for both sexes at all developmental stages. Additionally, this study will also quantify protein expression of several of these developmental and phototransduction genes to confirm that protein expression trends match those found for mRNA. Finally, this study will compare expression trends found for E. carcharodonta and E. morini. 60
Chapter 7: Methods
Collection of Euphilomedes morini. Euphilomedes morini were collected at
Stearns Wharf Pier (34° 24′ 38.41″ N, 119° 41′ 14.5″ W). and Goleta Pier (34°24'53.1"N
119°49'44.0"W), both in Santa Barbara, California. Sand was collected off the pier using a standard Ekman grab (Wildco) and preliminary sorting was performed on-site using a
No. 5, No. 35, and No. 45 test sieves to remove most of the fine sediment (W.S. Tyler
Industrial Group). Animals and remaining sediment were stored in a partially covered bucket filled with collected seawater. The bucket was surrounded externally with ice during return transport to the University of the Pacific, about 10 hours. Once in the laboratory, animals in sediment were stored at 12-14°C with an aquarium bubbler until sorted. Sand was sorted using a dissection microscope and paintbrushes to identify E. morini within the sand. Animals were removed from the sand with transfer pipettes and stored in petri dishes containing artificial seawater at 4°C. Seawater was replaced weekly with additional artificial seawater.
Dissection. Euphilomedes morini were staged and dissected under a dissecting microscope using size 1 enameled insect pins. Animals were staged in a manner similar to the protocols described in Sajuthi et al, 2015, with modifications for the morphology of
E. morini. Eyes were severed from the eye stalk and placed immediately into 800 μL of
Trizol (Life Technologies). Eyes in Trizol were stored at 4°C. The eyes from a total of
10 animals of the same stage and sex were pooled into each sample for a sufficient amount of tissue for RNA isolation. 61
RNA isolation and cDNA synthesis. RNA was isolated using modified manufacturers protocols. 5 μL of glycogen and 160 μL of chloroform were added to the
Trizol and eye samples. The samples were shaken vigorously then incubated 5 minutes at room temperature. Samples were centrifuged in a 5430 centrifuge (Eppendorf) at
13200 rpm for 15 minutes at 4°C. The aqueous layer was removed, and 40 μL 3M sodium acetate was added and the sample was vortexed briefly. Then 300-400 μL isopropanol was added, and the solution was precipitated by centrifugation at 13200 rpm at 4°C for 15 minutes. The supernatant was removed and the pellet was washed with 1 mL of cold 75% ethanol. The samples were centrifuged for 10 minutes at 4°C, and the supernatant discarded. The samples were allowed to air-dry and the isolated RNA was resuspended in 30uL nuclease-free water. Following RNA isolation, samples were treated with the DNAse Max Kit (Mo Bio Laboratories Inc.) according to the manufacturer’s directions. DNAse-treated RNA was stored at -80°C.
RNA was quantified with a Qubit Fluorometer (Life Technologies) using manufacturer’s protocols for the Broad Range RNA Detection kit. 40 ng of RNA was used in each 20 μL reaction for cDNA synthesis. cDNA was synthesized using the
Superscript III First Strand Synthesis System (ThermoFischer Scientific) following the manufacturer’s protocols for first-strand synthesis using random hexamers. cDNA was used immediately for qPCRs or stored up to a week at -20°C.
Generation of qPCR standard curves. Using the published transcriptome and annotations of Euphilomedes carcharodonta, a sister species to our model, Euphilomedes morini, we designed primers against several genes. E. morini degenerate primers for
Shaven were designed using the published sequence of E. carcharodonta Shaven. The 62 primers were synthesized by Integrated DNA Technologies. PCRs were performed using
E. morini cDNA as a template and using GoTaq Master Mix (Promega). The cycling parameters were as follows: 94°C for 30 s, 55°C for 30 s, 72°C for 1 min for 24 cycles; followed by 95 °C for 30 s, 45 °C for 30 s, 72 °C for 2 min for 12 cycles; followed by
72°C for 10 mins.
PCRs resulting in bands of an appropriate size were ligated into pGEM-T Easy
(Promega) and transformed into TOP10 or DH5alpha competent cells following manufacturer’s protocols. Plasmids were purified using the GenCatch Plasmid DNA
Mini Preparation kit (Epoch Life Sciences). Isolated plasmids were sequenced by
Sequetech (Mountain View, CA) and analyzed with CLC Bio Workbench software (CLC
Bio). The sequence data was used to design qPCR primers in primer 3 to amplify a region of about 50-100 base pairs.
To make standard curves for qPCR, three plasmid standards were used per gene at 1:10 dilutions, beginning with a concentration of 0.1-200pg/μL.
Each PCR reaction contained 2 μL of cDNA or plasmid standards, 0.4 μL of the forward qPCR primer (10µM), 0.4 μL of the reverse qPCR primer (10µM),10 μL of
SybrGreen (Invitrogen), and 7.2 μL of water. The reactions were run with a StepOnePlus
RT-PCR Thermocycler Block (Applied Biosystems) using the StepOne Software and the following program: 94°C for 30 min; followed by 94 °C for 15 s, 52 °C for 1 min, 72 °C for 30 s repeated for 40 cycles; followed by 72°C for 5 min; followed by a final melting curve protocol: 94°C for 15 s, 60°C for 15 s, and 94°C for 15 s. Runs with low efficiency
(<80%) or a low R2 value (<0.98) were discarded and omitted from analysis.
Additionally, thresholds were set to discount any samples that went into exponential 63 phase after cycle 35. The last set of cycles was to amplify any spurious secondary PCR products for detection in Melting Temperature analysis. We used NORMA-Gene to normalize data (108). NORMA-Gene was also used to determine which replicates were considered too variable. Replicates which had high variation are shown in grey in
NORMA-Gene’s results. This grey color is associated with a standard deviation larger than 15% of the mean value.
64
Chapter 8: Results
Gene expression differences. The data from our trials were not consistent enough for use in drawing conclusions. Technical replicates as well as biological replicates had high amounts of variation, defined as standard deviation as 15% of the replicate mean or greater (grey cells in tables). This is likely due to use of old samples.
During this study, field collection of E. morini was relatively unsuccessful, and only
Table 1. Expression data for TOY. Data from qPCR experiments normalized with NORMA-Gene. Replicates are shown in rows next to each other. Technical replicates with high variation (standard deviation > 15% mean) are shown in grey.
8/8/ 8/18/ 8/19/ 8/23/ 8/30/ 9/29/ 9/29/ 10/27/ TOY 2016 2016 2016 2016 2016 2016 2016 2016 Adult 1.8E- male 06 0.000228 0.000905 0.000336 0.026533 0.000421 0.000355 1.75E- 06 0.000414 0.000999 0.000631 0.507857 0.000561 0.000662 5.06E- 5 male 06 0.000121 0.000182 0.000152 0.496517 0.000573 4.83E- 06 0.000369 6.28E-05 0.001449 0.6679 0.000588
4 male 0.000471 0.000453
0.000173 0.000356 Adult female 0.001804
0.002534 Early 5.94E- embryo 06 0.001548 0.001036 0.003285 0.001661 9.1E- 06 0.000782 0.000347 0.001661 0.001312 Late embryo 0.000108
0.000746
65
Table 2. Expression data for ELAV. Data from qPCR experiments normalized with NORMA-Gene. Replicates are shown in rows next to each other. Technical replicates with high variation (standard deviation > 15% mean) are shown in grey. 8/8/ 8/18/ 8/19/ 8/23/ 8/30/ 9/29/ 9/29/ 10/27/ ELAV 2016 2016 2016 2016 2016 2016 2016 2016
adult male 0.000346 0.637737 18.3731 3.057806 0.178073 0.022019 0.022005 0.00052 1.131742 8.70824 1.786289 0.09591 0.035546 0.053654 5 male 0.000368 0.0554 46.9101 5.343876 0.154832 0.000731 0.089652 68.4985 2.513143 0.244241 4 male 0.151696 0.00193 0.125847 0.003123 Adult female 0.006742 0.0048 Early embryo 0.001319 0.00273 21.28092 16.1459 0.001474 0.002124 0.005402 63.58176 30.78398 0.001867 Late embryo 0.002936 0.004989
small numbers of animals were found during each collection. Because eye tissue samples
were pooled samples of around 10 animals, incomplete samples often remained in storage
for several weeks, if not for months, before RNA extraction. This storage seems to have
led to degradation of samples. Furthermore, storage of RNA after extraction could also
have experienced degradation during storage. Other sources of error could include
contamination of reagents, such as in primers or water. Pipetting error is also another
potential source of error, but is unlikely, as quantification values and R2 values for
standards did not see the same degree of variation between replicates. Ultimately, the
most likely source of error is aging samples, so future studies must: 1) use freshly
dissected samples as soon as possible for analysis; 2) repeat trials from this present study
to obtain more reliable data. 66
Table 3. Expression data for PLC. Data from qPCR experiments normalized with NORMA-Gene. Replicates are shown in rows next to each other. Technical replicates with high variation(standard deviation > 15% mean) are shown in grey.
PLC 8/8/2016 8/18/2016 8/19/2016 8/23/2016 8/30/2016 9/29/2016 10/27/2016 adult male 0.000557 1.48E-05 5.9E-06 1.42E-05 0.787272 0.0064 0.000973 4.58E-06 1.13E-05 1.29E-05 0.076366 0.001409 5 male 0.002333 5.03E-06 0.007624 2.074397 0.003434 2.71E-05 0.001351 1.46E-05 0.000546 3.279094 0.002124 2.13E-05 4 male 9.12E-06 0.010132 1.84E-06 0.033153 Adult female 0.005485 0.002549
Table 4. Expression data for DAC. Data from qPCR experiments normalized with NORMA-Gene. Replicates are shown in rows next to each other. Technical replicates with high variation (standard deviation > 15% mean) are shown in grey.
DAC 8/23/2016 10/20/2016 10/23/2016 10/27/2016 Early embryo 1.401186 9.02E-07 1.06E-07 2.03E-08 1.453052 1.25E-07 9.83E-08 2.03E-08
Table 5. Expression data for SIA. Data from qPCR experiments normalized with NORMA-Gene. Replicates are shown in rows next to each other. Technical replicates with high variation (standard deviation > 15% mean) are shown in grey.
SIA 10/20/2016 10/23/2016 10/27/2016 Late embryo 5.95E-07 1.32E-06 4.48E-05 4.3E-06 1.42E-06 3.8E-06
67
Table 6. Expression data for SO. Data from qPCR experiments normalized with NORMA-Gene. Replicates are shown in rows next to each other. Technical replicates with high variation are shown in grey.
SO17 8/8/2016
Adult male 0.005734 0.002242 5 male 0.015128 0.013788 Early embryo 0.006615 0.002682
68
Chapter 9: Discussion
Unfortunately, the quality of the data collected in the present study were insufficient to make reliable comparisons of gene expression during eye development.
Future studies must use fresher tissue samples to determine expression differences.
An alternative technique which may prove promising for quantification could be mass spectrometry and proteomics. In addition to establishing an Euphilomedes eye proteome, mass spectrometry could provide important information on the quantity of protein expression in eye tissue, and verify that trends seen in RNA expression directly translate to protein expression.
Aside from quantification studies, further directions of this research could include immunohistochemistry or in-situ analysis to visualize the location of gene expression.
These staining techniques could reveal whether gene expression in the developing
Euphilomedes eye has specific spatial patterns. This is a likely scenario, as many genes in the developing Drosophila eye, as discussed previously, have very specific expression patterns, and the location of expression is crucial in specifying the correct fields for specification (88). This could reveal gene expression differences that were not initially apparent with quantification studies – for example, two tissues may express the same quantity of RNA, but the localization of the expression could differ, a subtlety lost by homogenization of the eye tissue.
69
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