Course Structure and Modules
Hydra is a freshwater cnidarian polyp, easily reared in the lab, which was identified by Abraham Trembley (1744) as the first animal able to fully regenerate any missing part of its body. Trembley also found that Hydra is a carnivorous animal that actively catches its food, walks and reacts to physical stimulation by contracting its body. Since then Hydra proved to be a powerful model system to investigate the molecular and cellular basis of stem cell biology, asexual reproduction (budding), body regeneration and low senescence in a non-bilaterian context. One major discovery was made by Ethel Browne (1909) who used pigmented and unpigmented animals to show that restricted regions in intact or budding or regenerating animals, nowadays named organizers, induce ectopic axis formation upon transplantation by recruiting cells from the host. Organizers were later evidenced in developing chordates such as the gastrula organizer (Spemann and Mangold, 1924 – Nobel Prize 1935). The cell biology is equally fascinating in these animals as three continuously cycling stem cell populations exhibit an unusual cell cycling behavior and a tight spatial regulation. Epithelial stem cells provide a high resistance to starvation and stress, while interstitial stem cells allow de novo neurogenesis and gametogenesis whatever the age of the animal. Modellers of developmental processes were inspired by Hydra to propose as general model of patterning the reaction-diffusion model (Turing, 1952, Meinhardt and Gierer, 1972) or the French flag model (Wolpert, 1969). Over the past 15 years, beside omics data, functional genomics were made possible with stable transgenic lines, gene silencing through RNAi and Crispr-Cas9 strategies.
In this module, students will observe the natural behavior and developmental processes in intact and bisected Hydra, identify the aging phenotype developed by some strains, follow the autophagy flux with the help of an autophagy sensor. They will also use transgenic lines to perform transplantation experiments and evidence the activity of organizers, to produce reaggregates– natural organoids – that undergo regeneration, to monitor through live imaging the wound healing process when ROS production is altered. Although the constraints of a short module will not allow sufficient time to perform gene silencing experiments, the students will have the opportunity to be creative in their experimental design to investigate the amazing developmental plasticity of this animal.
Planarians are free-living representatives of the phylum Platyhelminthes, a group of some 50,000 species of flatworms. Flatworms are among the simplest bilaterally symmetric animals: they are acoelomates, yet they possess derivatives of all three germ layers organized into complex organ systems. Thus, Platyhelminthes have been thought to occupy an important position in Metazoan evolution. Current models place the Platyhelminthes in a large assemblage of protostome invertebrates, known as the Lophotrochozoa, a sister group to the Ecdysozoa (to which insects and nematodes belong).
Planarians are best known for their capacity to regenerate complete individuals from minuscule body parts, as well as for their ability to “de-grow” when starved. Such extraordinary plasticity in the adult is in direct contrast to the rigidity displayed by currently used invertebrate models such as Caenorhabditis elegans and Drosophila melanogaster. The difference lies in a population of adult somatic stem cells, called neoblasts, that are distributed throughout the planarian body. Neoblasts are the only mitotically active cells in planarians, and their division progeny generate the 30-40 different cell types found in these organisms. In intact planarians these stem cells replace cells lost to normal physiological turnover; whereas, in amputated animals, they give rise to the regeneration blastema, the structure in which missing tissues are regenerated.
Until the mid-20th Century, planarians were a key model for studying development and regeneration. Yet, as attention shifted towards animals amenable to classical genetic analysis, the use of planarians declined. Recently, however, the successful introduction of cell, molecular, and RNAi techniques in planarians, along with heightened interest in stem-cell biology and the plasticity of the differentiated state, has re-kindled interest in these fascinating organisms. Part of this renaissance, includes an ongoing Genome sequencing project, being carried out by the University of Washington Genome Sequencing Center in St. Louis, MO (http://genome.wustl.edu/projects/planarian/). The species Schmidtea mediterranea was selected for sequencing since it is a stable sexual diploid with a genome size of approximately ~8 x 108 bp. S. mediterranea provides a vital resource for the development of a unique model to study metazoan evolution, regeneration, and the regulation of pluripotentiality. Mechanistic insights into these basic biological problems will have deep and obvious implications to our understanding on biology and perhaps for the improvement of human health.
In this module students will learn many techniques and biological phenomenon used to study planarians, including amputating animals for regeneration analysis, injecting animals with dyes, performing tissue transplantation, observing wild type regenerating animals, watching neoblast deficiency phenotypes, observing RNAi-induced phenotypes in Schmidtea mediterranea, examining planarian embryos, performing antibody staining, observing other planarian species, and learning cell isolation.
In 1997, a new molecular phylogeny revealed that C. elegans and Drosophila were much more closely related to each other than had been thought previously. Before then, Drosophila was thought to be more closely related even to humans than to C. elegans. The 1997 work placed the nematodes (which include C. elegans) and arthropods (which include Drosophila) together in a clade now known to include six other animal phyla. These eight phyla together are named the Ecdysozoa, or molting animals. Ecdysozan phyla closely related to the nematodes and arthropods might serve as valuable models for evo-devo biology and for modern comparative biology more generally, assuming it would be possible to find a tractable lab model among these animals. Animals related to but not within the nematodes and arthropods might be used to take advantage of having two reference model systems — C. elegans and Drosophila. In the long term, the use of a relative of C. elegans and Drosophila might dramatically expand the set of genes and mechanisms that are of interest for comparative studies beyond the narrower set of genes and mechanisms that are known to have conserved functions only across a greater breadth of animal diversity. But little modern work had been done in any of the phyla closely related to nematodes and arthropods, suggesting that it might be necessary to develop a new model. One ecdysozoan phylum in particular, the tardigrades, has useful characteristics for lab study.
Tardigrades, also known as water bears, are a phylum of microscopic, eight-legged animals estimated to include thousands of species, over a thousand of which have been described to date. One species, Hypsibius exemplaris, can be cultured in the lab, frozen as live stocks, and has a short, 12-day generation time. Embryos are laid by mothers just before molting, and the embryos are left behind in the clear cuticles, making it easy to film synchronous batches of embryos.
Tools for studying Hypsibius exemplaris have now been developed, and these animals can be used as an evo-devo model and as a model for understanding how living materials can survive extreme conditions. In this module, students will get their own culture of this species, and they’ll learn to fluorescently mark components of cells and film embryogenesis. Students will also have an opportunity to collect wild tardigrades along with the other kinds of microscopic animals that share an unusual superpower: they all survive desiccation. One of the joys of working with an emerging model system is that many of the things we decide to look at have never been seen before, so students will be able to try many new things.
The chick (Gallus gallus) is an excellent model system to study vertebrate embryogenesis and organ formation. It is one of the major amniote model systems used in developmental biology, the other being the mouse. Unlike anamniotes (amphibians and fishes), amniotes generally produce very few eggs, which they protect either by a shell or by keeping the embryo inside the mother. The amniotic membrane envelops the embryo to provide a fluid environment protecting the embryo from drying out, while the evolution of the yolk sac endoderm allows the embryo to take up nutrients and oxygen from outside the embryo. These ‘amniote inventions’ allow embryos to grow for a long time in their protected environment before hatching or birth, and unlike anamniotes, to activate the zygotic genome immediately. Unlike the mouse, the early anatomy and tissue organization of chick embryos is very similar to that of human embryos: both initially develop as a flat disc of cells, the blastoderm, and gastrulation occurs through the primitive streak rather than a blastopore.
Fertilized chick eggs are readily available and the equipment needed is minimal – a humidified incubator (380C, no CO2 required), a dissecting microscope, simple microsurgical tools, and either a hand-held mouth pipette or a micromanipulator and picospritzer are used for electroporation and labelling. The eggs can be stored (usually at 150C) for up to 1 week before use.
The chick embryo is easily accessible and comes encased in its own container, the hard eggshell. Through a hole in the eggshell, the embryo can be visualized and easily manipulated with microsurgical tools or gene constructs, the egg is then sealed and allowed to continue development in ovo to determine the consequences of the experimental manipulation. Alternatively, various culture methods are available for early embryos that allow ex ovo growth from stages just after laying to embryonic day 3-5 (or longer) depending on the method.
The main strength of the chick is that the embryos are fairly large, making tissue transplantation and manipulation experiments possible at many different stages, and that gene manipulation can be performed in a temporally and spatially controlled way. “Cut-and-paste” experiments have been used very successfully to determine e.g. tissue interactions controlling organ development, time of commitment etc. paving the way for many fundamental concepts about development. These “cut-and-paste” techniques can be learned relatively easily, and can be combined with molecular gain and loss of function experiments to test both sufficiency and necessity for a molecule of interest or with reporters for gene expression and signaling readouts.
The most important advantage of the chick is that molecular manipulations can be done in a temporally and spatially controlled manner. Gene misexpression or knock down is achieved by electroporation to introduce antisense oligonucleotides (morpholinos), sh-RNA or plasmids containing a cDNA of interest into a particular tissue at a particular time. This allows us to study gene functions relevant to the process under investigation, even for genes that are used multiple times at different developmental stages. This is different from systems where constructs have to be injected into 2-4 cell stage embryos, or from gene inactivation using genetics, where the phenotypes observed at late stages can represent the cumulative effect of many processes gone wrong at different times and in different places. Overall, the speed at which these experiments can be done, owing to the ease of manipulation and the large numbers of eggs that can be obtained, makes the chick embryo a powerful system to test gene function.
More recently, the chick has emerged as a very rapid system to test the activity of regulatory regions, without the need to generate transgenic lines. Electroporation of cell specific enhancer-GFP constructs are also a powerful tool to label specific cell populations and e.g. to isolate such cells for transcriptome analysis or similar molecular approaches.
The genomic resources available for the chick have also increased rapidly. The chick genome has been sequenced and large collections of chicken ESTs are available (http://www.chick.manchester.ac.uk, http://www.chickest.udel.edu). Affymetrix sells chick DNA microarray chips for transcriptome profiling and RNAseq technology is possible from dissected tissue or FACS-sorted cells, making gene discovery much easier. There are also public resources providing useful links to many databases and cataloguing gene expression (http://geisha.arizona.edu/geisha/). With only 1×109 nucleotides the chick genome is more compact than the mouse or human genome (1/3 of the size) containing less repetitive regions and spacers. The chicken evolved 85-90 million years ago from a galliform ancestor (van Tuinen M, Dyke GJ. Mol Phylogenet Evol. 2004, 30:74- 86), yet the tissue and molecular interactions are highly conserved with mammals. Thus, genomic comparisons that include chicken sequence data aid tremendously e.g. in the search for evolutionarily conserved regulatory elements.
A disadvantage of the chick embryo is the relative lack of genetics. Although it has been possible to make transgenic chickens, the time to sexual maturity and the space required to maintain a flock of chickens places it beyond the normal capacity of a lab to rear their own experimental stock. However, GFP-transgenic chicken eggs can now be obtained, different transgenic reporter lines are now available (e.g. Notch-reporter) and several different quail transgenic lines are now available in the USA.
During the course, students will learn in ovo and ex ovo methods, some of the most commonly used transplantation experiments, and various ways of manipulating gene expression. Results will be analyzed using imaging, video time lapse movies, antibody staining, in situ hybridization, or through many other techniques.
Understanding how genes control growth and development in mammalian embryos is driven by our curiosity of how human form is generated and how it evolved from much simpler organisms. During the past century, the mouse has become firmly established as the primary experimental mammal, due to its small size, resistance to infection, large litter size and relatively rapid generation time. Mice were also favored because of an interesting collection of mutations affecting coat color, hair morphology and pigmentation that were readily available from “mouse fanciers”, breeders and collectors of interesting pet mice. Together with the rediscovery of Mendel’s laws of inheritance in the 1900’s, this initiated the study of mammalian genetics. Subsequently, the mouse was the first species whose genome was genetically modified by transgenesis, and the first species in which a gene was knocked out. These are indeed wonderful times to be studying mouse genetics as the mouse germ line can be experimentally manipulated in just about every conceivable way, including direct injection of DNA into zygotes, genetic modification of embryonic stem cells, via TALEN and CRISPR mediated genome editing, and by chemical or irradiation mutagenesis of spermatogonial stem cells.
Mammalian embryology in contrast is a much older field, and historically was closely associated with the study of human and veterinary reproductive physiology. In more recent times, the mouse has been instrumental in the discovery of imprinting, in the development of methods for in vitro fertilization, the discovery, maintenance and manipulation of embryonic stem cells, the demonstration of reprogramming via the generation of induced pluripotent stem cells, cloning via somatic cell nuclear transfer, and the development of hybridoma cells, providing a never ending source of antibodies.
In this module students will learn how to properly handle, anesthetize and euthanize mice as well as perform surgeries including oviduct injection and transfer, and tissue transplantation under the kidney capsule. Students will have the opportunity to learn how to edit the mouse genome using the CRISPR/Cas9 system in preimplantation stage mouse embryos, both in vivo and in vitro. Students will learn how to isolate, culture and manipulate, post-implantation whole mouse embryos and their organs; how to manipulate key signaling pathways in vivo; and how to visualize gene products, cells and tissues of the skeleton, musculature, vasculature and nervous systems. Through a subcomponent of the module, known as the “Zoo Lab” students will have the opportunity to explore evolutionary development and comparative anatomy questions in a diverse range of species that typically includes mouse, lizard, chameleon, snake, bat, snail, turtle and pig embryos.
Zebrafish (Danio rerio) are a small tropical fish native to southeast Asia. With the goal to employ genetics for understanding the development of the nervous system, George Streisinger recognized in the 1970s the potential of zebrafish and established it as a model for vertebrate genetics. Since then they have become a prominent vertebrate model for basic genetic and developmental biology and behavioural studies, and more recently for phenotype-based drug screening. This is due to the ease of embryological and genetic manipulation and their small size and relatively simple husbandry.
Large scale genetic screens have identified hundreds of mutants to study a wide variety of biological processes, most prominently developmental biology, as well as models for human disease. The CRISPR/Cas9 genome editing revolution enabled the generation of targeted gene knock-out and knock-in zebrafish lines, enhancing the reverse genetics potential of zebrafish in basic and biomedical research.
In contrast to other genetic vertebrate systems, zebrafish have the capacity to regenerate the majority of tissues, including for instance the heart and sensory hair cells, making them exciting models to investigate why humans have lost this potential.
A unique feature among vertebrates is that zebrafish embryos and larvae are transparent and together with their small size allow the live observation of dynamic developmental and regenerative processes, including those of internal organs. The availability of a plethora of transgenic fluorescent reporter lines for various cells and tissues (e.g. heart, blood vessels, liver, sensory hair cells and neuronal subpopulation), cellular compartments (e.g. membrane, nucleus, cytoskeleton), as well as intra-cellular signaling indicators (e.g. calcium-signaling) and more recently optogenetic tools has opened the door to imaging fast dynamic events at subcellular resolution.
In this module students will learn how to handle, stage and anesthetise zebrafish embryos, including micro-injection at the one-cell stage. A large variety of RNAs, DNAs for fluorescent labelling of single cells or subcellular structures will be available for injection. Likewise, different methods for interfering with gene function will be introduced: antisense morpholino oligonucleotides for gene knockdown and CRISPR/Cas9 genome editing for gene inactivation. The latest insights into the biology and possible advantages and disadvantages of either technique will be discussed. Moreover, students will be able to stimulate or block major signalling pathways during embryo development in a temporally precise fashion by drug incubation. In addition, students will learn to transplant labelled cells between embryos, a powerful method to test cell autonomous and non-autonomous gene functions and cell behaviours. Utilizing the transparency of the embryos, students will be taught how to live-image entire embryos, as well as dynamic processes such as cell migration of fluorescently labelled cells using the latest models of state-of-the-art microscopes and equipment, such as light-sheet, spinning disc and confocal microscopes. Using this diverse set of tools students will design and investigate their own research question.
The first thing that springs into mind when thinking about amphibian studies is likely the induction of twinned axes by Spemann-Mangold organizer graft. However, this is but only one example of a long list of important contributions of amphibian research to embryological concepts. Pioneering experiments from Holtfreter brought focus on cell adhesion and movements as well as the concept of developmental potentials, and Nieuwkoop’s tissue separation, recombination and transplantation studies broadened the views of embryonic induction and patterning.
These classical experimental embryology studies rely on a salient feature of amphibian embryos that parts of the embryos (explants) can be isolated and cultured in simple salt solutions for developmental programs to continue. The “cut-and-culture” and “cut-and-paste” experiments are carried over to the modern days of amphibian research and remain a crucial tool in embryo manipulations nowadays. The toolset is further enriched by incorporation of advanced molecular and cellular technologies, such as using fluorescence microscopy to investigate cell morphology, cytoskeleton dynamics, cell and tissue movements, and signaling mechanisms.
The ease to introduce exogenous molecules into early embryos allows gain- and loss-of-function studies. The application of CRISPR/Cas9-mediated genome editing technology complements the antisense morpholino-based approaches for analysis of endogenous gene activities. The abundance of embryos (thousands of eggs from a single adult female in a day) also makes –omics research readily available, including RNA-seq, ChIP-seq, ATAC-seq, proteomics, and metabolomics studies. Today, the favored amphibian species has shifted from a range of frogs, salamanders, and newts to the African clawed frogs Xenopus laevis and Xenopus tropicalis, and the Mexican axolotl Ambystoma mexicanum, but the topics of research have diversified.
Some of the examples include studies of molecular nature of and signal tuning and crosstalks at the Spemann organizer, embryonic patterning along different axes, mechanical and signaling control of tissue morphogenesis both at early developmental stages (gastrulation and neurulation) and during organogenesis (e.g. heart formation, gut looping, ciliated skin cell development), wound healing, tissue regeneration, and epigenetic control of embryogenesis (including the Nobel prize winning experiments by John Gurdon that demonstrated nuclear equivalence and cellular reprogramming). In addition, the Xenopus model is increasingly used for studies of genetic variants that are associated rare human diseases. All these demonstrate that Xenopus continues to be a powerful system ripe for in depth investigations that lead to vital discoveries.
In the Xenopus module, students will be introduced to the more than 100 years of history of experimental embryology in amphibians and be given the opportunity to apply modern molecular and imaging technologies to the Xenopus embryos. Students will learn how embryos are generated via in vitro fertilizations, how to prepare and inject embryos, how to make fine surgical dissections to obtain various explants and perform tissue transplant experiments, and how to mount fixed or live samples for imaging. They will be provided with reagents to inactivate or alter gene expression and instructed to design and analyze their experiments. They will also be provided with embryos from another amphibian species, the Mexican axolotl, for comparative and cross-species experiments. This module will give students the necessary skills to use the frog to determine the function of a novel gene along with the ability to apply modern methods to uncover molecular and physical mechanisms that underlie the classic observations made by the famous embryologists of the past century.
Modern developmental biology has largely focused on models contained within two of the three main groups of animals: the deuterostomes and the ecdysozoans. The third group, known as the Spiralia (or Lophotrochozoa), contains nearly half of all animal phyla and a vast range of adult morphologies. Even among this group, cephalopod molluscs have a highly derived body plan and a suite of innovations with no obvious correlates in other molluscs, including a crown of sucker-lined arms, highly developed eyes, and a complex central nervous system, making them a compelling model to the study of the evolution of novelty. Many cephalopod innovations have been thought to be the result of competition with teleosts (Packard 1972), and therefore also serve as a model for the study of convergence.
Embryogenesis in cephalopods is a complete departure from the otherwise highly conserved lophotrochozoan developmental sequence of holoblastic spiral cleavage followed by larval stages. Instead, cephalopods have superficial cleavage on top of a large yolk followed by direct development. The initial cell divisions produce a blastodisc at the animal pole, which continues to divide, spreading over and eventually enveloping the yolk by epiboly. The profound differences in early cephalopod development relative to other molluscs represents both a true novelty and an additional example of convergence with teleost fish.
Particular traits in different cephalopod species have been leveraged as biological models. For example, the longfin inshore squid Doryteuthis pealeii has long been used as a system to study the neurophysiology of the giant axon, the cuttlefish Sepia officinalis has been a model to understand adaptive coloration, and neuroanatomy and behavior has been studied in Octopus vulgaris. Though cephalopods have been important scientific models, particularly in neurobiology, many modern techniques applied in other animal clades are undeveloped in this group. Recent work at the MBL has focused on developing one or more model cephalopod species, leveraging advances in sequencing, genome editing, and molecular techniques for the study of these remarkable animals. These efforts are focused on the Hawaiian bobtail Euprymna scolopes, the flamboyant cuttlefish Metasepia pfefferii, the stumpy cuttlefish Sepia bandensis, the pyjama squid Sepiolina lineolata, and the California two spot octopus, Octopus bimaculoides. In this module, students will have access to embryos of a variety of cephalopod embryos from those listed above. A number of antibodies and vital stains work well in these species, but students are encouraged to try new techniques to observe these animals, as many have yet to be explored.
Ctenophores (‘comb-bearers’) are a monophyletic group of globally distributed gelatinous, primarily pelagic, marine predators. The sea walnut, Mnemiopsis leidyi, is a lobate ctenophore widely distributed along coastal Atlantic environments and can be found during summer months in Eel Pond, the NOAA jetty and other nearby harbors. The gelatinous adults produce a blue-green bioluminescence when disturbed, are self-fertile hermaphrodites, and typically release thousands of eggs in a single spawning event. Experimental embryologists have a long history of using Mnemiopsis for cell lineage analyses due to easy access to large numbers of synchronized, highly transparent, rapidly developing embryos. These studies have been instrumental in describing cell fates associated with a ctenophore specific invariant early cell cleavage program. Moreover, the Mnemiopsis leidyi genome has been sequenced, facilitating the adoption of functional genomics approaches to this model system. A resurgent interest in ctenophore biology stems from recent phylogenomic analyses that have highlighted the early branching relationship of ctenophores relative to other animal phyla, making Mnemiopsis a particularly interesting and informative system for investigating character trait evolution during metazoan diversification.
The ctenophore module will introduce students to Mnemiopsis as a ctenophore model system and provide a number of reagents for cellular/molecular biology that can be used to explore ctenophore embryonic development. Gametes and live embryos at different embryonic stages will be available for DIC and fluorescent imaging with the wide range of microscopy resources available to the course. Also single cell embryos for lineage tracing and CRISPR/Cas9 genome editing via microinjection will be available to students in the course.
Molluscs—derived from the latin word for soft—make up the second most speciose animal phylum (after arthropods), and are bilaterally-symmetric, unsegmented members of the protostome (lophotrochozoan) superclade, the Spiralia. Molluscs have adapted to virtually every habitat on earth (save for the sky, as there are no flying forms), and exhibit a stunning array of morphological, physiological, and behavioral diversity. Members include the bivalves (oysters and mussels), chitons, gastropods (snails and limpets), and cephalopods (octopus, cuttlefish and squids). Although they form a monophyletic group, few synapomorphies define the entire group; one highly conserved feature is the mantle, an organ in the dorsal body wall that secretes biominerals in shelled forms. Molluscs have been employed for over 100 years in the study of developmental mechanisms, cell biology, population genetics, materials sciences, neurobiology, behavior, physiology, evolution, and aquaculture. So far, two Nobel prizes have been awarded for discoveries made using molluscs: the 1963 Nobel Prize in Physiology or Medicine for uncovering the ionic mechanism of action potentials using the giant squid axon, and the 2000 Nobel Prize for Physiology or Medicine for discoveries concerning signal transduction in the nervous system using the gastropod sea hare Aplysia. Recently, CRISPR/Cas9 has been successfully used to knock in a fluorescent transgenic reporter using the gastropod snail Crepidula, the first demonstration of this approach in any lophotrochozoan.
Some of the earliest developmental studies using molluscs were carried out in the late 19th century by E.G. Conklin working at the MBL with slipper snails, including Crepidula fornicata. Since Conklin’s seminal work, mollusc embryos have been used to study cell lineage, spiralian fate maps, asymmetric cell division, sub-cellular localization of mRNAs, organizer signaling, chirality at the levels of cells and organs, evolution of biomineralization, and the formation of the nervous system, to name a few. The spiral cleavage program is ancestral for molluscs (notably lost in the cephalopods) and provides a powerful framework for comparing developmental mechanisms among morphologically diverse molluscan species, and even between molluscs and other phyla like annelids and nemerteans that share this highly conserved mode of development (as members of the Spiralia/Lophotrochozoa).
In this module students will have access to a variety of molluscan embryos that can be used for cell lineage tracing, embryological manipulations, laser ablation, gene-perturbation, immunohistochemistry, and live imaging using fluorescently tagged bio-sensors. You will also have access to the larvae and adults of several closely-related species (such as the slipper snails) that exhibit different modes of development, including direct (lecithotrophic) versus indirect (planktotrophic) development. Students will also be able to study the formation and regeneration of the shell.
Sea urchins and Sea stars are Echinoderms and have long been used for experiments to understand how development works. These animals are found in abundance in marine environments around the world and there are several species of each that have been studied in some detail. The genomes of a number of Echinoderms have been sequenced and a number of cellular and molecular approaches are available for experimentation. Sea urchins and sea star embryos are model embryos for the laboratory for a number of reasons. They are easy to obtain, shedding gametes and fertilizing the embryos is simple, the embryos develop rapidly, the embryos are transparent so that many imaging methods can readily be used, and the embryos are especially valuable for the study of gene regulatory networks, for dynamics of morphogenesis, for cut and paste experiments, and for many different molecular perturbations to study function. Each of these approaches will be introduced and available for experimentation in the module.
The sea urchin and sea star module will introduce methods of animal husbandry to obtain gametes and embryos, and will introduce methods of egg injection, embryo microdissection, cell recombination, morpholino and CRISPR perturbations, antibody staining, and a number of reagents will be available for experimentation. In each case the embryos will then be available for time lapse, confocal, light sheet, immunofluorescence, DIC, and other forms of microscopy in each case providing pairing a highly photogenic embryo with the rich microscope resources available for student use in the course. Students will be encouraged to think of their own hypotheses to test and the instructors and TAs will also provide suggested avenues to explore.
C. elegans, a free-living nematode, was chosen by Sydney Brenner in 1963 as an experimental organism because it is transparent, easy to propagate, and is a hermaphrodite capable of selfing as well as outcrossing, which facilitates genetic manipulation. The entire cell lineage of C. elegans, from the zygote to the adult has been determined—hermaphrodite adults contain exactly 959 somatic cells. The worm’s transparency and reproducible anatomy make it possible to identify each cell at all stages of development. These features, coupled with its amenability to forward genetics, reverse genetics, systems-level approaches, and molecular studies, have allowed developmental, physiological, and behavioral events to be identified and characterized. Work on this small worm has led to Nobel Prizes for Physiology or Medicine in 2002 to Sydney Brenner, Robert Horvitz, and John Sulston for their characterization of organ development and programmed cell death, the 2005 award to Andrew Fire and Craig Mello for their characterization of RNAi, and the 2008 Nobel Prize in Chemistry to Martin Chalfie and colleagues (including Osamu Shimomura from Woods Hole!) for their use of the jellyfish green fluorescent protein (GFP) as an experimental reagent.
C. elegans has two sexes: self-fertile hermaphrodites (which produce both sperm and oocytes) and males that can inseminate hermaphrodites. Hermaphrodites can produce large populations without mating; you can generate many worms by placing a single hermaphrodite on a petri dish with sufficient food (worms are grown on a lawn of E. coli). Genetic crosses are performed by mating males with hermaphrodites, with male sperm precedence increasing the likelihood of producing cross progeny over self progeny.
In this module, students will learn how to knock down genes with RNAi, look at protein dynamics with fluorescence recovery after photobleaching (FRAP), conduct experiments with optogenetic approaches, carry out laser ablation of selected cells, conduct analysis of differentiated cell types by fluorescent protein markers, and observe development and behavior in wild-type, mutant, and wild-caught worms. One of the main advantages of C. elegans for genetic studies is its short generation time (3 days). Although the constraints of a short module will not allow sufficient time to perform genetic experiments, you will have the opportunity to take advantage of many reagents and experimental conditions to live image worm development.
Ascidians are the largest group within the tunicates. Tunicates are invertebrate chordates and are the sister group to vertebrates. Ascidians are marine animals and they have a long history as an experimental model system for development. Edwin Grant Conklin followed the cell lineage of the ascidian Stylela canopus during studies at the Marine Biological Laboratory and published his results in a 1905 monograph. The embryos of Styela have naturally pigmented cytoplasms that are segregated into specific cell lineages during development. Today, most laboratories that study ascidian development have focused on a few model species including Ciona robusta, C. savignyi, Halocythia roretzi and Botryllus schlosseri. The genomes of about a dozen species have been, or are currently being, sequenced. Many of the experimental tools utilized in other systems, CRISPR/Cas-9, RNA-SEQ, and gene knock-downs have been adapted to ascidians. One of the significant advantages of ascidians is the ability to generate transgenic embryos using a simple electroporation process. The ability to manipulate gene function in these animals makes them well-suited for studying the molecular mechanisms of development, including the characterization of gene regulatory networks that govern tissue-specific patterns of cell differentiation.
With rare exceptions, tunicates are usually self-sterile hermaphrodites and fertilization occurs between the mixing of gametes from two or more individuals. As tunicates are marine animals, they are broadcast spawners with gametes being freely shed into sea water. Many species of ascidians have been spread throughout temperate ocean waters in large part because the adults can grow on the bottoms of cargo ships. When in port, adults attached to the ships release gametes and new generations of animals can gain footholds in the new environments. In some species, the block to self-fertilization is less restrictive, so even small numbers of adults can initiate new populations of animals.
In this module, students will learn about different species of ascidians, how to spawn the animals and how to generate transgenic embryos. Additionally, students will learn more classically-oriented experiments including the dissection of early embryos into specific sets of cells. Students will have an opportunity to examine cells lineages by using a variety of fluorescent protein transgene constructs and mis-express transcription factors and signaling proteins to alter cell fate. Lastly, students will be able to use the CRISPR/Cas9 system to both knock-out genes of interest and to knock genes into the genome.
Acoel worms belong to a lineage (Phylum: Acoelomorpha) that is likely the sister group to all other animals with bilateral symmetry (bilaterians). Acoels are “wormy”, with clear anterior-posterior and dorsal-ventral axes; they have organized subepidermal nervous systems, but lack eyes, excretory organs, and a true gut. Acoels diverged from other bilaterians 550 million years ago, a node on the animal tree that corresponds with the origins of many features, e.g., bilateral symmetry, a centralized nervous system, and mesoderm (the third germ layer) and true muscle. Therefore, the study of acoels is important for many questions about the evolution of development during early animal evolution.
The acoel Hofstenia miamia, commonly known as the three-banded panther worm, is a new laboratory model for molecular studies of development and regeneration. The animals can be maintained as a sexually reproducing population, producing juvenile worms in substantial numbers to enable large-scale experiments. mRNA and protein localization can be studied via whole-mount in situ hybridization and immunostaining respectively. Hofstenia are amenable to studies of gene function via RNA interference (RNAi) upon soaking or injecting the animals with dsRNA. Genomic resources are also available – the transcriptome can be downloaded from NCBI, and a genome assembly is currently being annotated.
Hofstenia reproduce sexually, producing hundreds of embryos in a single a day. Hofstenia embryos undergo a stereotype cleavage program called “duet cleavage”. Hofstenia embryos are readily accessible and amenable to experimental procedures such as in situ hybridization and manipulations such as microinjection of dyes, mRNAs, and guide RNAs with Cas9 enzyme.
Regeneration in Hofstenia is similar to regeneration in planarians – both species generate anterior and posterior missing tissues when cut transversely. Acoels utilize similar patterning mechanisms to specify the identity of new tissue along the anterior-posterior (Wnt signaling) and dorsal-ventral axes (Bmp-Admp signaling) as planarians. Furthermore, Hofstenia has a parenchymal population of proliferating cells that are required for regeneration and express piwi, reminiscent of the neoblasts in planarians. Given that acoels and planarians diverged from the last common bilaterian ancestor 550 million years ago, these findings suggest that the genetic mechanisms of regeneration (including those for pluripotent stem cells) may be fundamental features of animal biology that were lost in vertebrates.
Studies in Hofstenia will complement and expand on many of the discoveries made in well- established model systems, and will inform us on the evolution of regenerative and developmetal mechanisms over animal evolution.
Activities in this module include, examining the anatomy of Hofstenia, observing regeneration, observing embryos, live imaging of embryos, whole mount antibody & phalloidin staining of animals, grafting animals together.
Annelids are the phylum of segmented worms, with over 17,000 described species in this clade. Annelids are characterized by their body plan, centralized nervous system, continued adult growth from addition of segments from a posterior growth zone, and adult regenerative abilities. Embryologists observed their development over a hundred years ago and annelids have been used for evo-devo studies due to their superficial similarities to other segmented animals (e.g. vertebrates and arthropods). Some annelids live in fresh water or are even terrestrial (e.g. leeches and earthworms), but most species of annelids live and reproduce in the marine environment, and many have an indirect life cycle, producing a free-swimming larval form called the trochophore. Two species that are emphasized at the MBL are Capitella teleta and Chaetopterus sp. Capitella is a small benthic worm with a cosmopolitan distribution and a high quality publically available sequenced genome (the first annelid to be sequenced!). There are distinct male, female, and hermaphrodite sexes. Chaetopterus is the parchment tube worm, has a highly tagmatized body plan, and is known for being able to regenerate both its head and tail. Recent molecular phylogenomic studies also place it as one of the earliest branching members of the Annelida.
Annelids exhibit spiral development, which means that there is a highly stereotypic program of early cleavages. The cell lineage of several annelids has been described in detail, and every cell in the early stage embryo of Capitella and Chaetopterus can be uniquely identified. In addition, the fates of these cells are known and a comprehensive fate map that identifies the early embryonic origin of larval features is available for Capitella. Because this pattern of early development is highly conserved, homologous cells can be compared across species, and even other animals such as molluscs. These observations provide interesting EvoDevo-type questions as to how such a highly conserved early cleavage program gives rise to such a highly diverse collection of adult body plans.
In this module students will be exposed to some of the diversity within annelids, both larval and adult forms. Students will learn animal handling techniques, how to obtain embryos from Capitella and how to set up an in vitro fertilization for the parchment tube worm, Chaetopterus. Students will have the opportunity to observe normal early development, and use immunohistochemical and staining methods to observe the developing larval nervous system and musculature across diverse larval forms. These types of studies are essential before any understanding of how ontological differences arise during annelid evolution. Students can also perform classical embryological manipulations, laser deletions of individual blastomeres, pharmacological perturbations, microinjections, make time lapse movies, or a wide range of other experiments.
In this module, students will learn about a variety of arthropod systems, including the model genetic system, Drosophila melanogaster. Most importantly, students will leave with the ability to analyze and compare the development of different arthropod embryos. In order to do that, students will be performing various molecular and embryological techniques, such as antibody staining, in situ hybridization, live imaging, and lineage tracing using both classical injection and modern genetic methods. Students will also analyze mutant phenotypes as well as try to create their own mutants using CRISPR/Cas9
At first, students will use a set of antibodies to detect the expression of important developmental proteins and RNAs in the fruit fly Drosophila melanogaster. This will allow students to master the procedure of antibody staining, examine the spatiotemporal expression of these proteins, and study the development of various tissues. Because many of these proteins are well conserved in all arthropods studied to date (and often in other phyla as well), a subset of these antibodies will allow students to stain all sorts of arthropods that either instructors provide or by collecting from around Woods Hole.
In this module students will also have the opportunity to look at aspects of post-embryonic development. In particular, students will look at wing imaginal disc development in Drosophila and scale patterning in butterflies. Students can stain imaginal disks with available antibodies and compare the expression pattern of various proteins between flies and butterflies.
Instructors will also give students several ideas for “projects” that will be discussed, but will also encourage students to try out as many techniques as possible on a range of species.