Developmental Biology: A Guide for Experimental Study, Second edition

table of

Contents

Preface x 1. Getting Started Things you will need 1-1 Your laboratory notebook 1-1 A format for your notebook 1-3 Formal laboratory reports 1-3 Anatomy of a scientific paper 1-3 Some hints about writing 1-4 Using the library 1-5 Accompanying materials 1-6 Selected bibliography 1-7

2. Embryological Tools Microknives 2-1 Microneedles 2-2 Hairloops 2-2 Pipettes 2-3 Embryo spoons 2-4 Instrument tray 2-4 Retooling metal instruments 2-5 Sterile technique 2-6 The workbench 2-6 Dissecting tools 2-6 Glassware 2-7 Fluids 2-7 Rules for the road 2-8 Accompanying materials 2-8 Selected bibliography 2-8 Suppliers 2-8 Glossary 2-10 Pre-lab questions 2-11

Contents 3. Using the Compound Microscope The microscope 3-1 Safety first 3-2 Stage 3-2 Magnifications 3-2 Oculars 3-2 Objectives 3-3 Koehler illumination 3-4 The condenser 3-4 Iris diaphragm 3-5 Other settings when maximum resolution is not desired 3-5 Oil immersion 3-5 Make your $1000 microscope into a $10,000 instrument for pennies 3-6 Color filters 3-6 Polarizing filters 3-7 Dark-field optics 3-8 Measurement under the microscope 3-8 Care and maintenance of your microscope: “Twelve good rules” 3-9 Accompanying materials 3-9 Selected bibliography 3-10 Suppliers 3-10 Glossary 3-11 Pre-lab questions 3-14

4. Cellular Slime Molds Life cycle 4-1 The vegetative stage 4-1 Aggregation 4-1 Pseudoplasmodium 4-3 Culmination 4-4 Sexual reproduction: A rare event 4-4 Preparing for your laboratory studies 4-4 Culture procedures 4-5 Experiments using Dictyostelium 4-6 Labeling with vital dyes 4-6 Transecting, grafting, and disaggregating pseudoplasmodia 4-7 Disruption of cyclic AMP levels 4-8 Behavioral studies 4-9 Completing your experiments 4-10 Accompanying materials 4-10

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Contents Selected bibliography 4-10 Suppliers 4-12 Glossa ry 4-13 Pre-lab questions 4-17

5. Gametogenesis Meiosis: An outline 5-1 Mammalian spermatogenesis 5-2 Stereology 5-4 Tunica albuginea 5-4 Seminiferous tubules 5-4 Spermiogenesis 5-5 Interstitial cells 5-6 Comparative sperm morphology 5-7 Mammalian oogenesis 5-7 Follicles 5-9 Theca 5-9 Oocyte 5-9 Corpus luteum 5-11 Histological Hitchcocking 5-11 Accompanying materials 5-11 Selected bibliography 5-12 Suppliers 5-12 Glossary 5-13 Pre-lab questions 5-18

6. Echinoid Fertilization and Development Collecting 6-1 Fertilization 6-3 Instructions for normal fertilization 6-4 Watching fertilization 6-5 Interfering with fertilization 6-5 Parthenogenesis 6-6 Cleavage, gastrulation, and larval stages 6-8 Accompanying materials 6-9 Selected bibliography 6-9 Suppliers 6-11 Glossary 6-12 Pre-lab questions 6-17

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Contents 7. Sea Urchin Development—Effects of Ultraviolet Radiation Ultraviolet radiation in Earth’s atmosphere 7-1 Biological effects of UV radiation 7-2 Mechanisms that protect against damage from UV 7-3 Using sea urchins as a model organism for UV studies 7-3 Setting up your experiments 7-4 Special lightbulbs that can be used 7-5 Filters that can be used to exclude UV-A or UV-B 7-5 Filters that block UV-B 7-6 Filters that block both UV-A and UV-B (UV opaque) 7-6 Filters that do not block either UV-A or UV-B (UV transparent) 7-7 Safety first 7-7 Monitoring your cultures 7-7 Keeping up with the issues 7-8 Accompanying materials 7-8 Selected bibliography 7-8 Suppliers 7-10 Glossary 7-11 Pre-lab questions 7-15

8. Development of the Fruit Fly Life cycle 8-1 Culturing Drosophila melanogaster 8-2 Collecting eggs 8-3 Collecting chamber 8-3 Mating behavior of adult flies 8-3 Observations of the egg 8-4 The chorion 8-4 Dechorionating an egg 8-5 Embryogenesis 8-6 Cleavage 8-6 Gastrulation 8-6 Later development 8-8 Embryonic staging series 8-8 Larval development 8-9 Anatomy of the larva 8-10 External anatomy 8-11 Internal anatomy 8-11 Development of imaginal discs 8-12 Dissection of imaginal discs 8-14

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Contents Eversion of imaginal discs 8-15 Whole-mount preparations of imaginal discs 8-15 Chromosome squash from salivary glands 8-16 Pupation 8-16 Accompanying materials 8-18 Selected bibliography 8-18 Suppliers 8-19 Glossary 8-20 Pre-lab questions 8-28

9. Early Development of the Chick The chick egg 9-1 The female chick reproductive tract 9-4 Cleavage 9-6 Gastrulation 9-7 24-Hour chick whole mount 9-9 Accompanying materials 9-12 Selected bibliography 9-12 Suppliers 9-13 Glossary 9-13 Pre-lab questions 9-20

10. 33-Hour Chick Embryo Central nervous system 10-1 Circulatory system 10-2 33-Hour whole mount 10-2 Folds and tucks: Morphogenesis proceeds 10-2 Ectodermal structures 10-3 A category unto themselves: Color them green 10-4 Endodermal structures 10-5 Mesodermal structures 10-5 33-Hour serial sections 10-8 Experiment with evolution 10-13 Create a puppet 10-13 Accompanying materials 10-13 Selected bibliography 10-14 Suppliers 10-15 Glossary 10-15 Pre-lab questions 10-21

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Contents 11. The Living Embryo and Making of Whole Mounts Incubation of eggs 11-1 Preparing whole mounts 11-2 Fixation 11-5 Washing 11-6 Staining 11-6 Dehydration and clearing 11-7 Mounting 11-7 Accompanying materials 11-8 Selected bibliography 11-8 Suppliers 11-8 Glossary 11-9 Pre-lab questions 11-12

12. Histological Techniques Fixation 12-1 Washing 12-3 Dehydration and clearing 12-3 Special microwaving techniques that save time 12-4 Paraffin infiltration 12-4 Paraffin molds 12-5 Paraffin embedding 12-6 Mounting and trimming paraffin blocks 12-6 Sectioning 12-7 Mounting sections 12-8 Staining 12-10 Using the hematoxylin, eosin, and alcian blue staining series 12-11 Hematoxylin 12-12 Mounting coverslips 12-12 Accompanying materials 12-13 Selected bibliography 12-13 Suppliers 12-14 Glossary 12-14 Pre-lab questions 12-17

13. Planarian Regeneration Collecting and maintaining planarians 13-1 Observations of normal anatomy and behavior 13-2 Anatomy 13-2 Behavior 13-4 The regeneration process: Patterns and theories of regeneration 13-5

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Contents The regeneration process 13-5 Pattern formation and positional information 13-6 Diffusion gradient model 13-6 Polar coordinate model 13-8 Patterns of regeneration 13-9 Experimental procedures 13-10 Planning your experiments 13-10 Procedures 13-12 Record keeping and analysis 13-13 Accompanying materials 13-13 Selected bibliography 13-13 Suppliers 13-15 Glossary 13-16 Pre-lab questions 13-19

14. Amphibian Development Breeding 14-1 Environmental hazards affecting amphibian development 14-2 Preparing for the field trip 14-3 Learn about your local amphibians first 14-3 Equipment needed and recording of data 14-6 Equipment for pH measurements 14-6 Come dressed for the field 14-7 Know government regulations 14-7 The field trip 14-7 Rules for the road 14-9 Night sounds 14-9 Back at the laboratory 14-11 Species identification 14-11 Normal development 14-11 Environmental effects on amphibian development 14-16 Adopt an egg mass 14-18 Stay in touch 14-18 Accompanying materials 14-18 Selected bibliography 14-19 Suppliers 14-21 Glossary 14-22 Pre-lab questions 14-25

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Contents 15. Zebrafish Development A new model organism 15-1 Fertilization 15-2 Cleavage 15-5 Blastula 15-5 Gastrulation 15-7 Organogenesis 15-8 Preparing for your laboratory studies 15-9 Culture procedures 15-10 Tanks and water 15-10 Sexing adult fish 15-10 Feeding 15-10 Obtaining embryos 15-10 Maintaining embryos 15-11 Raising larvae 15-12 Experiments using zebrafish embryos 15-12 Observing normal development 15-12 Removing chorion 15-13 Temperature experiments 15-13 Chemical teratogens 15-14 Lithium 15-14 Vitamin A 15-15 Alcohol 15-17 Endocrine disruptors 15-17 Testing water below the pipe 15-18 Behavioral studies 15-18 Completing your experiments 15-18 Preparation of whole mounts 15-19 Fixation 15-19 Washing 15-20 Staining procedures 15-20 Clearing and mounting 15-20 Accompanying materials 15-21 Selected bibliography 15-21 Suppliers 15-24 Glossary 15-25 Pre-lab questions 15-36

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Contents Web Links to Appendices Web pages for the appendices below can be found at www.developmentalbiology.net The hyperlinks below go directly to each of the individual pages.

Appendix 1: List of Recipes Appendix 2: List of Suppliers and Addresses Appendix 3: Full Glossary Appendix 4: Laboratory Safety

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Preface

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Preface Development is a magnificent and mysterious journey that we all participate in until the cold ground claims us. It is a journey so complex that we travel mostly unaware of the means by which we are speeding along. But the means of travel are well worth our attention. This book invites you to look through the windows that science provides—you may be the one who sees what others have missed.

The Book at a Glance This guidebook begins with a discussion of the tools you need for studying developing organisms and moves on through the events of development, first in simple and then in more complex organisms. Chapters 1–3 and 12: Tools of the trade Much of the study of developmental biology involves craft. The first three chapters and chapter 12 will school you in the methods of this craft, teaching you how to record and report data, how to create your own tools, how to be master of your microscope, and how to prepare tissue for microscopic study. Chapter 4: A simple organism Not all organisms use sexual reproduction as the preferred method for passing on their genes. Sex is expensive, burdened as it is with the costs of meiosis. The frugal cellular slime mold, consisting of only a few cell types, seldom indulges in sex, instead undergoing repeated cycles of asexual reproduction. Chapter 4 shows you how to design experiments that use this organism to explore principles of development operating in the realm of simple organization. Chapter 5: Gametogenesis In organisms that do use sexual reproduction, gametogenesis marks the beginning of development. Chapter 5 shows you how to sleuth your way through the events that lead to the formation of dimorphic gametes—eggs and sperm, the cells that each carry half a load of genetic material for the new organism. Chapter 6: Fertilization Mature gametes have their cellular triggers cocked; fertilization is the explosion that is set off when egg and sperm meet, fusing the two gametes and propelling the new individual off along the road of development. Chapter 6 invites you to explore questions surrounding the mechanisms of this event using sea urchin gametes. Chapters 6 –11: Cleavage, gastrulation, and organogenesis The journey started at fertilization continues at breakneck speed through cleavage. Cleavage provides the embryo with a large number of cells for continuing the trip. Its product, a beautiful blastula, may be the hollow ball of cells you will see among the sea urchin embryos spiraling across your dish in the experiments in Chapters 6 and 7; a ring of cells surrounding a yolk mass, as you will see in fruit fly embryos in Chapter 8; or a plate of cells balanced on a yolky sphere, as you will see in chick embryos in Chapter 9.

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Following cleavage, cells start using a new set of instructions, newly transcribed messenger RNAs (mRNAs), for their developmental journey. Some of these messages instruct the cells to don new clothes, in the form of new cell surface molecules. These molecules identify the cells according to germ layer: ectoderm, mesoderm, or endoderm. Once they put on their new clothes, the cells can’t sit still. They begin moving to new positions in the process of gastrulation, which brings the germ layers into positions appropriate for their specific fates. You will see how these movements are choreographed in the sea urchin, fruit fly, and chick in Chapters 6 through 10. With the germ layers in position, the trip enters a phase of dynamic interactions known as organogenesis. Cells in the embryo relay information back and forth, and these interactions lead to organ differentiation. The embryo is now complete; the embryonic portion of the trip is over. In Chapters 6 through 11 you will glimpse this final leg of the embryonic journey as you raise sea urchin larvae, dissect and experiment with imaginal discs of fruit fly larvae, and test the responses of one of the earliest products of organogenesis in the chick, the embryonic heart. Chapter 13: Regeneration and pattern formation Development is a long journey that extends well beyond the event of hatching or birth. One of the fascinating side roads is that of regeneration. Few organisms are capable of such impressive feats of regeneration as those seen in planarians. Forming complete replicas of the intact worm from fragments as small as 1/279th the size of the original organism, the planarian, as you will see in Chapter 13, allows you to venture into the mysteries of regeneration. Chapters 7, 14, and 15: Development and the environment Development is always influenced by the environment in which it takes place, and often environmental hazards affecting the delicate embryonic stages of an organism threaten its survival. Organisms such as sea urchins and amphibians, which must entrust their poorly protected embryos to the open wilds, can be particularly vulnerable, and therefore make sensitive barometers of environmental hazards. In Chapter 7, the hazards of ultraviolet radiation, an ever increasing threat, are examined using the delicate sea urchin embryo. In Chapter 14 we explore the woods and streams for evidence of threats to embryos of our cool and thin-skinned amphibian relatives. And in Chapter 15, we end our explorations with a study of hazards that we expose ourselves to in our daily lives and determine whether they affect the development of the zebrafish, with the premise that if they harm zebrafish development, then they also pose a threat to the development of other species, including humans. One goal of this book is to help students be independent scientists. All the exercises are described in sufficient detail that students should be able to perform them on their own. No secrets have been sequestered to a separate instructor’s manual. In addition to instructions and background information, the book provides recipes for solutions, bibliographies for further study, and lists of scientific suppliers. The illustrations in the book are line drawings designed to clearly delineate components and to draw attention to general principles of structure. Photographs and videos for all chapters can be found in the corresponding Vade Mecum modules of the CD-ROM.

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Vade Mecum Vade Mecum: An Interactive Guide to Developmental Biology, by Mary S. Tyler and Ronald N. Kozlowski, and published by Sinauer Associates, was developed primarily to augment this laboratory manual. The goal of including the manual with Vade Mecum on a single CD is to provide students with all that they need to be truly independent learners. Each chapter of the laboratory manual is represented by a corresponding Vade Mecum module, including over 150 videos and 300 labeled photographs that explain the development of the organisms and give step-by-step explanations of techniques. For example, where useful, color-coding is superimposed on living embryos to illustrate positioning of different germ layers. A complete set of cross sections of a 33-hour chick embryo and whole mounts with definitions of terms are included. A “virtual microscope” section shows how to achieve Koehler and dark-field illumination and how to use polarizing filters. For each chapter, there are study questions and websites listed. A section on laboratory safety is also included. Vade Mecum can be purchased as a stand-alone item, or bundled with the printed version of the laboratory manual at reduced cost, and also comes packaged with Scott Gilbert’s Developmental Biology, Sixth and Seventh Editions.

Acknowledgments Many people have contributed ideas, inspiration, and hard work to these pages, and to them I am very grateful. I thank all the reviewers who added their wisdom and careful comments: Nikki Adams, John Dearborn, Brian Hall, Malcolm Hunter, Richard Kessin, John Lucchesi, Robert Mead, David McClay, Drew Noden, John Ringo, Brian Sullivan, Seth Tyler, Bonnie Wood, Joel D’Angelo and Scott Gilbert. To those at Sinauer Associates, publisher Andrew Sinauer, production specialist Chris Small, and editors Jason Dirks, Kathaleen Emerson, Mara Silver, Chelsea Holabird, Marie Scavotto, and Carol Wigg, I extend a special warm thanks for their work. Their creativity, attention to detail, and support were invaluable. And to my always-curious students, I am beholden for their attentiveness, which has taught me to see more than I would alone. Mary S. Tyler Orono, Maine

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chapter

Getting Started

As you embark on your exploration of developmental biology, the most important thing to bring with you is curiosity. Ask questions, for these will lead to hypotheses, and hypotheses open the mind to discoveries. Keep careful records of your questions and answers. Above all, enjoy exploring.

Things You Will Need The list of things you will need to bring is short. Laboratory notebook A standard 8.5”×11” looseleaf notebook is the most useful. The pages are large, and you can add pages where you need them. Drawing paper Any thickness, white, unlined, 8.5”×11” paper. Most experiments will require drawings to record data. If you erase a lot or simply enjoy putting pencil to paper, you might consider having the more expensive heavyweight paper that holds up under many erasures. Colored pencils Red, yellow, blue, and green. When studying germ layers, you will be color-coding your diagrams according to the standard embryological code: red = mesoderm, yellow = endoderm, and blue = ectoderm. Green often is used to designate neural crest. Dissecting kit Blunt probe, dissecting needle, scalpel, and scissors. For the most part, you can use microdissecting tools that you make yourself. However, a dissecting kit will be useful at times.

Your Laboratory Notebook Everything pertaining to the laboratory is to be kept in your laboratory notebook: notes, drawings, data, speculations, thoughts in the middle of the night. This is a working notebook of the type every experimenter keeps. It is not necessary for your records to look like a polished, neat report (unless, of course, this is your natural style). Your notebook must be orderly and complete, however. Since the quality of your science can be seen from the quality of your laboratory notebook, your laboratory instructor may check yours from time to time. To help make the quality of your science the very highest, here are some important hints. Always write in pencil Laboratory spills can wipe out whole pages of data written in pen. Always take careful, complete notes on all observations Minutiae often lead to great discoveries. Record with descriptions, measurements, and drawings wherever possible. Diagram to scale, noting the scale used. Drawings need not be artistic, but they do need to be accurate. If you can see it, you can draw it. If you can’t draw it, you’re not seeing it. Just look harder.

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Never recopy notes Valuable information can be lost. Remember, this is a working notebook. Follow a specific format so your notes are orderly, and make sure you’ve written neatly enough so that you can read it. It’s your notebook—not the instructor’s or anyone else’s.

ALT-1 Date, Time TITLE OF STUDY General procedures This section should describe the general experimental procedures used. In many cases, these will be t h e procedures given in a laboratory manual. You do not need to recopy these, but you do need to indicate any changes or additions you make. Experimental setup This section describes the specific procedures for each individual experiment within the study. You number each experiment and state exactly what you did in setting up this experiment. You also state what you are testing: What questions are you asking of the organism? Observations and results This section is a careful record of everything you do and observe during the course of the study. Every time you make a set of observations, you start by dating the page, and then recording your observations using the code you invented for each experiment. Observe, measure, and draw. Don’t crowd information. Leave space between your notes for one experiment and those for the next. Remember, these are notes. They need not be in complete sentences, but they should be extremely precise and clear. These are your data—the most important part of a l l your work. Hypotheses and conclusions can be wrong; data must stand the test of time. Summary of results When the experiments are wrapped up, the excitement builds. You can now collate your data to figure out what they’re telling you. Assemble the data into graphs and charts; look for trends and timing of events; determine how the data from one experiment compare with those from the others. Milk the data. And remember, this section is fact—it is a summarization of everything your organism has told you. Conclusions Here th e fun expl odes to party l evel s. Wh ere before you were adh eri ng to just-th e-facts-Mac, you now can expand i nto conjecture. Deci de wh at th ose facts coul d mean. Specul ate, th eori ze. Use wh atever background i nforma ti on you h a ve from courses, textbooks, l a bora tory ma nua l s, or oth er l i tera ture. Th i s secti on does not h a ve to pa ss th e test of ti me—you’re a l l owed to be wrong. Th i nk h a rd a nd h a ve fun.

Figure 1.1 A typical format for recording data in your laboratory notebook.

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A format for your notebook Any research scientist establishes a format for recording data. The one shown in Figure 1.1 is typical. You will need to invent a coding system for numbering your studies and each experiment within a study. In a classroom full of scientists, it is convenient to use your initials as part of the code. For example, if your initials are A.L.T., ALT-1 could refer to your first study. ALT-1-1 then could refer to the first experiment in your study, ALT-1-2 to the second, and so on.

Formal Laboratory Reports A formal laboratory report is no more and no less than a scientific paper in a format for publication. Be impressed, and know that you can produce work worthy of publication. In order to publish, though, one must repeat experiments a number of times, and you probably won’t have the luxury of this in the confines of a one-semester course. Your laboratory reports, therefore, will be a training ground for the scientific papers many of you will publish in the future (perhaps sooner than you think). Undergraduate students are perfectly capable of producing good, publishable papers, just as seasoned scientists are perfectly capable of producing bad ones. The secret is to practice the former and be able to recognize the latter. Presented here is a format for a formal scientific paper. Whether you use the entire format or a truncated form of it, you should at least know the anatomy of a scientific publication. Each section has a specific purpose. If you understand that purpose and adhere to it, your chances of producing excellent papers will increase. Anatomy of a scientific paper Abstract This is a concise rendering of the question you posed, the methods you used to investigate it, your results, and your conclusions. Usually only 100–200 words, it is almost always composed after the rest of the paper is written, even though it is placed first. Introduction This is a statement of the problem you are investigating and its significance, with enough background information to understand the next two sections of the paper. It is not a long section; most of the background information that needs to be included in the paper can be saved for the Discussion. Your main objective in the introduction is to get the readers to understand what you were asking and why you bothered to ask it. You want, very concisely and effectively, to hit them between the eyes with the meaning of your work, showing them how it fits into the bigger scheme of things. If you don’t succeed here, they may not bother to read further. Materials and methods This is the boring section. It is a very careful explanation of your methods, the precise materials you used, and where you got those materials. List species names, collecting sites, and suppliers of reagents, along with an explanation of the experimental design for each experiment. This allows you and others to repeat your experiments. Results This is a just-the-facts section. It is the section that should remain when all others burn in the fires of future truths and hindsight. It is created from the “Summary of Results” section from your laboratory notebook as described above. Data are presented in digested form, summarized in charts, graphs, and descriptions. You do not conjecture as to what these data are telling you—you simply present them.

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Discussion This is where you can let go and wax eloquent. A discussion is usually divided into two parts: first a statement of your conclusions (the “Conclusions” section from your laboratory notebook), then a discussion of the significance of these conclusions as they relate to other studies published in the literature. You must state how your results corroborate or disagree with previous work. In a paper for publication, you need to have done an exhaustive literature search before writing this section. (For a student laboratory report, your instructor should give you guidelines on the number of papers to be considered in your Discussion.) When you cite another person’s work, you need to write in parentheses the names of the authors and date of their publication. Readers then can find the full citation of this work in your Literature Cited section. You want to leave the reader with a full understanding of the importance of your results and their implications as they relate to bigger questions. Literature cited This is your bibliography. It is a listing of all the literature you cited throughout your paper. The format for this is rigid, and each journal will have its own rules. Whatever format you use, you must be consistent. An example of one acceptable format is given below. Journal names are abbreviated (usually using the abbreviation the journal itself uses or that used by Biological Abstracts, which has a listing called Serial Sources of the Biosis Data Base that is published each year). Following the journal name comes the volume number, then the page numbers of the article. Notice that the titles of books and journals, as well as species names and the names of genes, are usually italicized or underlined. You can find more examples of citations in the Selected Bibliography section at the end of each chapter in this book. Journal article: Smith, J. S. and J. P. Jones 1999. The significance of nothing. J. Biol. Nonsense 14: 13–17. Article from a book: Watler, J. N. 1999. Drosophila without per still uses clock. In The Anatomy of a Fly, M. I. Fruit and R. U. Fly (eds.). Unbelievably Esoteric Press, New York, pp. 78–91. Book: Tyler, M. S. 2000. I Didn’t Say That. Ununited Professors Press, Wabash, CT.

Some Hints about Writing Writing often does not come easily to scientists, and I’m convinced this is primarily the result of beating ourselves out of our natural styles. When you are caught in the grip of unintelligible prose, I suggest you apply these few rules to the afflicted areas. The cure is often immediate. Don’t pull teeth Writing should never be painful. If you are not having fun, the writing is probably not very good. Relax. Don’t be an editor as you write. Edit later. Enjoy being yourself on paper. Hit, don’t be hit The most common fault that makes writing dense and indecipherable is the use of the passive rather than the active voice. Instead of saying “the egg was fertilized by the sperm,” say “the sperm fertilized the egg.” In more complex sentences, this invariably clears away clouds of confusion. Accent the important All sentences have a natural stress point. Make sure the important information in your sentence is at this stress point, usually at the end of the sentence. Keep the train cars connected Sentences of a paragraph, like the cars of a train, must be connected. This is often done by making the subject of one sentence the subject of the next, or by placing old

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information before new in a sentence. Whatever the technique, one sentence must lead naturally to the next. Read good writers You learn by example. Choose writing that is effective and dissect it. Ask, What did the writer do that makes it good? Read about writing There are a number of books and articles about writing, some of which are listed in the Selected Bibliography at the end of this chapter. (If you devote no more than an hour to this step, read the 1990 article, “The Science of Scientific Writing” by Gopen and Swan. Its examples of obtuse writing will make it obvious that a great deal of scientific writing is bad writing. You can do better.)

Using the Library The library is a source of buried treasure, so bring the correct digging tools. Find out what computerized databases are available. Your library’s website is a good place to start. Some of the databases that will be of use are Expanded Academic Index, Medline, UnCover, and WorldCat. Look up websites for specific journals; often they allow you to conduct electronic searches of their articles. Abstracts and sometimes the entire text of articles may be available at these sites. A few examples include BioEssays (www.interscience.wiley.com), Developmental Biology (www.sciencedirect.com), FASEB Journal (www.fasebj.org), Nature (www.nature.com), Science (www.sciencemag.org), Scientific American (www.sciam.com), and Science News (www.sciencenews.org). Electronic databases are exceedingly useful, but realize they are only one method used in compiling a reading list on a subject. The most important first article you can get is a recent review article about the subject. Use its bibliography to make your first short list of papers to read. Also, learn to use scientific indices; some of the useful ones are listed below. Biological Abstracts and Science Citation Index are two that are available in electronic form over the web—your library may subscribe to them. These indices always include instructions for their use, but they still may be confusing. Ask a librarian to help—librarians consider this part of their job, so don’t be shy. Often they have instruction sheets they can give you. And finally, browse the current literature. The current issues of journals are usually displayed in a separate place from the bound back issues. Know which journals deal with your subject and get used to browsing through them. Included below are lists of specialized developmental biology journals and the more general journals that are important to developmental biologists. (These are only short lists; many excellent, important journals have been left off.) If you scan only ten of these on a weekly basis, you quickly will be learning the art of being a scientist. Scientific indices Biological Abstracts Biology Digest Science Citation Index Zoological Record Journals specific to developmental biology BioEssays Cell Differentiation Development (formerly Journal of Embryology and Experimental Morphology) Developmental Biology

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Development, Growth, and Differentiation Differentiation Roux’s Archives of Developmental Biology (alternate title Wilhelm Roux’ Archiv; formerly Wilhelm Roux’ Archiv für Entwicklungsmechanik der Organismen Journals that cover development and other topics American Zoologist Cell Experimental Cell Research Journal of Biological Chemistry Journal of Cell Biology Journal of Experimental Zoology Nature Proceedings of the National Academy of Science (USA) Science Journals for the broader audience American Scientist Scientific American Science News

Accompanying

Materials

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This

Manual

Each chapter of this lab manual has a corresponding chapter in Vade Mecum that gives background information, illustrates techniques, and shows the living organisms in movies and labeled pictures. The website that augments this laboratory manual as well as Vade Mecum and the FlyCycle CD can be found at www.developmentalbiology.net. The website includes staging series, recipes, suppliers, interactive puzzles, a searchable glossary and other information. The CD-ROM FlyCycle-II, by Mary S. Tyler and Ronald N. Kozlowski, 2002, Sinauer Associates, Sunderland, MA, is an adaptation of the 45-minute film, Fly Cycle: The Lives of a Fly, Drosophila melanogaster, 1996, by M. S. Tyler, J. W. Schnetzer and D. Tartaglia, Sinauer Associates, Sunderland, MA. This covers the life cycle of the fruit fly as well as a number of the mutants used in research. There are a number of superb texts and films in developmental biology. Listed below are three that are cited in each chapter where relevant to help coordinate lecture and laboratory. If you are using a different text in your course, just use the text’s index to find the pages relevant to each laboratory exercise. Text: Gilbert, Scott F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. Films: Fink, Rachel (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Fink, Rachel (ed.). 1995. CELLebrations. Sinauer Associates, Sunderland, MA. Tyler, Mary S., Jamie W. Schnetzer and David Tartaglia. 1996. Fly Cycle, the Lives of a Fly, Drosophila melanogaster. Sinauer Associates, Sunderland, MA. * An earlier version of this CD-ROM is also available as FlyCycle by M. S. Tyler, R. N. Kozlowski, and L. Iten, 1998, Sinauer Associates, Sunderland, MA.

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Selected Bibliography Gopen, G. D. and J. A. Swan. 1990. The science of scientific writing. Am. Sci. 78: 550–558. This article gives excellent, concise descriptions of many common mistakes that make scientific writing unnecessarily obtuse. McMillan, V. E. 1988. Writing Papers in the Biological Sciences. St. Martin’s Press, New York. This is very useful for its suggestions on presentation of data, formats for literature citations, and tips on how to write clearly and accurately. Meyer, A. W. 1939. The Rise of Embryology. Stanford University Press, Stanford, CA. It is instructive to see the laboratory notebooks of famous scientists; unfortunately, facsimiles are not readily available. This book, however, includes a number of plates showing original drawings and text by scientists, including drawings of chick development by Marcello Malpighi and diagrams of mammalian sperm by Antonj van Leeuwenhoek. Moore, R. 1992. Writing To Learn Biology. Saunders College Publishing, Philadelphia. An excellent, delightfully written book. Not only will it help you to write clearly and to recognize bad writing, but it will entertain you along the way with a myriad of humorous and clever examples. Pechenik, J. A. 1997. A Short Guide to Writing About Biology, 3rd Ed. Longman Publ., New York. An excellent writing manual that includes chapters on writing laboratory reports and giving oral presentations. Slack, J. M. 1983. From Egg to Embryo: Determinative Events in Early Development. Cambridge University Press, London. This is a sophisticated, well-written summary of embryonic development. It will introduce you to the major questions rattling around in the minds of presentday developmental biologists. Strunk, W., and E. B. White 1999. The Elements of Style, 4th Ed. Simon and Schuster, Inc., New York. No writing manual has ever topped this one for its helpfulness and conciseness. Wolpert, L. 1991. The Triumph of the Embryo. Oxford University Press, Oxford. Truly a delightful book about embryonic development, this serves as an excellent introduction to the subject and can’t fail to entice you further into the field.

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2

Embryological Tools

Most of an embryologist’s tools are so fine that they have to be made by hand. The ones you will be making are those used most routinely by embryologists. The only indispensable microdissection tool that you will not be making is fine forceps. You can usually buy these (#5) from any biological or electron microscopy supply company (see the end of this chapter for addresses), or your laboratory may have a supply from which you can borrow. Tools to be used on living material must be kept separate from those that have been in fixative. Your dissecting tools from other labs (such as comparative anatomy, where you were up to your elbows in fixed material) must never touch those that you make for living material. Fixative is nearly impossible to remove completely from a tool, and the smallest trace can severely damage living embryological material.

Microknives Knives as fine as the iridectomy knives used by eye surgeons can be made from chips of razor blades mounted on wooden dowels. Protective eye gear and leather work gloves must be worn when breaking the razor blades. Break a razor blade into a number of small pieces by cutting it with heavy-duty scissors, aviation tin snips, or a pair of dikes. If there is an exhaust hood or dissecting chamber in the lab, cutting should be done behind the protection of the glass. The ideal knife is shaped like a half-spear with a long shank (see Figure 2.1A). Prepare a knife handle by first using a pencil sharpener to give a slight taper to the end of a 6-inch length of quarterinch dowel, then clamp the dowel in a vise and make a slit in the tapered end by lightly hammering a heavy-duty scalpel blade half an inch into the dowel end. Holding a prepared piece of razor blade with forceps, insert the shank of the piece into the slit in the dowel. If the blade needs further securing, glue it in place with fingernail polish. Make two microknives. Store the knives with a length of a soft-drink straw over the end to protect the knife blade. Microknives can be sterilized before use in 70% ethanol or in boiling water. Author’s note: Many of the techniques in toolmaking and sterile technique that I pass on were taught to me by my major professor, Dr. H. Eugene Lehman. The many hours I spent in the quiet of his tiny lab were hours of discovery and wonder. It was there, in those cramped, hot quarters, that I was initiated as an embryologist to the mysteries of my subject and the tools that would help me explore those mysteries. I stand in a long line of oral and bench-tutoring tradition, and it is this that I try to pass on to you in these few pages.

Embryological Tools Razor blade chip

Insect needle

2-2

Hair

Cotton plug

Microknife (A)

Microneedle Hairloop (B)

(C)

Bent-tip pipette (D)

Figure 2.1 Embryological tools.

Microneedles Microneedles (Figure 2.1B) are extremely valuable for fine manipulations and can be used as microscissors by drawing one needle past another. You will be using insect pins (#00) mounted in the ends of wooden dowels. Break the plastic ball off the end of an insect pin (you can crush it with forceps). If you would like a shorter pin, cut off a length with wire cutters. Hold a 6-inch length of quarter-inch dowel in a vise; with a hammer and fine finishing nail, make a quarter-inch or longer hole in the end of the dowel. Force the blunt end of the insect pin as far down into the dowel as you can, securing it in place with fingernail polish. Make two needles. Store needles with a length of soft-drink straw over the end to protect the tip. Microneedles can be sterilized before use in 70% ethanol or in boiling water.

Hairloops Hairloops (Figure 2.1C) are one of those romantic additions to an embryologist’s arsenal. Dating from the days of Hans Spemann, when experimental embryology was in its infancy, they have been used to gently coax delicate tissues around a culture dish or to deliver grafts to a graft site. Blond baby’s hair makes the finest quality loops, and for years embryologists have been known to hang around barber shops waiting for the first haircut of a blond child. You will probably want to experiment with several types of hair. Fine blond hair will make the smallest loops. Coarser hair will make larger loops. Basically, you want to force the hair into the smallest loop possible to give you a firm instrument.

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You will be using a Pasteur pipette to hold the hair. Since the opening of the pipette is too large to hold the hair securely, you will be narrowing the opening by heating the bill and drawing it out. You must use protective eye gear for these procedures. Use heavy-duty forceps to hold the pipette at either end, and pass about 2 inches of the bill of the pipette back and forth through the flame of a Bunsen burner to heat it evenly. When the glass is soft, remove it from the flame and quickly pull on both ends using the heavy forceps. This will stretch the glass into a long fiber. When the glass is cool, use a diamond marking pencil to score the glass at a level where the diameter of the bore is approximately 1 mm. Break the glass at this point by pulling in opposite directions on either side of the score line. This will give a clean, perfect break. Do not bend the glass to break it. This will shatter the glass and give a ragged break. Take a length of hair (at least several inches; longer lengths tend to be easier to manipulate) and push both ends into the small end of the pipette. Make sure that the protruding loop is no more than 5 mm long. A large, floppy loop is relatively useless. Melt a small amount of paraffin on a glass slide by placing the slide on a hot plate set at a low temperature or by holding the slide with forceps in a flame. Touch the end of the pipette to the edge of the puddle of melted paraffin. Capillary action will draw the paraffin up into the pipette. Allow the paraffin to harden. Any excess paraffin on the hairloop can be removed by warming a slide, placing filter paper on the warm slide, and touching the hair to the filter paper. Make one or more hairloops. Hairloops can be sterilized in 70% ethanol but not in boiling water or in a flame.

Pipettes Pipettes are among of the most valuable instruments you can have. The appropriate bore size can perform magic at the bench, and you won’t know the appropriate size until you’re sitting there needing it. It therefore is very important to have made a variety of sizes in advance. Always wear protective eye gear when breaking glass or using an open flame. To make widemouthed pipettes, break off the end of a Pasteur pipette where the diameter of the bill begins to widen. As you did when making hairloops, score the glass with a diamond marking pencil where you want the break to be, and give a straight pull on the glass on either side of the score line. To avoid a rough opening, fire-polish the end by passing it quickly through a flame. If you heat the end too much, the opening will get smaller or even close. Make several widemouthed pipettes. To make micropipettes, you will be using Pasteur pipettes and pulling them in a Bunsen burner flame, as you did when making hairloops. Hold both ends of the pipette with heavy-duty forceps and heat approximately 2 inches of the bill until the glass is soft. Quickly remove the pipette from the flame and pull on both ends until the hot glass is pulled out into a thin fiber. When the glass is cool, use a diamond marking pencil to score the glass at a level that will give a bore that is smaller than the original opening. Break the glass by giving a straight pull on either side of the score line. If you choose to firepolish the end, do so with a very quick swipe through the flame. These small openings occlude easily, and often a clean break is better left unpolished. You will be putting a 20° bend in the ends of some of your pipettes (Figure 2.1D). This creates a cradle for your tissues when you are transporting them around a dish. The tissue settles into the bend of the pipette rather than getting stuck way up in the bill of the pipette. To create the bend, heat approximately 2 inches of the bill until the glass is soft. Quickly remove the pipette from the flame, and with heavy-duty forceps, make a 20° bend in the glass about one-half inch from the end. Put bends in some or all of your micropipettes as well as in several unmodified short-billed pipettes.

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All your pipettes must be plugged with cotton (Figure 2.1D). The cotton will filter any contaminants out of the air when the pipette is in use, and it is absolutely necessary for sterile procedures. Take a small wad of cotton and flatten it into a nickel-sized patty. Then take a much smaller wad—about the size of a thick pencil point—and place it in the center of your nickel patty. Use a blunt probe to push against the smaller wad as you push the entire cotton round into the large open end of the pipette. Do not twist the cotton while pushing it in, because this will create air channels that will serve as avenues for contaminants. Make sure no cotton protrudes from the end of the pipette, since this will get caught in the pipette bulb when using it and will cause the plug to be pulled out. Put cotton plugs in all the pipettes that you made as well as in several unmodified short-billed Pasteur pipettes. The pipettes should be sterilized in an autoclave. To prepare the pipettes for autoclaving, wrap them in aluminum foil and attach a name tag written in pencil to the bundle using string or masking tape. You may want to first separate your pipettes according to size, making smaller bundles and marking these according to size. If you do, do a second wrapping so that they are together in a single larger bundle that is tagged with your name. Your instructor will probably autoclave the pipettes for you, doing those of the entire class at the same time.

Embryo Spoons An embryo spoon is one of those tools you seldom use and can’t do without. It is nothing more than a perforated spoon that allows you to pick up embryos and eggs in the several-millimeter to severalcentimeter range without having them float off the spoon. A metal embryo spoon can be bought for around 75 dollars, or you can make one from a cast-off plastic spoon for less than 2 cents. A really cheap, thin, relatively small plastic spoon is best. Heat a clean dissecting needle in a Bunsen burner until it is glowing red, then perforate the bowl of the spoon, making 12–15 holes, evenly spaced, and enlarging each hole with the heated needle so that each is approximately 2 mm in diameter. You will have to reheat the needle periodically. Once the holes are made, you will notice that their edges are very rough. Sand both the upper and lower surface of the spoon with sandpaper until your fingers can no longer feel any rough edges. This sanding is extremely important, because a rough edge can seriously damage an embryo. Test your spoon by putting water in it. The water should drain through the holes. If the spoon holds water instead, the holes need to be enlarged. Your plastic embryo spoon can be sterilized in 70% ethanol, but not, of course, in either boiling water or a flame.

Instrument Tray A tray holds your instruments so that they are both protected and ready for use. The tray should be made out of metal, since this is easily sterilized. A piece of sheet metal 8”×4” can be bent so that it looks like a three-sided pencil tray (Figure 2.2). Use metal cutters to cut V-shaped grooves on each side, so that pairs of grooves line up. The instruments will rest in these grooves. Your instrument tray can be sterilized in a variety of ways, but wiping it down with 70% alcohol is usually the simplest method.

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Instrument tray

Figure 2.2 Instrument tray.

Retooling Metal Instruments Tools such as forceps and scalpels must constantly be rehoned to keep them useful. It is best to use a waterstone for this rather than an oilstone, to avoid oil contamination on your tools. An Arkansas stone (though often used with oil, it can also be used with water) is an economical, high-quality stone. Japanese waterstones are also excellent, but are usually more expensive. None are cheap. Use a soft Arkansas stone or an 800–1200 grit Japanese waterstone for the first coarse honing, and a hard Arkansas stone or a 4000–8000 grit Japanese waterstone for the final polish. Always keep the stone wet with water while sharpening—this keeps metal particles from clogging the pores of the stone—and use the whole surface of the stone to avoid uneven wear. Wash the stone with water after use to remove metal residues and store the stone dry in a soft cloth bag. For a scalpel, as with knives such as those in your kitchen or pocket, sharpening should be done by stroking the blade against the stone in a forward stroke, with the blade tipped at a shallow angle and the cutting edge pointing forward. When the tilt is correct, you will feel the blade biting slightly into the stone. Notice that you are pushing the cutting edge of the blade into the stone, rather than away from it. Trust me, this is the right way to do it. After finishing one stroke, turn the blade over and repeat the stroke on the other side. Check the blade under a dissecting scope. Continue the coarse honing until there are no obvious nicks. Always follow a coarse hone with a fine hone, which should take few strokes in comparison. Always clean the blade carefully between a coarse and fine hone, and after the final hone. Clean the knife by using water and a soft cloth or paper, and wipe toward the cutting edge, never away from it. The retooling of forceps is an art that is easily learned but seldom taught. (This method will not work on your heavy stainless steel forceps, but will on the very fine—#5 or less—forceps, whose tips are so easily damaged.) First examine your forceps under the dissecting scope, closing them slowly to determine if the tips meet precisely, or if they are out of alignment. If the tips are badly damaged, hold the forceps straight up and down and rub them vigorously against the coarse stone to remove the damaged tip. Then, holding the forceps closed, at a slight angle against the stone, and oriented so that the two halves are both touching the stone with equal pressure, grind back and forth. Turn the forceps over and repeat the operation. Now grind the two remaining sides. Examine the forceps under the dissecting scope. You should be regaining a fine tip to the forceps, and the two halves should meet precisely. Do the final set of grindings on the fine stone. If the tips are not damaged but the two prongs are out of alignment, then grind the sides as described above. If your forceps are merely bent slightly at their tip, this sometimes can be fixed by putting the stone between the two prongs and pulling the forceps backward, putting equal force on the two halves. With enough pressure, the ends usually straighten as they come off the stone.

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Sterile Technique Sterile technique is no more than a matter of common sense. If you simply imagine that everything about you is filthy—your hands, your hair, your breath—and that germs are raining down from the heavens, and let this principle guide your actions, then you will master sterile technique quickly. The ammunition that you can use against this filth is limited: it must be germicidal but leave no residue that could harm your tissues. Our list includes ultraviolet (UV) light, 70% alcohol (any higher or lower percentage is less germicidal), wet heat (such as boiling water or an autoclave), and dry heat (such as an oven, autoclave, or open flame). All work areas, instruments, solutions, and cultureware must be sterilized. The workbench A sterile workplace is extremely important. If you have the luxury of building your own workplace, you will want to construct a culture hood (Figure 2.3), paint it with a high-gloss enamel, and install two lights, one a work light and the other a germicidal UV light. The UV light is extremely hazardous to germs, to living things in general, and to your eyes. When the UV light is on, the culture hood must be covered with an opaque curtain (coated bench paper works well) to avoid eye exposure. An hour of exposure to UV is sufficient to sterilize the area, but researchers often leave the UV light on overnight before they work. Never have the UV light on when you are working under the hood, and never leave your cultures in the hood with the UV light on. Most of you will not have the opportunity to build your own culture hood. In this case, your work space on the lab table should be wiped down with 70% alcohol. You must also use 70% alcohol to wipe down the stage of your dissecting scope, if you are using one, as well as its knobs, your instrument tray, and any other items that you will be using. Dissecting tools Your dissecting tools must be sterilized, and then resterilized after each use. Most tools can be sterilized by soaking them for 10 minutes in 70% alcohol, by placing them in boiling water, or by

UV fluorescent bulb Regular fluorescent bulb

Figure 2.3 Homemade culture hood.

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autoclaving them. Specific suggestions for each tool were given in the earlier toolmaking section. For most of your tools, it will suffice to soak them in 70% alcohol. Obtain a lidded jar suitable for this; push cotton to the bottom as a cushion for the tips of your instruments. Label the jar in pencil with your name and fill it with 70% alcohol. (The same alcohol can be used many times.) When you are doing an experiment, sterilize your instruments, tips down, in this alcohol, then rest them on an instrument tray to dry. No alcohol must be left on the instrument when it comes in contact with your culture tissue. Alcohol kills. Some researchers prefer to sandwich their instruments between the folds of a paper towel that has previously been soaked in 70% alcohol instead of using an open instrument tray. This is an extremely effective method of keeping germs away from the instruments, but it has the disadvantage of making the researcher fish blindly for the instrument that is needed. Figure out a method that best suits you. Every time you use an instrument, dip it back in the alcohol (a dip will do), and return it to the instrument tray or sterile towel envelope. Glassware Any glassware that is used must have been cleaned carefully using detergent such as Alconox that is safe for culture use. Glassware must be rinsed very carefully, then re-rinsed in distilled water and airdried. It is then wrapped in aluminum foil, folding over each seam twice so that the seams are airtight. Glassware with no cotton stoppers or plastic tops can be autoclaved in dry heat or sterilized in an oven at 400°F for an hour. Let the glassware cool slowly in the oven after the oven is off before removing it. If the glassware has cotton or plastic tops, or contains fluids, it must be autoclaved in moist heat. Bottles with tops do not need to be wrapped completely in aluminum foil. Cotton stoppers should be covered in foil, and plastic tops should be screwed on only loosely. Once the autoclaving is completed, the screw caps are tightened. Most researchers now use disposable, presterilized plastic dishes for culture chambers. If you can afford them, they are well worth the expense for the convenience. Be aware that they are virtually impossible to resterilize, but they can be reused for nonsterile purposes. Fluids Fluids can be sterilized by boiling, autoclaving, or filtering. The method used depends upon the solution. Water can be boiled. Many solutions, however, cannot, since boiling will change their concentration. Then autoclaving in moist heat will often suffice. Bottles should be filled less than three-quarters full, and tops should be loose during the autoclaving. After autoclaving, when the fluids have cooled, the tops can be tightened. Many solutions, such as physiological saline solutions, cannot tolerate either boiling or autoclaving. Under heat, various of the components, such as calcium chloride and sodium bicarbonate, combine to form a precipitate (in this case, calcium carbonate). In these situations the fluid can be sterilized by passing it through a filter with a porosity that excludes bacteria—a porosity of 0.22 µm or less. There are many sterilizing filter setups available. Some use filtration under vacuum for large quantities of fluids, and others attach to a syringe for smaller quantities. It is an extremely fast method of sterilizing a solution and very handy for last-minute needs. You must have a sterile bottle ready to store the fluid in, of course.

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Rules for the road 1. Never breathe directly over a culture dish. Observe your cultures while they are covered, or from the side whenever possible. 2. If you need to observe your culture uncovered, keep the cover raised above the culture dish to protect it as much as possible from your breath and falling germs. If you must place the cover down, invert it before putting it on your work surface. 3. When not working on your culture, keep your culture dish covered. 4. Never breathe directly into a solution bottle. If you need to smell a solution, waft the vapors toward you with your hand. 5. When using a solution, remove what you need from the stock bottle by pouring or using a sterile, cotton-plugged pipette. Keep the top to the bottle in one hand, pointing downward; do not lay it down. Flame the opening to the bottle after use and immediately put the top back on.

Accompanying Materials See Vade Mecum: “Tools.” This chapter of the CD illustrates, with movies and still pictures, how to make microknives, microneedles, hairloops, micropipettes, and embryo spoons. “Laboratory Safety.” This module illustrates a set of rules that should be followed in the laboratory to ensure safety.

Selected Bibliography Cameron, G. 1950. Tissue Culture Technique. Academic Press, New York. Because of the modern surge in expensive disposables, the art of the reusable is submerged in the older literature. This dated volume is a wonderful, compact addition to the low-cost lab and discusses space, equipment, and sterilization procedures. New, D. A. T. 1966. The Culture of Vertebrate Embryos. Logos Press, London. It is hard to find a volume that discusses embryological tools. This book has a short chapter on several of the classics such as glass bridges, microneedles, and micropipettes. Rugh, R. 1948. Experimental Embryology: A Manual of Techniques and Procedures. Burgess Publishing, Minneapolis, MN. This very old, out-of-print text is both a classic and a gold mine, and it should be bought whenever it is discovered on the dusty shelves of a used bookstore. An introductory chapter discusses some toolmaking techniques and is followed by a wealth of information on amphibian development and experiments.

Suppliers Any good hardware store: Dowels Sheet metal Aviation tin snips, dikes

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Any good biological supply house, such as: Fisher Scientific 585 Alpha Dr. Pittsburgh, PA 15238 1-800-766-7000 www.fishersci.com Syringe filters for sterilizing fluids; order 0.22-µm pore size Vacuum filter units for sterilizing fluids; order 0.22-µm pore size Culture dishes; a number of types are listed under petri dishes Insect pins Germicidal lamps (8 watt); they come in 12- through 36-inch lengths. General Electric makes these and distributes them through science and medical supply houses.

Fine Science Tools, Inc. 373-G Vintage Park Drive Foster City, CA 94404 1-800-521-2109 www.finescience.com This supply house carries very fine and expensive dissecting tools. They also carry excellent fine forceps (Dumont), graded as student forceps, that can be bought in bulk inexpensively. They also carry special razor blades that break easily and insect pins down to #000.

Nasco 901 Janesville Ave. Fort Atkinson, WI 53538-0901 1-800-558-9595 www.nascofa.com This supply house carries inexpensive fine forceps and dissecting tools, with additional savings when items are bought in bulk.

Any good craft or woodworking store, such as: Tandy Leather Company Outlets all over. Catalog from: 1400 Everman Pkway Fort Worth, TX 76140 1-888-890-1611 www.tandyleather.com

Woodcraft 210 Wood County Industrial Park P.O. Box 1686 Parkersburg, WV 26102-1686 1-800-225-1153 www.woodcraft.com Arkansas stones and Japanese waterstones

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Glossary Arkansas stone: A high-quality, but inexpensive, sharpening stone that can be used with oil or water. Autoclave: A machine that is used to sterilize solutions, tools, and glassware, and other materials capable of withstanding heat. The machine uses steam under pressure. Usually the sterilizing chamber is heated to about 121ºC and is under 138-172 kNm of pressure for 12 minutes or longer. Culture hood: An enclosure, designed for laboratory use, that provides a sterile environment for doing culture experiments. It contains a germicidal UV light, that is turned on to sterilize the surfaces and turned off before the hood is used (UV-radiation from a germicidal UV light kills cells and is exceedingly harmful to the skin and eyes). The walls of the culture hood have an easily washable surface that can be sterilized with alcohol or other antimicrobial agent. Commercial hoods called laminar flow hoods include a system of air flow that prevents microbes from entering the hood even though the front of the hood is open. Embryo spoon: A small perforated spoon that allows embryos to be picked up without their floating off the spoon. It can be made inexpensively from a plastic spoon and a heated needle, used to perforate the bowl of the spoon. Germicidal lamp: A UV-bulb that emits light enhanced in the UV-C range. It is used to sterilize an area. It is dangerous to the eyes and skin, and should be turned off before a person enters the work area. Hairloop: An instrument, used by embryologists almost exclusively, that is made from a loop of fine hair (usually baby hair) that is secured in the tip of a Pasteur pipette. It is ideal for manipulating fragile tissues or embryos within a culture dish. Hone: Sharpen. Instrument tray: In an experimental culture set up, it is a tray that holds the tools that will be used. The important aspect of the tray is that it is made of a material, such as metal, that can be repeatedly sterilized either by heat or alcohol. Japanese water stone: A high-quality, though relatively expensive, sharpening stone that is used with water. Microknife: A knife with a small blade, suitable for microdissection under a dissecting microscope. It can be made inexpensively from chips of razor blade, a wooden dowel, and superglue. Microneedle: A needle that is very thin, suitable for microdissection under a dissecting microscope. It can be made inexpensively from inset pins, a wooden dowel, and superglue. Pasteur pipette: A pipette with a long narrow bill with a relatively small bore. It can easily be modified into a pipette with smaller bore by heating the bill, pulling it out when the glass is soft, and breaking it off where the bore is the desired size. Sterile technique: Culture techniques that ensure that cultures are not contaminated by microorganisms, such as bacteria and fungi. Most sterile technique is simply common sense—use sterile instruments, solutions, and containers, and don’t introduce any microorganisms through your hands or the air. Sterilizing methods include ultraviolet light (never expose yourself or your organism to UV-radiation), 70% alcohol, wet heat such as boiling water or an autoclave, or dry heat such as an oven or open flame. Sterilizing filter: A filter that can be used for sterilizing solutions that can not withstand heating. The porosity of the filter must be 0.22 µm or less in order to exclude bacteria.

Embryological Tools Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. What use does an embryo spoon have? After making one, how do you test it to see if it will work properly?

2. What percentage alcohol do you use for sterilizing tools? (This percentage is the most germicidal)

3. What type of needle is useful in making microneedles?

4. How can you use two microneedles to simulate microscissors?

5. What methods can you use for sterilizing your microknives and microneedles?

Name: ______________________

Tools - 2

6. What can a hairloop be used for?

7. Why is it useful to put a slight bend in the bill of a micropipette?

8. What is the function of the cotton plug that you place in the ends of your micropipettes?

9. What method can you use to sterilize your cotton-plugged pipettes?

10. When making a hairloop, which will make a smaller loop, a fine or a coarse hair?

chapter

3

Using the Compound Microscope

The Microscope The compound microscope is an exquisitely refined, expensive piece of equipment. Learning how to use it correctly not only will allow you to achieve the greatest resolution your microscope can give, but it also will avoid costly damage. A microscope has a number of components, all of which you must be familiar with (see Figure 3.1). By the time you finish this lesson, not only will you be able to say “Koehler illumination” three times fast, but you will know what it is, how to use it, and why.

Ocular Ocular focus knob

Arm Nosepiece Stage Objective Mechanical stage control

Iris diaphragm control

Coarse focus

Condenser

Fine focus

Condenser focus

Field diaphragm control

Figure 3.1 Schematic diagram of the compound microscope.

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Safety first First, learn how to carry the microscope. Always carry it by its arm (Figure 3.1), supporting the base with your hand. Once it is at your lab station, determine whether the light source is built in or is an external lamp. If it is built in, plug in the cord and briefly look at all parts of the scope. Find the on-off switch, which is usually built into the base and is often on a rheostat that allows you to vary the intensity of the light with the turn of a knob. If there is a meter to tell you how much light you are using, often this meter will have a red zone at the high end of its scale. Very seldom would you need to have the light on this brightly, and it is very hard on the bulb to do so, significantly reducing its life. Keep light settings low. Stage Examine the microscope’s stage. Most stages have a clip for holding a microscope slide in place. This will be either a pair of stationary spring clips or a spring-loaded lever attached to mechanical controls that can be used to move the slide on the stage back and forth. Mechanical stage controls are usually two knobs set below the stage. Find these knobs and move the mechanical stage to learn which knob governs which direction of movement. Now place a microscope slide in place. If you have a springloaded slide holder, pull the lever back, slip the slide in place, and very gently release the lever. Many a slide is broken by an inadvertent clumsy release of this lever allowing the holder to snap against the slide. If you have a mechanical stage, look at it again to see if it has two vernier scales—one along the top of the stage, the other along the side. As you move the stage, notice that the position of the slide is indicated by these scales. Vernier scales are particularly handy when working with serial sections—for example, when you want to keep track of particular sections out of the many that are on the slide. This is especially useful in photomicrography for recording precisely what objects have been photographed. Magnifications Now look at the objectives held in the nose piece. You probably have three or four. The magnification of each objective is written on its barrel. Find out which magnifications you have. Now look at the magnification of your ocular(s). You have one ocular if your microscope is a monocular and two if it is binocular. The magnification of the ocular is either written on its rim or etched on its barrel. (Barrel markings can be seen by very carefully removing the ocular.) The magnification that you see looking through the microscope is the magnification of the objective multiplied by that of the ocular. Write down the magnifications that your microscope gives. If one of your objectives is 100× oil, then this is an oil-immersion lens and can be used only in combination with immersion oil. Never use this lens unless your instructor allows immersion oil to be used. Oculars Before using a binocular microscope, you must adjust the oculars so that the distance between them matches the distance between your eyes (your interpupilary distance). Usually you can do this simply by pulling the objectives away from each other or pushing them together. Now, rotate the nosepiece so that the lowest-power objective is in place. The microscope should always be stored with this objective in position. Find the focus knobs on your scope. There should be both a coarse- and a fine-focus. As long as the lowest-power objective is being used, you can start focusing by first adjusting the coarsefocus knob to move the objective as close to the slide as it will go. There will always be plenty of

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clearance between the slide and the lower lens of this objective. Make sure that the slide is positioned so that there actually is something in view; then, while looking through the ocular(s), use the coarsefocus knob to slowly move the objective farther from the slide. Objects on the slide will suddenly come into focus. You then use the fine-focus knob to adjust the final focus. If your microscope is binocular, usually one of the oculars will be adjustable by means of a focus knob around its circumference. To set the oculars for your eyes, first fine-focus the microscope on a particular detail, using only the eye that is looking through the nonadjustable ocular. Then close that eye and look through the adjustable ocular with the other eye. Rotate the ocular’s focus knob (not the microscope’s focus knobs) until you again see the same detail in sharp focus. Now look with both eyes through the oculars. Everything should look crisp, and your eyes should feel relaxed. If you do not adjust the oculars to your eyes in this manner, in no time you will start feeling nauseous or have a pounding headache, as your eyes try to compensate for the out-of-focus ocular. It will take the fun out of the day. Sometimes a person feels slightly sick after using the microscope even when the oculars are properly adjusted. In this case, the person is unconsciously focusing images with their eyes rather than focusing the microscope. The trick is to completely relax your eyes as you use the microscope; do all the focusing with the focusing knobs. Objectives We now come back to the objectives. On most microscopes, these are parfocal with one another. “Parfocal” means that, once the specimen is in focus using the lowest power objective, all the other objectives will be roughly focused on the specimen when moved into position. A microscopist always starts focusing with the lowest-power objective in place. Only the sloppy novice bulls ahead using a higher-power objective first. Chide the sloppy novice, for they, in one click of a long-barreled objective, can crash that objective into the slide, cracking both the slide and the objective lens. This is a $400 to $1000 mistake. Always start focusing with the lowest-power objective and move progressively up through higher-power objectives, fine-focusing each one, until you reach the objective that you want. When moving from a lower-power to a higher-power objective, always check from the side to make sure there is sufficient clearance for the objective. A thick slide will never fit under a high-power objective. The quality of the image you achieve with your microscope depends to a great extent on the numerical aperture, or NA, of your objectives and the degree to which the objectives have been corrected for aberrations inherent in lenses. The NA of a lens is a measure of its light-gathering ability—the wider the cone of rays that can be gathered, the higher the NA. And the higher the NA, the better the resolution of that objective. An oil-immersion lens can have a NA greater than 1 (up to 1.3), but all dry objectives have an NA of less than 1. Look at the barrel of each objective to find the number that corresponds to its NA and record this in your lab notebook. Which objective has the lowest NA? Which the highest? Which objective (other than the oil-immersion) gives the best resolution? A microscope lens is subject to spherical and chromatic aberrations. In spherical aberration, the periphery of the lens focuses more strongly than its center. In this case, if you focus on the center of the image, things on the periphery are out of focus. In chromatic aberration, different colors are focused to different planes, causing fringes and halos of color in the image. Focus on a slide using each of your objectives, and determine for which of these aberrations, if any, your objectives have been corrected. Now check to see what is written on the side of each objective. (The type that it is will be abbreviated.) Chromatic aberrations can be corrected to various degrees. Achromat lenses are the least corrected, followed by semiapochromat (fluorite), and finally apochromat lenses, which are the most highly

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corrected for chromatic aberrations. Objectives that are highly corrected for spherical aberration have the word “plan,” meaning flat-field, added to their name. These are top-of-the-line lenses and are highly recommended for photomicrography. A planachromat, therefore, is a very good lens, and a planapochromat is the Mercedes of all lenses. Record what type lenses you have. Now that you are familiar with the basics, it is important that you learn how to achieve maximum resolution from your microscope. The resolution of a microscope is the precision with which you can use it to distinguish separate items. The closer two items can be and still have the microscope resolve them as two items instead of blurring them into one, the better the resolution of the microscope. The resolving power of most classroom microscopes is superb, regardless of how old they are. The physics of optics sets a limit to resolving power of light microscopes, and this limit was achieved by the end of the 1800s. By this time lens makers were able to correct to a great extent the aberrations in glass lenses. (Improvements in contrast techniques, however, continue to be made.) A good microscopist knows how to adjust the settings on the microscope to achieve the best resolution—that is, to set apertures and illumination so that the effects of aberrations in the lenses are minimized. This array of settings is known as Koehler illumination. (This very German word, “Koehler,” is pronounced almost like the very American word “curler.”)

Koehler Illumination The condenser Examine the parts of your microscope that lie below the stage. Most microscopes have a set of lenses here called the condenser. The condenser’s job is to focus light from the light source onto the specimen. Look for a knob that when turned moves the condenser up and down. By moving the condenser, the focal plane of the light is moved. To make sure that it is focused right at the plane of the specimen, you must do the following. First, focus your microscope on an object. Then, if your light source is built in, see if there is also a field diaphragm (see Figure 3.1) on the base of the microscope that can be opened and stopped down to change the diameter of the cone of light coming from the light source. If you do not have a field diaphragm, you can make a temporary one with a heavy index card. Cut a circle of card so it can sit directly on the field lens, use a hole-punch to cut a central hole in the card, and center it on the field lens (where the light is coming through). If you have a field diaphragm, stop it all the way down. Now look through the microscope. You should see a field of light that is smaller than the full field of view; the edges of this field are probably fuzzy. This will be the case if the condenser is not properly positioned. Move the condenser up and down until the edges of the lit field come into sharp focus. Do not change the focus of the microscope during this operation. Remember, you are only moving the condenser up and down. When the edges of the light field are in sharp focus, the condenser is properly positioned to give you the maximum resolution for the particular objective you are using. Now check the position of the condenser. It should be fairly high, several millimeters from its highest position. The field diaphragm now can be opened back up until the cone of light just fills the field of view. If it is opened wider than this, you lose some resolution.

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Iris diaphragm The iris diaphragm must also be adjusted for best resolution. You should find it attached to the condenser (Figure 3.1); it is regulated by a lever. This diaphragm, when properly adjusted, excludes stray, refracted light that would interfere with sharp imaging. The only professional way to adjust this diaphragm is annoyingly time-consuming. You must remove an ocular and look down the barrel of the microscope. You are now looking at the back lens of the objective. Start with the iris diaphragm completely open, then close it down until you see that light is filling two-thirds of the back lens of the objective. Since the setting differs with each objective, the iris diaphragm must be readjusted every time a different objective is clicked into place. If you are doing photomicrography, it is well worth the time to set the iris diaphragm exactly, but it really isn’t worth the time or the risk of dropping the ocular if you are simply studying slides. You can get a good approximate setting by closing down the iris diaphragm just to the point at which the illumination appears to dim. Get used to how objects look when the iris diaphragm is properly set. In general, if the objects look grainy, the iris diaphragm is closed down too far; if they look washed out, the iris diaphragm is open too wide. You have now achieved Koehler bright-field illumination and the maximum resolution for your microscope. Congratulations.

Other Settings When Maximum Resolution Is Not Desired There are times when you do not want maximum resolution from your microscope. Since resolution is gained at the expense of contrast, there may be times when you want to sacrifice resolution to increase contrast. In any type of microscopy, as well as in photography, if there is a need to increase contrast, it must be done at the expense of resolution. Resolution and contrast are inversely related. The higher the contrast, the lower the resolution. Often, specimens on slides are only faintly stained, or you are looking at unstained material. Under these circumstances, contrast must be increased in order to see anything. There are several ways to do this, which the lengthy discussion of Koehler illumination above should reveal to you. Try to discover them on your own before taking the easy way and reading about them below. The first thing a microscopist tries when increased contrast is required is to stop down the iris diaphragm. This gives a very unpleasing grainy texture to the object being viewed, but dramatically increases contrast. Remember, if your specimen is already adequately stained, then stopping down the iris diaphragm seriously compromises the details that you can see. A second method of increasing contrast is to jack the condenser down to a low level. Play around with your microscope settings, view the results, and record both what you did and the results in your laboratory notebook.

Oil Immersion It is very seldom that you would have to use oil immersion. Your lab instructor may prefer that you not use it at all because of the difficulty of properly cleaning the objective lens and slide after use. An example of a specimen for which oil immersion would be useful is a slide of sperm. It is exceedingly important that any slide you use for oil immersion is not a thick one. A wholemount preparation of a chick embryo, for example, would be too thick. There is not enough clearance for the long 100× objective, and it can be damaged by contact with the slide. (Even a cheap oilimmersion lens can cost $150.)

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You can tell if your microscope has an oil-immersion lens because “oil” or “öl” will be written on the barrel along with the magnification. Very fancy microscopes may have lower-power oil-immersion lenses, but most microscopes only have a 100× oil-immersion objective. A special oil, which has the same refractive index as glass, is used as a medium between the slide’s coverslip and the objective. This oil increases the light-gathering power of the lens. With dry lenses, light rays have to pass through glass, into air, and back through glass, and at each interface some light is scattered. With immersion oil, the interfaces scatter less. Resolution is enhanced because more light from the specimen (more information) reaches the lens. The first step in setting up an oil-immersion lens is to focus the specimen, starting on the lowestpower objective and working up to the 40× objective. Set the controls for Koehler illumination at 40×. Then swing the 40× objective out of position without changing the focal plane, put a drop of immersion oil on the slide precisely where the 40× objective had been, and swing the oil-immersion objective into place. The oil should be caught by the lens of the objective and fill the tiny space between coverslip and lens. If there is still an airspace, add a second small drop from the side. Now focus the specimen using the fine-focus only. Remember, your objective lens is very close to the coverslip, and a slight movement of the coarse-focus could bring the lens crashing into the coverslip. You can now reset the settings for Koehler illumination at 100×. You can move the slide around with the mechanical stage. If you don’t have a mechanical stage, moving the slide will be difficult, since a slight movement with the hand will register as an enormous distance under the lens. If you do move the slide, keep checking to make sure that there is still enough oil between the lens and the coverslip. Cleaning up after using oil immersion is critical. If oil is left to dry on the lens, it will take a professional technician to clean it adequately. Use lens paper designed specifically for microscopes. (Never use lens paper for eyeglasses, for example. It is coated with a compound that comes off on the lens and is a trial to remove.) Wipe the lens once in one direction, move to a clean area of lens paper, and wipe the lens again. Repeat this until no more oil shows on the lens paper. Clean the oil from the slide as well. In this case, Kimwipes™ can be used, and even a small amount of soapy water, if necessary. A note of caution: In the past, a type of immersion oil was routinely used that contained PCBs (polychlorinated biphenyls), which are both carcinogenic (cancer-causing) and estrogenic (acts as an estrogen). New federal regulations require that only PCB-free immersion oil be used, but occasionally an old bottle shows up. Check the immersion oil you are using. Make sure that its label says “safe” or “contains no PCBs.”

Make Your $1000 Microscope into a $10,000 Instrument for Pennies Color filters Color filters can be used to increase resolution and contrast. They are extremely useful both for photomicrography and for studying a specimen. You can make a $100 set of filters for approximately $5. Obtain a set of differently colored pieces of cellophane, each approximately 2 inches square. The most useful colors will be red, blue, green, and yellow. The cellophane squares can be mounted in cardboard mounts. These can easily be made from posterboard, or you can use premade mounts that are used for projection slides. Before mounting the cellophane, cut the opening in the cardboard mount so that it is as large as possible. Cut the square of cellophane so that a narrow border of cardboard is

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left around the rim that isn’t covered by the cellophane. Glue or tape the edges closed. Projection slide mounts often are preglued, and an iron is used to close these. You can use an iron specifically made for mounting slides or a regular household iron on a very low setting. Iron only the edges; do not let the iron slide over or touch the cellophane. The filters are used by placing them on the field diaphragm. To increase resolution, use filters with shorter visible wavelengths. Within the visible spectrum, red has the longest wavelength (700 nm); orange, yellow, green, and blue are progressively shorter, in that order; and violet has the shortest (400 nm). Put a slide of a specimen on your microscope. Use each filter you’ve made, and note in your laboratory notebook any differences in resolution. It is helpful to focus on something such as cilia, which are near the limits of resolution of your microscope. You may find that you see best when using a green filter. Even though blue and violet have shorter wavelengths and therefore give better resolution to the microscope, your eye is most sensitive to green. It points out that, ultimately, resolution is a function of both the microscope and your eye. You can also use color filters to increase contrast in a stained section. This is done by using the filter that is the complementary color to the color of the section. A red-stained tissue will look much darker through a green filter, for example. Red and green are complementary, as are blue and orange. Find slides that will allow you to test each one of your filters, and record your observations in your laboratory notebook. Polarizing filters You can make your microscope into a polarizing microscope very easily using polarized film. The polarization system that you can make for less than $2 costs over $500 from a microscope supply company. Polarized film transmits light that is vibrating only in one direction—all other rays are excluded. You will use this property in several applications. Cut the polarized film to make two 2-inch squares. If the polarized film comes with a plastic protective sheet, this can be removed. Etch an arrow into the plastic film to show the direction of light transmittance through each polarizer. You can determine this by looking through the filter at a glare surface, such as a puddle with sunlight reflecting from it. Turn the filter until the glare is eliminated. The direction of polarization in your filter is now at right angles to the glare surface. Check your polarized sunglasses in the same way. In which orientation would you expect the polarized lenses to be mounted? Using polarized light will increase the resolution of your microscope. Put one of your polarizers on the field diaphragm, and focus on a slide. Do you notice an increase in resolution? How does it compare to using a blue filter? Combine both a blue filter and a polarizer. What do you notice? Record your observations in your laboratory notebook. Polarized light can also be used to detect birefringence. Birefringence is the ability of a substance to rotate polarized light and can be used as a diagnostic characteristic of a structure. The highly keratinized layers of skin, for example, are birefringent. (Another word for birefringent is “anisotropic.”) To check for the presence of birefringence, you need two polarizers. One is placed on the field diaphragm, and the other is placed over the eyepiece. The polarizers must be rotated so that they are crossed; that is, their directions of transmittance are at right angles to one another. The easiest way to determine if your polarizers are crossed is to rotate them until the field of view is dark. The light that passed through the first polarizer is blocked out by the second polarizer (called the analyzer). Anything between the two polarizers that is birefringent will rotate light coming through the first polarizer, and this rotated light can now pass through the second polarizer. The effect you see is a

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shining object against a dark field. Highly parallel arrays of closely spaced, linear structures are birefringent. Focus on a slide of adult mammalian skin with hair follicles. Now position your two polarizers so that they are crossed. One should be on the field diaphragm, and the other on the eyepiece. If you do not see any birefringence, rotate the two polarizers so that they maintain their relative orientation to one another until you see the keratinized layers of skin shining. On an actual polarizing microscope, the stage is able to rotate so that you don’t have to rotate the polarizers. Rotating the object in relation to the crossed polarizers maximizes the amount of light that is transmitted. Try slides of other material. Diatoms are exquisite under polarized light. Other good examples of birefringent substances are cellulose, starch, and mitotic spindles. Record your findings in your laboratory notebook. Dark-field optics Dark-field optics can dramatically increase contrast. By placing an opaque disk in the middle of the field lens, a hollow cone of light can be made that illuminates the object but does not enter the objective. Any light that is refracted by a structure will enter the objective and make its image appear self-luminous against a dark field. Dark-field is often used on living material such as ciliated organisms. Since cilia and flagella are near the limits of resolution of your light microscope, they are nearly invisible in bright-field, but the severe contrast that dark-field introduces will make them stand out sharply. Even objects smaller than the resolution of the light microscope, such as microtubules and bacterial flagella, can be seen in dark-field microscopy. To achieve dark-field, you need only to create a hollow cone of light that has a diameter appropriate for the objective you are using. A simple way to do this is to center coins on the lens of the field diaphragm—a quarter for the lowest power objective, a nickel for a higher power, and a dime for the highest. If the diameter is correct, when looking through the ocular you will see just a bare fringe of light around the edge. Try looking at several samples of living material—a scraping from the inside of your cheek, a protozoan culture, a fly wing. Record your observations in your laboratory notebook. Measurement under the microscope The device normally used to measure structures under the microscope is a micrometer. This can be made as a slide or as an insert for the ocular. Micrometers are relatively expensive (about $30), consisting of finely etched glass showing several millimeters, each divided usually into tenths or hundreths. You can make a crude slide micrometer with a glass slide, an etching tool, or a photocopying machine. Get a glass slide, a diamond- or carbide-tip marking pen, and a millimeter ruler. Work under a dissecting scope. Place the slide on top of the ruler under the dissecting scope and etch several centimeters with their millimeter markings into the glass. In the same manner, make a grid micrometer by etching the outline of a square centimeter and dividing it up into millimeter squares. To use your micrometer, place it upside down over a slide that is already in focus. This puts the markings as close to the coverslip as possible. Make sure that there is clearance for this rather bulky prep, and only use the lowest two powers of your microscope. A thinner micrometer can be made by photocopying a millimeter ruler and grid onto a transparency and cutting it out. This makes a very thin preparation, and is therefore preferable to the slide micrometer, but it cannot be sterilized.

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With your micrometer and grid, you will be able to measure the diameters of structures in millimeters and to estimate subfractions of millimeters. Your micrometer and grid will be especially useful under the dissecting scope. Using both your dissecting scope and compound microscope, measure the thickness of the epidermis on the skin slide. Choose something else to measure under the dissecting scope. Record your measurements in your laboratory manual.

Care and Maintenance of Your Microscope: “Twelve Good Rules” “‘The time has come,’ the Walrus said, ‘to talk of many things.’” And those things for us are a most annoying, but most necessary, list of dos and don’ts that will make you a safe microscopist rather than a liability. 1. Always carry the microscope with one hand grasping the arm and one hand supporting the base, keeping the microscope level. 2. Never increase the light so that the meter shows in the red zone. If brief incursions into the red zone are necessary (for photomicrography, for example), keep them short, reducing the light as soon as possible. 3. Never leave the light on when the microscope is unattended. 4. Never force a knob or control. Get help from the instructor if something won’t move. 5. Always move from the lowest-power objective to successively higher-power objectives. 6. Never unscrew an objective unless instructed to do so. If you remove an ocular for adjusting Koehler illumination, do not let go of it, and replace it immediately after the adjustment is made. Dirt that gets down the ocular tube impedes microscopy and can be removed only by a technician. 7. Always clean an oil-immersion lens immediately after use and clean any oil that might have spilled on the stage. Never use alcohols when cleaning; these will dissolve the glue that holds the lenses in place. A small amount of xylene or toluene may be used, but always check with your instructor first. 8. Never leave a slide on the stage when storing the microscope. 9. Always put the lowest-power objective back into place for storing the microscope. 10. Always cover the microscope when storing it. If there is no microscope cover available, get a plastic bag large enough to come down over the scope to a level lower than the stage so that the condenser is adequately covered. 11. Always review these rules until they are reflex. 12. Always take pride in being a fine microscopist, and keep the naive show-offs away from your precision instrument.

Accompanying Materials See Vade Mecum: “Microscope.” This chapter of Vade Mecum gives step-by-step instruction on how to achieve Koehler illumination and how to use color and polarizer filters, and it shows the results that are seen through the microscope. A virtual microscope is also shown in a QuickTime Virtual Reality (QTVR) video in which the parts of the microscope are identified and defined.

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Selected Bibliography Bradbury, S. 1980. Getting the best out of your micrscope. Royal Microscopical Society Proceedings 15: 270–279. A concise and valuable set of tips and explanations. Davidson, M. W. 1991. A photography primer. The Science Teacher 58(7): 12–18. This nontechnical, well-illustrated paper gives practical information about adapting the microscope for polarized and reflected light as well as photography. Delly, J. G. 1988. Photography through the Microscope. Eastman Kodak Company, Rochester, NY. This is a beautifully illustrated pamphlet that concentrates primarily on photomicrography but has an excellent discussion of the principles of light microscopy, Koehler illumination, filters, dark-field, and polarizing methods. Gage, S. H. 1920. The Microscope. Comstock Publishing Co., Ithaca, NY. Originally published in 1908, this book is a testament to how sophisticated microscopes were even in the early 1900s. Packed with information about microscopy and histological preparations, it is still extremely useful. Lacey, A. J. (ed.). 1989. Light Microscopy in Biology: A Practical Approach. IRL Press/Oxford University Press, New York. This is a very sophisticated, but readable, volume that includes an such excellent chapter on the principles of light microscopy. Other chapters are on specialized techniques as immunochemistry, fluorescence, and video microscopy. Needham, G. H. 1968. The Microscope: A Practical Guide. Charles C. Thomas, Springfield, IL. This older volume has lost none of its usefulness. The illustrations are clear, the language straightforward, and the basic information will always be relevant. Sluder, G. and D. E. Wolf (eds.). 1998. Methods in Cell Biology. Vol. 56, Video Microscopy. Academic Press, San Diego. A sophisticated volume that helps make sense of terms and techniques in microscopy.

Suppliers Edmund Scientific Company 101 East Gloucester Pike Barrington, NJ 08007-1380 1-800-728-6999 www.edsci.com Economical scientific equipment and gadgetry Color filters (listed under photography); filters can be bought as individual sheets or a book of 6 Polarizing sheets (listed under optics), including an inexpensive, experimental grade

Any good photography store: Cardboard mounts for projections slides (e.g., Kodak Ready-Mounts)

Any good scientific supply company, such as: Ward’s Natural Science Establishment, Inc. P.O. Box 92912 5100 West Henrietta Road Rochester, NY 14692-9012 1-800-962-2660 www.wardsci.com

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Diamond- or carbide-tip marking pen. (A carbide-tip drill bit from any hardware store is an inexpensive alternative to the more expensive marking pens. For use, wrap all but the tip of the drill bit in tape to protect fingers.) Immersion oil

Glossary Achromat lens: A lens that is corrected for chromatic aberrations for the light of two wavelengths (red and blue). This is the type of lens found in the objectives of most student microscopes. Anisotropic: Having properties that rotate polarized light. Birefringent. Apochromat: A lens that is the most highly corrected for chromatic aberrations, being corrected for four wavelengths of light. Arm: The connecting piece between the top and base of the microscope. When carrying the microscope, always hold onto the arm with one hand and support the base with the other hand. Birefringence: The ability of a substance to rotate polarized light. Birefringence indicates a highly organized parallel array of linear substances, for example, the arrangement of light-meromyosin molecules in a skeletal muscle sarcomere, are birefringent, as are the keratin filaments in the keratinized cells of skin. Chromatic aberration: An aberration in a lens that distorts the colors seen through a lens. Different colors are focused to different planes, causing fringes and halos of color in the image. Coarse and fine focus knobs: Two knobs that control the distance between the objective lenses and the specimen. Usually the larger of the two knobs is the coarse focus knob. Always start focusing your microscope with the lowest-power objective in place. Most microscopes are parfocal; that is, once the specimen is in focus using the lowest power objective, the other objectives will also be roughly in focus when moved into position, and only fine focusing is necessary. Compound microscope: A microscope that uses multiple lenses to increase total magnification. Condenser: A set of lenses that focuses the light onto the specimen. This light must be properly focused to attain maximum resolution. When the condenser is out of focus, resolution is diminished and contrast of the microscope is increased, which can be useful when viewing unstained or poorly stained material. Dark-field optics: A method for using a compound microscope that greatly increases contrast. Basically, an opaque disk is positioned in the middle of the field lens to create a hollow cone of light that illuminates the object but does not enter the objective. Light that is refracted by a structure within the specimen and that enters the objective will be observed. This makes the image appear self-luminous against a dark field. Dark-field is often used on living, unstained material. Field diaphragm: Microscopes with a built-in light source usually have a field diaphragm. This regulates the diameter of the cone of light coming from the light source. This must be properly adjusted to attain maximum resolution of the microscope. Iris diaphragm: A diaphragm attached to the condenser that when properly adjusted excludes stray, refracted light from coming through the condenser that would interfere with sharp imaging. If you close down the iris diaphragm too much, you produce a grainy image with increased contrast at the expense of resolution. Iris diaphragm control: Usually a lever attached to the condenser that adjusts the iris diaphragm. If you close down the iris diaphragm too much, you produce a grainy image with increased contrast

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at the expense of resolution. This can be useful, however, when observing poorly, or unstained, specimens. Isotropic: Having properties that do not rotate polarized light. Not birefringent. Keratinized: Containing high amounts of keratin filaments in organized, parallel arrays. The outer layers of mammalian skin are keratinzed. Koehler illumination: The adjustment of settings on the microscope to achieve the best resolution. These set the apertures and illumination so that the effects of aberrations in the lenses are minimized. Micrometer: A device used to measure structures under the microscope. These are usually made as a slide or as an insert for the ocular. NA: Stands for numerical aperture. Numerical aperture: A measure of the light-gathering ability of a lens—the wider the cone of rays that can be gathered by the lens, the higher the NA, and the higher the NA, the better the resolution of that objective. An oil-immersion lens can have a NA greater than 1 (up to 1.3), but all dry objectives have an NA of less than 1. Objectives: The sets of lenses that fit into the nosepiece of the microscope. There are usually three or four objectives, each marked with its magnification. Ocular: The lens that forms the eyepiece. The magnification seen through the microscope is the magnification of the ocular multiplied by that of the objective. Oil immersion lens: An objective lens on a microscope that is used by placing oil between the lens and the microscope slide. A special oil, which has the same refractive index as glass, is used. This increases the light-gathering power of the lens, and thereby enhances resolution. Parfocal: The ability of a set of objective lenses to keep focus from low power to high power. When lenses are parfocal, once the specimen is in focus using the lowest power objective, all the other objectives will be roughly focused on the specimen when moved into position. Planachromat: The term “plan” refers to flat field. A planachromat lens is an achromat lens that has been corrected for spherical aberrations. Planapochromat: The term “plan” refers to flat field. A planapochromat lens is an apochromat lens that has been corrected for spherical aberrations. This is the highest quality lens for a compound microscope. Polarizer: A filter on a microscope that transmits light vibrating only in one direction—all other rays are excluded. Polarizing microscope: A microscope equipped with two polarizers which allows for the detection of birefringence in specimens. Resolution: The precision with which an optical device can distinguish separate items. The closer two items can be and still be resolved as two items instead of blurred into one, the better the resolution of the device. Semiapochromat: A lens that is corrected for chromatic aberrations for three wavelengths of light. Serial sections: Histological sections of a specimen that are taken sequentially, and kept in order. Usually every section is mounted in correct order for microscopic observations. Spherical aberration: An aberration in a lens that distorts the image seen through the lens. The periphery of the lens focuses more strongly than its center, so that as one focuses on the center of the image, things on the periphery are out of focus. Stage: The platform upon which the slide is placed. Most stages have clips for holding a microscope slide. A mechanical stage has a set of control knobs that allow you to move a slide—one knob for moving the slide from side to side, and another for moving the slide backward and forward.

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Compound Microscope Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. If you are using a 40X objective and a 10X ocular, what is the final magnification that you are viewing through the microscope?

2. What does the condenser do?

3. If the image looks grainy, what might be incorrectly adjusted on the microscope?

4. If the image looks washed out, what might be incorrectly adjusted on the microscope?

5. You are looking at a specimen that, being stained with eosin and hematoxylin, looks primarily red. What color filter would you use to increase the contrast of the image?

Name: _________________________

Microscope - 2

6. If you are looking at a specimen and put a single polarizing filter between the light source and the specimen, have you improved the resolution of the image? Have you improved the contrast of the image?

7. If you observe a specimen under crossed polarizers and see something shining brightly, what does this tell you about the brightly shining structure? What is this phenomenon called?

8. You are given a slide of living, unstained cells. What adjustments would you make to your microscope to ensure that you were able to best observe the cells?

9. What are some of the ways in which you can increase the contrast of your microscope? You should consider adjustments that can be made with the condenser, iris diaphragm, various filters, etc.

10. What adjustments might you make to increase the resolution of your microscope? Consider adjustments to the condenser, iris diaphragm, various filters, etc.

chapter

4

Cellular Slime Molds

Mycetozoa: Dictyostelium discoideum

Life Cycle Scuffing through the moist leaves on a woodland path, you might have disturbed the tiny myxamoebae of the quietly present and seldom seen cellular slime molds of the group Mycetozoa. Myxamoebae are the vegetative stage of these primitive eukaryotes. Classified among the protozoa, cellular slime molds—maligned by their name—are neither slimy nor molds. Their vegetative stage is just the beginning of a simple and fascinating life cycle. This simplicity is one of the reasons for studying these organisms. With only a limited number of cell types, development and behaviors can be examined at the cellular, biochemical, and even genetic levels. The species of Mycetozoa most widely used for such studies is Dictyostelium discoideum.

The vegetative stage The life cycle of Dictyostelium (Figure 4.1) starts with the hatching of myxamoebae from the cellulose encasements of the spores. Under favorable conditions of moisture and warmth, each spore releases a single, haploid (N = 6) myxamoeba (or simply, amoeba). These amoebae crawl on the substrate, feeding on bacteria and dividing by mitosis. In nature, this vegetative stage is probably the stage in which the organism spends most of its time. The amoebae move toward the bacteria, attracted by the folic acid the bacteria secrete. When the food supply is depleted, though, a second stage begins, that of aggregation. Aggregation Upon starvation, the amoebae go into a 4- to 8-hour refractory period when nothing appears to be happening. Using the fine resolution of a biochemist’s magnifier, however, we see that there is an extraordinary list of activities. Starvation triggers a burst of gene activity in the amoebae. Possibly as many as 100 genes are called into action as a complex of new biochemical machinery is assembled: a set of glycoproteins that will enable cell– cell adhesions appear in the cell membrane; adenylyl cyclase, an enzyme that catalyses the formation of cyclic AMP (cAMP; adenosine 3´, 5´-cyclic monophosphate) from ATP is formed; cAMP receptors are put into the cell membrane; and a membrane-bound phosphodiesterase that breaks down cAMP appears, as well as a phosphodiesterase inhibitor. How will all this biochemical machinery be used? Surely something dramatic is about to happen. Suddenly, directed cell movement begins. Amoebae within a certain area start migrating toward a single central amoeba, as if that amoeba had shouted, “OK, on the count of three. . . .” That call to aggregation turns out to be a chemical signal that is secreted first by the central amoeba, and then by

Cellular Slime Molds

Figure 4.1 Life cycle of the cellular slime mold, Dictyostelium discoideum. D. discoideum spends much of its life cycle as feeding myxamoebae on the forest floor. When food runs out, the myxamoebae aggregate to form a migratory pseudoplasmodium (or slug), which searches for a new environment that will be suitable as a feeding ground. Once this is found, the slug settles on its posterior end, taking a “Mexican hat” form, which then culminates into a mature fruiting body. The spores from the fruiting body are released and hatch into myxamoebae, starting the cycle over again.

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Spore mass Spores Stalk

Mature fruiting body

VEGETATIVE STAGE

CULMINATION

Myxamoebae

Mexican hat AGGREGATION

Pseudoplasmodium

all the amoebae in the area. The signal is cAMP. This is the same molecule (often called a second messenger) made by your own cells in response to hormones such as adrenalin, your morning coffee, or your asthma medication. In all of these cases, cAMP is made by adenylyl cyclase and kept within the cell. In the case of Dictyostelium aggregation, however, intracellular cAMP levels first rise by induction of adenylyl cyclase, and then cAMP pours out of the cell. This use of cAMP in the extracellular environment has never been demonstrated in any other organism. The extracellular cAMP acts as a chemotactic agent, and the amoebae move in response to its presence, traveling up its concentration gradient. This is one of the clearest, most experimentally accessible examples of chemotaxis that we as developmental biologists have. With all amoebae pulsing out cAMP, a concentration gradient is established that is greatest at the center of a group of amoebae. The cAMP binds to the cell surface cAMP receptors of the amoebae, and this induces the amoebae to move in the direction of a higher concentration. When the cAMP-receptor sites are all filled so that there is no distinction between the front and back of the cell, an amoeba can no longer sense the gradient, and movement stops. The high levels of bound cAMP activate the phosphodiesterase, which breaks down the cAMP into 5´-AMP, thereby freeing the receptor sites. A phosphodiesterase inhibitor that turns off phosphodiesterase ensures that there is never too much phosphodiesterase made. The cycle of cAMP secretion and amoeboid movement is repeated about once every 6 minutes. An amoeba pulses out cAMP and moves up the cAMP gradient for about 60 seconds, then remains stationary until the next pulse. As the cells move toward the central amoebae, they bump into one another, and because of newly made glycoprotein adhesion molecules in their cell membranes, they adhere and form clumps. This

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results in streams of amoebae, traveling together toward the center. You will see this in your dishes as beautiful branched patterns radiating out from a central mound. There are dozens of questions you can ask about this stage. For example, what would happen if you flooded the substrate with cAMP? If you increased the amount of phosphodiesterase? Pseudoplasmodium Aggregation takes from 8 to 12 hours from the time of starvation. The assembly of amoebae first form a mound, which elongates and finally tips over, becoming a finger-shaped pseudoplasmodium, grex, or slug (Figure 4.2). The slug, usually 2–4 mm long, can consist of only a few hundred or as many as 100,000 cells. It is motile, has anterior and posterior ends, moves only forward, and is attracted toward light, higher temperatures, and higher humidity—conditions that would attract it to the surface in its natural environment. In your own dish, you may see it wandering, seemingly aimlessly, crossing and recrossing its own tracks. You will be able to see a complete record of its wanderings, since the slug lays down a trail—a cellulose sheath—that is easily visible. This sheath, secreted by anterior cells, provides a sleeve through which the slug moves and which collapses in its wake. Though the slug may look relatively undifferentiated at first glance, it contains several cell types. The anterior cells, constituting about 20% of the slug, are prestalk cells. They will form the stalk in the mature fruiting body. The posterior cells of the slug are primarily prespore cells, which become the spores of the fruiting body. Scattered throughout the posterior region is a third cell type, discovered quite recently, called anterior-like cells, which become the basal disc of the fruiting body and the upper and lower caps of the spore mass. Many have asked what the factors are that determine these cell types. As in the development of higher organisms, these sophisticated questions of induction and differentiation can be asked of this simple organism. Not all answers are in, but what is known is tantalizing. Cyclic AMP, as you might have suspected, continues to play a major role. High levels of cAMP appear to induce prespore cells, and a morphogen known as DIF (differentiation-inducing factor), a low molecular weight lipid found in the anterior region of the slug, determines prestalk cells. Little is known about the anterior-like cells. Surprisingly, cells may already be determined prior to aggregation, and their regional separation in the slug may be the result of sorting out of the cell types. One of the major questions of developmental biology is whether the genetic programs of differentiated cells can be changed or even reversed. This is something you easily can ask of Dictyostelium in the laboratory. Suppose you were to divide the slug into separate pieces. Would the cells of each piece form only what they were originally destined to form, or would they regulate to form normal fruiting bodies? What would happen if you disaggregated the cells of the slug—would they reaggregate? If you provided them with bacteria, would they revert to a feeding stage? How stable or labile are these cell types?

Prespore cells Prestalk cells

Figure 4.2 Formation of the pseudoplasmodium. After aggregation, prestalk and prespore cells arrange themselves into a slug by first forming a mound, which elongates and finally tips over, becoming a migrating pseudoplasmodium. The prestalk cells position themselves anteriorly, the prespore cells posteriorly.

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Culmination Transformation of the slug into a fruiting body starts when the slug settles onto its posterior end, looking very much like a Mexican hat (and so it is called), and the cells start to reorganize. You might have guessed already that this would have to happen. The cells in the slug are positioned inappropriately for fruiting body formation. The prespore and prestalk cells will have to switch places. This starts as the anterior tip of the Mexican hat begins secreting a cellulose tube. Cells just behind the tip start migrating up the outside of the tube and into its open end. More posterior cells then start their journey up the sides of the tube, and the tube continues to be laid down and lengthens. In this way, the order of the cells is reversed. The anterior prestalk cells of the slug now form the stalk of the fruiting body, and the more posterior prespore cells of the slug become the spore mass of the fruiting body. This reorganization takes approximately 8–10 hours and results in a beautifully delicate mature fruiting body, usually 1–2 mm high. The stalk cells, now dead, have raised the spore mass high above the substrate. Poised to start the cycle over again, it awaits the proper stimulus to release its spores to the substrate. The fruiting body is a stage that you will want to watch carefully, since it will give you important information about previous experiments. If, for example, you had cut a slug in half and each half formed a fruiting body, the relative sizes of the stalk and spore mass will tell you whether cells were reprogrammed to become different cell types. Also, you can manipulate different stages from Mexican hat to mature fruiting body to ask questions about regulation in these cells. Remember, however, that stalk cells die, and dead cells won’t reprogram. Sexual reproduction: A rare event Though normally reproducing asexually, Dictyostelium can reproduce sexually under conditions such as excessive water and darkness. For sexual reproduction, different mating types must be present. The cycle begins in the aggregation stage when two amoebae in an aggregate fuse, creating a giant cell. This giant cell engulfs the surrounding cells of the aggregate, then encysts in a thick cellulose wall forming a macrocyst, which is resistant to harsh conditions. Within the macrocyst, the giant cell divides meiotically, becoming haploid, and then divides mitotically, producing a number of amoebae that eventually are released as feeding amoebae. Sexual reproduction is rare in these organisms, and it is not always successful. For example, of those macrocysts formed by Dictyostelium discoideum that have been observed in the laboratory, none have been seen to germinate.

Preparing for Your Laboratory Studies The advantages to using Dictyostelium discoideum for our studies are many. The organism is extremely resistant to damage. You can hit it with a hammer and find that you’ve merely asked an interesting question: “What happens when the organism is disaggregated?” Also, the organism’s simplicity invites us to examine complex principles in the context of a minimum of cell types. Developmental biologists have recognized these advantages for years and have used Dictyostelium to ask questions about mechanisms of cell movement and chemotaxis, about the biochemistry of differentiation, and about the genes involved in differentiation and pattern formation. Though genetic manipulations are beyond the scope of this laboratory, many other realms are open to you. Use your imagination. Design simple experiments that will yield sophisticated answers. Keep in mind that a chemical usually has more than one effect. And enjoy this experimental playground.

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Culture procedures The following is a conventional method for culturing slime molds. Petri dishes may already have been prepared for you with nutrient agar, a culture medium described below. A divided petri dish with four wells of nutrient agar makes an excellent set of experiment chambers. If your petri dish is not divided, you can make barriers with sterilized (use 70% alcohol or boiling water) coverslips pushed into the agar. The agar should have a moist but not excessively wet surface. Use sterile distilled water to wet the surface if needed. This culture medium will provide nutrient for Escherichia coli, the bacterium upon which the slime mold amoebae will feed. It is a substrate that is perfectly adequate for all stages of the slime mold’s life cycle. Wherever the amoebae clean out an area of bacteria, they will aggregate to form slugs, and eventually there will be a rich population of slugs and fruiting bodies in the dish. Nutrient agar culture for E. coli Bacto-peptone Bacto-dextrose Να2ΗΠΟ4·12Η2Ο KH2PO4 Bacto-agar Distilled water

10.0 gm 10.0 gm 0.96 gm 1.45 gm 20.0 gm 1000 ml

Buffer to pH 6.6 (usually doesn’t need adjustment). The medium can be autoclaved, or boiled for 3–5 minutes, then poured into petri dishes. Recipe is enough for pouring about 35 plates.

Sterilize a bacteriological loop in a flame by heating it until it is red-hot. Allow it to cool, then immediately use it to transfer E. coli to the first well or small region of your agar plate. This is done by scraping some bacteria from the surface of the stock tube onto the loop and then lightly moving the loop back and forth over the surface of the agar in your plate. This will create streaks of bacteria. (A suspension of bacteria in sterile distilled water also can be used and smoothed over the surface of the agar with a bent glass rod. This will give an even lawn of bacteria rather than streaks.) Resterilize the bacteriological loop and use it in the same manner as above to transfer mature slime mold fruiting bodies and spores to the same well or small region of the petri dish where the bacteria have been placed. Label the dish with your name and date, and place the dish in a protected area of the lab. The optimum temperature range for growth of the slime mold is 22–24°C (room temperature) and optimum relative humidity is 70–75%. The spores will germinate, releasing myxamoebae that will increase by mitosis and feed on the E. coli. This will continue as long as the supply of E. coli remains. Check on your dish as often as time allows during the week, keeping a precise record of the development you observe. Diagram what you see, note variations among different groups: What are the sizes of the aggregating populations that you see? What sizes of slugs do you observe? What different morphologies do you see among fruiting bodies? By the next week, all stages of development should be available to you in your culture dish, since this method does not produce synchronized development. You will do the rest of this lab at that time. If the laboratory instructors have already prepared cultures for you, you will be starting your laboratory exercise at this point. When you receive your dish, answer the questions stated in the paragraph above.

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Make as many observations as you can about the organism before proceeding to the experimental exercises. Examine different stages under the dissecting scope. Put various stages on a slide, and examine them under the compound microscope. Put a spore mass on a slide, and use a needle to burst it open. Look at the spores under high power; there will be about 70,000 in one spore mass. Each spore is ellipsoid and about 5 µm long. Make diagrams of what you see. A note to the instructor: Preparing cultures The most economical way of preparing cultures for class is to make subcultures from stock tubes bought from a biological supply house. In this way, just a few tubes of D. discoideum and E. coli can supply enough material for even a hundred students. Timing is the critical factor. If the class is small, making subcultures 5 days in advance of the class and keeping them at room temperature should work well. If the class is large, it is best to make subcultures 13 days prior to class. Use these to seed further cultures for the individual students 5 days in advance of class.

Experiments Using Dictyostelium You will be designing your own experiments. What follows are suggestions of techniques you can use. When you start an experiment, state clearly in the “Experimental Setup” section of your laboratory notebook the question or questions you are asking. This is absolutely essential. You need to know what you are asking before you can know what the answers are. In addition to your observations of normal development, it is suggested that you do three experiments. Labeling with vital dyes You can mark individual organisms or regions of organisms with vital dyes. A vital dye is one that is not toxic to cells when used in low concentrations. It is toxic at high concentrations, however; the rule of thumb is that a vital dye should not be used at concentrations higher than 0.1%. This is considered a stock solution. The solution must be sterilized before use. This can be done by filtering it through a 0.22-µm porosity filter into a sterile test tube. When choosing a dye, you should always be aware of what elements in the cell the dye stains. Often valuable biochemical information can be gained by noticing the pattern of staining. Useful vital dyes include Nile blue sulfate, neutral red, methylene blue, and toluidine blue (Table 4.1). You can use a vital dye to stain any of the stages in the life cycle of the slime mold. You could stain one group of amoebae and follow them through the rest of the life cycle. You could stain a slug and look for differential staining between prestalk and prespore cells. Move the structure you are staining to an unused quadrant of your culture dish (unless you are staining aggregating amoebae, which should be left in place so as not to disturb their configuration—unless a disturbance is part of the experimental design). Place a drop of dye directly on the cells you are staining. Excess dye will sink into the agar and doesn’t need to be removed. Record which stain you used. Diagram the staining pattern. If you used a metachromatic dye, note regions of primary and secondary colors. What conclusions about biochemical differences can you draw from the differences you see? Observe the organism throughout the rest of its life cycle, noting any changes in color patterns.

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Table 4.1 Some useful vital dyes Vital dye Some properties Nile blue sulfate

More or less specific for phospholipids and lipoproteins of cell membranes.

Neutral red

Beautiful dye with variable staining properties that are difficult to interpret chemically. Stains autophagic vacuoles in prestalk cells of D. discoideum.

Methylene blue

A basic dye. It is a “redox” dye–it is blue when oxidized and changes to colorless when reduced.

Toluidine blue

A basic dye. Extremely useful for its property of metachromasia, which means it has more that one color form. Its primary color, blue, shows when the dye binds to nucleic acids (DNA and RNA). Hence, the chromosomes in the nucleus will stain blue, as will ribosomes and other RNA-containing structures in the cytoplasm. The secondary color, pink, occurs when the stain binds to glycoproteins and mucins (mucins are not found in D. discoideum).

Transecting, grafting, and disaggregating pseudoplasmodia Pseudoplasmodia (slugs) can be transected (cut) into halves, thirds, and so on. They also can be disaggregated into individual cells. A number of interesting questions can be asked in this way. For example, are the fates of the cells of a slug irrevocably set? If a slug is cut in half, what will happen? Can a half-slug survive? Can it differentiate? If so, what will it form? Will the anterior half form only stalk, or will some of the cells alter their fate to become spore cells, thus forming an entire fruiting body? If so, is the fruiting body that results normal in its proportions? Do the results vary depending upon how much time the slug wanders before it settles down to differentiate? What happens if you cut off just the tip of a slug? What do your results say about the flexibility of cells whose fates have already been determined? The more questions you ask of a single experiment, the more acute your observations will be, and the more data you will collect from each experiment. To divide a slug into two or more pieces, first move the slug to an unused quadrant of your culture dish. This can be done by lifting the slug on the flat side of a microknife, or even by lifting out a section of agar along with the slug so as not to risk hurting the slug. You will have an easier time cutting the slug if it is thoroughly cooled first. Place your dish in the refrigerator for 10 minutes, or place it on crushed ice held in a larger dish. Use a microknife to cut the slug. Cuts can be made through the slug into the agar below. The knife is then drawn through the agar without touching the cells. Always make careful diagrams of the position of your cuts. Devise a scheme that will allow you to distinguish one piece from another. For example, each piece could be stained with a different vital dye. Cutting the slug often results in bursting the cellulose sheath that encloses the cells, and instead of two discrete pieces you have a puddle of disaggregated slug. This is fine—don’t despair. Your experiment has simply changed to being a disaggregation experiment. What will happen when you disaggregate a slug? Will the cells reaggregate? If so, will they reaggregate immediately, or will there be a lag period prior to reaggregation? What does this mean at the biochemical level? Will the disaggregated cells behave differently if you give them food (E. coli)? Can you force the life cycle to go backward? Just how flexible is a cell that has already become differentiated as a prespore or prestalk cell?

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Grafting one piece of a slug onto another is rather more difficult than a simple cut, but can be used to ask more complex questions. What would happen if two anterior ends were grafted together? Would there be a battle for anterior dominance? In which direction would the resulting slug move? What would it develop into? The difficult part of grafting is keeping the two pieces together until they have annealed (joined together). It will help if you cool the slugs before cutting. It will also help if the slugs to be cut are close together initially, which diminishes the distance each piece must be moved. Choose the pieces you want to anneal, transfer one to the other by lifting it on the flat side of a microknife or other suitable instrument, and gently push the two pieces together. It will help to keep the dish cool during the first few hours of annealing. You could also dig a shallow depression in the agar where the pieces could sit. The shape of the depression should be such that it helps to force the pieces together. Use diagrams to indicate exactly the regions that were used in the graft. Disruption of cyclic AMP levels Altering cAMP levels will undoubtedly have profound effects on the organism. It is important to know that exogenous (extracellular) cAMP will not cross cell membranes to enter a cell and raise intracellular cAMP levels. There are various chemicals that you could use to disrupt extracellular levels of cAMP. You could use cAMP itself, soaked into an agar cube or flooding an entire area of agar. The concentration of cAMP you use is your choice, but you should at least know some effective ballpark figures. Aggregating amoebae can respond to cAMP levels as low as 0.01 µg/ml. Levels used in chemotaxis experiments are often much higher than this, ranging from 0.05 to 1.2 mg/ml. At 1.2 mg/ml, amoebae fail to continue development. Since cAMP may be something your laboratory cannot supply, there are other sources readily available. The tip of the migrating slug, for example, secretes cAMP and can be removed and placed among amoebae. Also, human urine has relatively high levels of cAMP (approximately 0.53–0.75 µg/ml), well within the range that amoebae respond to. Urine, of course, is a complex substance, and this must be kept in mind when you are drawing your conclusions. For example, urine also contains cGMP (approximately 0.09–0.23 µg/ml), which acts to block the cells’ phosphodiesterase. (This should actually augment the rise in extracellular cAMP levels.) If you use urine, measure its pH (on average, this will be pH 6.3, but normal ranges are from pH 4.5 to pH 8.0), and be aware that it contains some ammonia (0.07–0.13% on average). Basic pH and ammonia are known to favor spore formation and decrease stalk formation. Another method of altering cAMP levels is indirectly, by stimulating or inhibiting the enzymes involved in making cAMP. Dictyostelium’s phosphodiesterase, the enzyme that breaks down cAMP, is secreted at low levels during the amoeboid stage, and it is found at much higher levels, inserted in the membrane just prior to aggregation. During the amoeboid stage, the extracellular phosphodiesterase probably is useful in breaking down the cAMP secreted by bacteria. This would prevent the amoebae from responding prematurely to cAMP. During aggregation, the membrane phosphodiesterase cleans off cAMP binding sites, getting them ready for each new pulse of cAMP. What do you think would happen if you stimulated or inhibited phosphodiesterase during aggregation? In higher organisms, caffeine or theophylline could be used to inhibit phosphodiesterase, thereby raising cAMP levels, but the phosphodiesterase of Dictyostelium is relatively unresponsive to these substances. Caffeine at a concentration of 10 mM (0.194 mg/ml) does not inhibit it, and theophylline will decrease its activity by 15% when present at a concentration of 10 mM (1.8 mg/ml). In fact, caffeine’s effect on Dictyostelium is just the opposite of what you might expect. It actually indirectly

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increases levels of phosphodiesterase activity by inhibiting adenylyl cyclase. Intracellular cAMP levels drop, and this, in turn, may raise phosphodiesterase activity. Five mM caffeine (0.97 mg/ml) can double the activity levels of phosphodiesterase in Dictyostelium amoebae. If you choose to use caffeine or theophylline, you can use the purified form, or a cheaper source—that cup of coffee or tea you’re not allowed to drink in the laboratory. Drinking strengths of coffee contain 0.5 to 1 mg/ml of caffeine, and a cup of tea has about 0.3 mg/ml of caffeine and 0.04 mg/ml of theophylline; these levels are well within the range that should affect amoebae. You may find, as J. T. Bonner did, that as little as 0.025 mg/ml of theophylline can delay aggregation. How might you explain this at the biochemical level? What do you think the effect of caffeine or theophylline would be on a migrating slug or Mexican hat? If you use coffee or tea to ask these questions, I suggest measuring out instant coffee or tea, diluting it with sterile distilled water, recording the concentrations you use, and measuring its pH before applying it. Behavioral studies Prior to starvation, Dictyostelium amoebae show chemotactic behavior toward the folic acid (one of the B vitamins) that bacteria secrete. You could test the various aspects of this behavior. What concentration of folic acid is effective? (A range between 10-6 M and 10 -2 M are ballpark concentrations to try: folic acid can be bought as a vitamin supplement, a 400 mcg pill dissolved in 10 ml distilled H2O makes approximately a 10-4 M solution.) Are cells of a disaggregated slug attracted to folic acid? Dictyostelium slugs show a number of behaviors. For example, they migrate toward light, higher temperature, higher humidity, low ionic concentrations, and the acidic side of a pH gradient. The cue for settling down into a Mexican hat may be the breaking of the surface film of water by the anterior tip of the slug, which it would do under normal circumstances if it were moving upward, toward light. The presence of ammonia, which is copiously produced by the migrating slug, inhibits culmination. You can design experiments that will test various parameters of behavior. For example, will slugs settle into Mexican hats in the absence of light? Are they more sensitive to certain wavelengths of light than others? If you expose a slug to ammonia will you prevent culmination? And if so, for how long? (10–100 mM ammonium hydroxide titrated to pH 7.5 with concentrated phosphoric acid is a concentration range you could use; household ammonia used for cleaning is approximately 3–4 M—diluting it to mM concentrations is something you could try.) What salts encircling an aggregate or slug prevent migration, and why do you think migration is inhibited? (It is presently not known why.) A number of questions can be asked. The trick is to design a setup that will do the asking. A box with a small hole at one end provides a good chamber for testing movement toward light, and different colors of cellophane can be used over the hole to test different wavelengths of light. You will have to observe your cultures over several days to collect your data. Dictyostelium fruiting bodies are surprisingly reactive. They move in response to air currents, for example. And if a dissecting needle is placed near the spore mass, the fruiting body may bend in response to the needle. You can experiment with a number of stimuli (sound, air currents, heat) or substances of different composition (wood, plastic, different metals) to try to determine what specifically the fruiting body responds to. You can also try to determine the stimuli that lead to the release of spores. Is it a response to physical touch, to water? Since these aspects of Dictyostelium biology have been largely ignored by researchers, your results can add new information to our knowledge of this organism.

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Completing your experiments Watch your cultures carefully throughout the rest of the laboratory period—a lot can happen in only a few hours. Also, check your cultures throughout the week as time allows. Keep careful records of any changes. Always use diagrams to document what you see. A great deal of data will be tied up in these diagrams, so make them accurate and proportional. You may run your experiments as long as you wish, or your energy allows. You do not have to terminate them before the next laboratory period. The only constraint is that they be completed before a report on them is due. Have fun and be imaginative.

Accompanying Materials See Vade Mecum: “Slime Mold.” This chapter of the CD shows, in movies, time-lapse photography, and still pictures, each of the life stages of Dictyostelium and techniques used to set up cultures of the organism. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA, Contains an excellent, concise summary of the life cycle, biochemistry, and genetics of Dictyostelium. Fink, R. (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Sequence 2 is an excellent abridged version of one of John Tyler Bonner’s films on Dictyostelium.

Selected Bibliography Berks, M. and R. R. Kay. 1990. Combinatorial control of differentiation by cAMP and DIF-1 during development of Dictyostelium discoideum. Development 110: 977–984. The authors show that differentiation-inducing factor (DIF) is crucial to the differentiation of stalk cells by activating stalk-specific genes and inhibiting spore-specific genes. Bonner, J. T. 1957. Fruiting in Dictyostelium. Precision Film Labs. This is an 11-minute film by John Tyler Bonner. It is superb, and you should find it tucked away in a film rental library at many Universities. Bonner, J. T. 1959. Differentiation in social amoebae. Sci. Am. 201(6): 152–162. Though old, this beautifully illustrated article is far from outdated. It is a wonderful source of ideas for simple but elegant experiments that could easily be done for this laboratory. Bonner, J. T. 1983. Chemical signals of social amoebae. Sci. Am., 248 (4): 114–120. A very readable account of sorting out behavior among two species of slime molds. Bonner, J. T. 1991. Researches on Cellular Slime Molds: Selected Papers of J. T. Bonner. Indian Academy of Sciences, Bangalore, India. Bonner’s work on Dictyostelium discoideum spans five decades and constitutes a major body of information on the organism. His papers are clear and easily understood, with extremely instructive illustrations. The papers are a treasure trove of observations and ideas that will serve students very well as a model for their own reports. Bonner, J. T. 1998. A way of following individual cells in the migrating slugs of Dictyostelium discoideum. Proc. Natl. Acad. Sci, USA 95: 9355–9359. This is yet another example of Bonner’s genius in devising simple ways of getting an organism to reveal details about its biology. Here, Bonner explains how he viewed individual cell migration by creating thin slugs caught between a coverslip and mineral oil on a slide. Bonner, J. T., D. S. Barkley, E. M. Hall, T. M. Konijn, J. W. Mason, B. O’Keefe III and P. B. Wolfe. 1969. Acrasin, acrasinase, and the sensitivity to acrasin in Dictyostelium discoideum. Dev. Biol. 20: 72–87.

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Acrasin is the old term for what is now known to be cAMP, secreted by aggregating amoebae; acrasinase is phosphodiesterase. This paper includes a smorgasbord of experiments, including ones using human urine and theophylline. (Urine acts as an attractant during aggregation, and theophylline inhibits or delays aggregation.) Bonner, J. T., T. A. Davidowski, W.-H. Hsu, D. A. Lapeyrolerie and H. L. B. Suthers. 1982. The role of surface water and light on differentiation in the cellular slime molds. Differentiation 21: 123–126. A paper that beautifully illustrates how, with a minimum of equipment and astute observation, significant discoveries are still to be made. Cavalier-Smith, T. 1998. A revised six kingdom system of life. Biol. Rev. 73: 203–266. How to classify the cellular slime molds has long been in question. It is now generally agreed that they belong among the protozoa, and Cavalier-Smith has designated the group as the Mycetozoa. Devreotes, P. 1989. Dictyostelium discoideum: A model system for cell–cell interactions in development. Science 245: 1054–1058. A sophisticated paper that discusses primarily the mechanisms of signal transduction through cAMP. Gerisch, G. and D. Malchow. 1976. Cyclic AMP receptors and the control of cell aggregation in Dictyostelium. Adv. Cyclic Nucleotide Res. 7: 49–68. Published at a time when cAMP was an extremely hot research topic, this paper summarizes much of the work on cAMP in Dictyostelium and provides a wealth of ideas for laboratory studies. Gomer, R. H. and R. A. Firtel. 1987. Cell-autonomous determination of cell-type choice in Dictyostelium development by cell-cycle phase. Science 237: 758–762. Gomer and Firtel put forward the theory that the determination of amoebae to prestalk or prespore is controlled by their position in the cell cycle at the onset of starvation. Gross, J. D. 1994. Developmental decisions in Dictyostelium discoideum. Microbiological Reviews 58: 330–351. This is a superb summary of much of what we know about the biology and biochemistry of Dictyostelium. Hall, A. L., J. Franke, M. Faure and R. H. Kessin. 1993. The role of the cyclic nucleotide phosphodiesterase of Dictyostelium discoideum during growth, aggregation, and morphogenesis: Overexpression and localization studies with the separate promoters of the pde. Dev. Biol. 157: 73–84. The complex story of phosphodiesterase in Dictyostelium has been unraveled with exquisite finesse by Richard Kessin and his colleagues. This paper summarizes much of that work and demonstrates the delicate precision with which genetics can be studied in this organism. Kay, R. R. and R. H. Insall, 1994. Dictyostelium discoideum. In Embryos, Color Atlas of Development. J. B. L. Bard (ed.). Wolfe Publ., London, pp. 23–35. A clear review of the life cycle of Dictyostelium with exquisite photographs. Kessin, R. H. and M. M. Van Lookeren Campagne. 1992. The development of a social amoeba. Am. Sci. 80(6): 556–565. If you read no other papers on Dictyostelium, read this one. It is a delightfully written, beautifully illustrated article on the general biology, biochemistry, and genetics of Dictyostelium discoideum. Parent, C. A. and P. N. Devreotes. 1996. Molecular genetics of signal transduction in Dictyostelium. Annu. Rev. Biochem. 65: 411–440. A clarifying review of the complex of transduction pathways that are part of Dictyostelium’s biology. Powell-Coffman, J. A. and R. A. Firtel. 1993. Cellular dedifferentiation and spore germination in Dictyostelium may utilize similar regulatory pathways. BioEssays 15: 131–133. This short review discusses the mechanisms of dedifferentiation—how a pseudoplasmodium can be caused to revert back to the stage of feeding amoebae.

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Raper, K. B. 1935. Dictyostelium discoideum, a new species of slime mold from decaying forest leaves. J. Agric. Res. 50: 135–147. This is the report of the discovery of Dictyostelium by Kenneth Raper, then a graduate student, who isolated the organism from soil samples taken on a camping trip while in the Smoky Mountains of North Carolina. Riley, B. B. and S. L. Barclay. 1990. Ammonia promotes accumulation of intracellular cAMP in differentiating amoebae of Dictyostelium discoideum. Development 109: 715–722. The paper describes experiments in which ammonia and high pH were used to promote spore and inhibit stalk formation. The ammonia caused an increase in intracellular cAMP by preventing its secretion. Extracellular cAMP levels were thereby lowered. Riley, B. B. and S. L. Barclay. 1990. Conditions that alter intracellular cAMP levels affect expression of cAMP phosphodiesterase gene in Dictyostelium. Proc. Natl. Acad. Sci. USA 87: 4746–4750. This paper discusses the effects of caffeine on Dictyostelium, showing that the resulting increase in phosphodiesterase activity leads to increased stalk and decreased spore differentiation. Shaulsky, G., D. Fuller and W. F. Loomis. 1998. A cAMP-phosphodiesterase controls PKA-dependent differentiation. Development 125: 691–699. An important paper on the control points of transduction pathways involving cAMP in Dictyostelium.

Suppliers Carolina Biological Supply Co. 2700 York Road Burlington, NC 27215 1-800-334-5551 www.carolina.com

Connecticut Valley Biological Supply Co., Inc. P.O. Box 326 82 Valley Road Southampton, MA 01073 1-800-628-7748 Dictyostelium discoideum. Four or five tubes will seed up to 50 petri dishes. Order two to three weeks in advance. If ordering in the winter, ask for cultures with mature stages of Dictyostelium. Spores can survive an accidental freeze, but earlier stages can not. Escherichia coli (non-mucoid) One or two tubes can be used to grow a lawn of bacteria on nutrient agar, enough for an entire class.

Any good biological supply company, such as: Fisher Scientific 585 Alpha Dr. Pittsburgh, PA 15238 1-800-766-7000 www.fishersci.com 0.22-µm porosity sterilizing filters and filtering apparatus

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Bacteriological loops Nutrient agar supplies Petri dishes Vital dyes

Any good chemical supplier, such as: Sigma Chemical Company P.O. Box 14508 St. Louis, MO 63178 1-800-325-3010 www.sigma-aldrich.com Adenosine 3´, 5´-cyclic monophosphate (cAMP) Caffeine Theophylline

Glossary Adenylyl cyclase: An enzyme that catalyses the formation of cyclic AMP (cAMP; adenosine 3´,5´cyclic monophosphate) from ATP. (This has previously been called adenyl cyclase and adenylate cyclase.) Agar: A gelatinous substance found in red algae. It can be extracted and used as a stiffening agent to produce a gel. In the laboratory, it is often used to make an agar-based culture medium. A 1.5% solution makes a solid but elastic gel when cooled to below 40ºC that will not melt again until it reaches temperatures above 85ºC. Agar is also widely used in the food industry for gelling and thickening; baking recipes may include agar instead of gelatin for thickening in making puddings, etc. Aggregation: The coming together of individual cells to form a single structure. In the cellular slime mold, aggregation of myxamoebae take place after a period of starvation. The aggregated myxamoebae form a migrating pseudoplasmodium (also called grex, or slug). Anterior-like cells: Cells scattered throughout the posterior region of the pseudoplasmodium that become the basal disc of the fruiting body and the upper and lower caps of the spore mass. Caffeine: A drug found in foods such as coffee, tea, and chocolate. In its purified form, it is white and crystalline. A bitter alkaloid, C8H10N4O2. One of its biological effects in higher organisms is to raise cAMP levels by inhibiting phosphodiesterase, the enzyme that breaks down cAMP. CAM: Often used as the abbreviation for cell adhesion molecule. Also used as an abbreviation for the chorioallantoic membrane. A CAM graft is a graft that is placed on the chorioallantoic membrane of a chick embryo. cAMP: Cyclic AMP. cAMP-receptor: A protein that binds cyclic AMP. In Dictyostelium, cells during aggregation have cellsurface cAMP receptors that bind to external cAMP. Cell adhesion molecules (CAMs): Glycoproteins found within cell membranes responsible for adhesion between cells. There are two major categories of cell adhesion molecules, calcium-

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dependent (primarily cadherins) and calcium-independent (those within the immunoglobulin superfamily of molecules). Cellulose: A polysaccharide that occurs as the major structural material in the cell wall of plants. Some fungi have it in their hyphal walls. In the cellular slime mold, it makes up the sheath around the grex, the material for the stalk of the fruiting body, and the capsule around the spores. cGMP: Cyclic GMP. Chemotaxis: Attraction by a chemical that causes cellular migration up a concentration gradient of the chemical. Culmination: The stage following the pseudoplasmodium stage during which the pseudoplasmodium is transformed into a fruiting body. The slug settles onto its posterior end, forming what is called a Mexican hat, and the cells reorganize to form a lower stalk and an upper spore mass. Cyclic adenosine monophosphate: Cyclic AMP. Cyclic AMP: Cyclic adenosine 3´,5´-monophosphate, made from ATP. Within a cell, it is used as a second messenger. Rarely, as in the aggregation of Dictyostelium amoebae, it is pulsed to the outside of the cell and in the case of Dictyostelium amoebae causes directional cell migration up a gradient of cAMP. When cAMP occurs outside of a cell, it does not normally enter a cell. Experimental forms of cAMP, such as dibutyryl cAMP, are capable of entering the cell. Cyclic GMP: Cyclic guanosine 3´,5´-monophosphate. A cyclic nucleotide made from GTP. Acts as a second messenger. Usually found in animal cells at a tenth the concentration of cAMP. DIF (differentiation-inducing factor): A morphogen found in the anterior region of the Dictyostelium slug, a low molecular weight lipid that determines prestalk cells. High levels of cAMP in the posterior region of the slug determine prespore cells. Diploid: Having two of each type of chromosome. 2N is used to indicate the diploid number of chromosomes. Disaggregate: To separate a group of cells into individual cells. E. coli : Escherichia coli, a species of bacterium. Escherichia coli : A species of bacterium. E. coli is a motile, gram negative, rod-shaped bacterium. It is a normal inhabitant of the large intestine of humans and other mammals. It crowds out diseasecausing bacteria and makes vitamin K. The toxic strains of this microbe are responsible for a number of diseases, including many incidences of traveler’s diarrhea. The mutant strain, O157:H7, that has recently been in the news for contaminating undercooked beef from several fast-food establishments, is very dangerous, causing hemorrhagic colitis. Fruiting body: The mature form of a life cycle. In the cellular slime mold it consists of a stalk that is capped by a spore mass, the entire structure is usually 1-2 mm high. Giant cell: The large diploid cell resulting from the fusion of two cells of different mating types as a form of sexual reproduction in the cellular slime mold, a rare event. A sexual phase is stimulated by severe conditions such as excess water and darkness. In this phase, two cells of differing mating types fuse, creating the giant cell, which then engulfs the surrounding cells and forms a macrocyst. Glycoprotein: A molecule containing protein bound to carbohydrates. Glycoprotein adhesion molecules: Cell adhesion molecules (CAMs) found within the cell membrane. There are two major categories of cell adhesion molecules, calcium-dependent (primarily cadherins) and calcium-independent (those within the immunoglobulin superfamily of molecules). Haploid: Having only one of each type of chromosome. 1N is used to indicate the haploid number of chromosomes.

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Macrocyst: A resistant form that results from sexual reproduction, an unusual event, in the cellular slime mold. It consists of a thick cellulose wall enclosing a giant cell, a diploid cell that divides meiotically to become haploid, and then divides mitotically to produce a number of amoebae that are eventually released as feeding amoebae. Metachromasia: The property of some histological stains of having more than one color form. Toluidine blue, for example, has a primary color (blue) when it binds to nucleic acids, and a secondary color (pink) when it binds to glycoproteins. Methylene blue: A basic vital dye. It is a “redox” dye—it is blue when oxidized and changes to colorless when reduced. Mexican hat: An early stage in culmination of the cellular slime mold, when the pseudoplasmodium sits down on its posterior end such that it looks like a Mexican hat. From here, the cells start to reorganize. Microknife: A knife with a small blade, suitable for microdissection under a dissecting microscope. It can be made inexpensively from chips of razor blade, a wooden dowel, and superglue. Morphogen: A diffusible molecule secreted at a distance from its target cells that specifies the fate of the target cells. It may specify more than one cell type through a concentration gradient of the morphogen, in which case different levels of concentration specify different fates. Myxamoeba (myxamoebae, pl): Single, haploid amoeboid organism that hatches from the spore of the vegetative stage of the cellular slime mold. It crawls on the substrate, feeds on bacteria, and divides by mitosis. Often referred to simply as amoeba, though it is not at all equivalent to the protozoan called an amoeba. Neutral red: A vital dye with variable staining properties that are difficult to interpret chemically. Stains autophagic vacuoles in prestalk cells of D. discoideum. Nile blue sulfate: A vital dye that is more or less specific for phospholipids and lipoproteins of cell membranes. Nutrient agar: A culture medium containing agar and nutrients that support bacterial growth, typically made with enough agar to make it a firm substrate for growing cells. Phosphodiesterase inhibitor: An enzyme that inhibits the activity of phosphodiesterase. Phosphodiesterase: An enzyme that breaks down cyclic AMP into (non-cyclic) 5′-AMP. Prespore cells: Cells of the pseudoplasmodium that will later form the spores of the fruiting body. Most of the posterior 80% of the pseudoplasmodium consists of prespore cells. Prestalk cells: Cells of the pseudoplasmodium that will later form the stalk of the fruiting body. The front 20% of the pseudoplasmodium consists of prestalk cells. Pseudoplasmodium: The aggregated form of a cellular slime mold, made up of individual myxamoebae within a cellulose sheath. It is migratory, moving forward and leaving a collapsed cellulose sheath in its wake. The front 20% consists of prestalk cells, and the posterior cells are primarily prespore cells. Slug: In reference to cellular slime molds, a common name for the grex, or pseudoplasmodium stage. Theophylline: A drug found in tea. In its purified form, it is white, and slightly water soluble. It is an alkaloid, C7H8N4O2. One of its biological effects in higher organisms is to raise cAMP levels by inhibiting phosphodiesterase, the enzyme that breaks down cAMP. Toluidine blue: A basic vital dye. Extremely useful for its property of metachromasia, which means it has more that one color form. Its primary color, blue, shows when the dye binds to nucleic acids (DNA and RNA). Hence, the chromosomes in the nucleus will stain blue, as will ribosomes and other RNA-containing structures in the cytoplasm. The secondary color, pink, occurs when the stain binds to glycoproteins and mucins.

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Vegetative stage: An asexual reproductive stage in a life cycle. Vital dye: A dye that stains cells and when used at low concentrations (typically 0.1%) is non-toxic to cells. Examples are neutral red, methylene blue, and toluidine blue.

Cellular Slime Molds Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. During what stage of the life cycle of Dictyostelium does cell division take place?

2. Upon starvation, the Dictyostelium amoebae go into a refractory period before aggregation begins. What do you think might be happening during this period? Consider what changes must occur to go from a non-aggregating to an aggregating stage.

3. If you supplied an external source of cyclic AMP to aggregating amoebae, what would you expect to happen?

4. If you placed a urine-soaked agar cube in the vicinity of aggregating amoebae, what results would you expect and why? (Note: urine is a complex substance that contains relatively high levels of cyclic AMP.)

Name: _________________________

Cellular Slime Molds - 2

5. An aggregate that is in the process of forming a slug can look similar to a slug that is beginning to culminate. What clues can help you determine which stage is which?

6. Does it make sense for the slug to contain more prespore than prestalk cells? Why?

7. Does a slug migrate toward or away from light? Toward or away from higher temperatures? Toward or away from higher humidity? What advantages do you think this behavior has for the organism?

8. How might you test whether the slug migrates towards light?

chapter

5

Gametogenesis

It is typical in general biology courses to learn about mitosis and meiosis all in one breath. This is unfortunate, for it dulls the fascination for the unique nature of meiosis. Think about meiosis: Where does it occur? Only in the gonads—the ovary and testis—and more specifically, only in the germ cells that are lodged in the gonads. Meiosis is the method of cell division that halves the number of chromosomes in the resulting egg and sperm cells. These cells, the gametes, can then fuse with gametes of the opposite type (an egg with a sperm) without multiplying the total number of chromosomes in the resulting zygote. Instead, the normal diploid number of chromosomes is restored. Meiosis is an exotic and profound event. It is the foundation for sexual reproduction. Without meiosis, there would be no sex. Ponder this, and think about its significance as you study the organs that specialize in this event.

Meiosis: An Outline Primordial germ cells arrive in the developing gonad after a long trip, often from the far reaches of the yolk endoderm. Their first task is to increase in number, which they achieve by standard mitosis. This is the gonial stage. Developing eggs in this stage are called oogonia and developing sperm, spermatogonia (Figure 5.1). Germ cells leave the mitotic stage to enter a stage of growth—the primary gonocyte stage—that includes primary oocytes and primary spermatocytes. Primary gonocytes then enter the first meiotic division. During this division, homologous chromosomes separate. The daughter cells are haploid, and each of their chromosomes is made up of two chromatids. This is the secondary gonocyte stage, which includes secondary oocytes and secondary spermatocytes. These enter the second meiotic division, during which the chromatids of each chromosome separate from one another. The resulting cells, gonotids (ootids and spermatids), are still haploid. As described above, every primary gonocyte has the potential to produce four gonotids. This is what occurs in developing sperm. The meiotic divisions are equal, and each primary spermatocyte produces four equal size, small spermatids. The spermatids then go through a further differentiation process called spermiogenesis to become fully formed spermatozoa. Meiotic divisions in eggs, on the other hand, do not produce four equal size cells. Instead, on each meiotic division a very eccentric spindle produces one large cell and one tiny cell. A single primary oocyte, therefore, produces a single large cell, the ootid, and two tiny cells, the polar bodies. (Sometimes the first polar body divides along with the oocyte on the second meiotic division, making a total of three polar bodies.) The polar bodies sit in a clump like bumps on the large spherical ootid and are eventually discarded. Egg development, therefore, involves a lot of genetic waste. For every egg that is produced, three potential egg cells are wasted, all for the sake of retaining as much cytoplasm as

Gametogenesis SPERMATOGENESIS

OOGENESIS

Mitosis

Spermatogonia

5-2

Oogonia

Growth Primary spermatocyte

Primary oocyte

First meiotic division

Figure 5.1 Schematic diagram of gametogenesis. Mitosis in the gonial stage is followed by a period of growth. Cells then enter meiosis. The meiotic divisions in developing sperm are equal. In the developing ovum, however, they are unequal, producing polar bodies and a single large ovum.

Secondary spermatocyte

Secondary oocyte

Second meiotic division

Spermatid

First polar body

SPERMIOGENESIS

Second polar body Ootid

Spermatozoan

possible in the single ootid. Why do you think this is important or necessary? Record your answer in your laboratory notebook. You will be using slide material to study this world of reduction divisions and genetic waste. We will concentrate on mammalian gametogenesis because of its inherent interest to us as gameteproducing mammals. You will be given sections of mammalian (probably rat) testis and ovary on slides. The sections are thin, approximately 6–10 µm, or the thickness of a cell. Think about how thick this is. Scratch your head for a flake of dandruff. The flake is a single epithelial cell from your scalp. It too is about 10 µm thick. Now look back at your slides. The sections are not the color of dandruff, but are an array of at least two contrasting colors. Check the label of the slides to see if you can determine which stains were used. The most commonly used histological stains are hematoxylin and eosin. Hematoxylin is blue-toblack and has an affinity primarily for nucleic acids. The nuclei, therefore, will be stained blue-black. Eosin, a red color, stains the cytoplasm. The contrast of blue against red facilitates the definition of individual cells. Place both slides side by side against a white background (such as a note card), and look at them under the dissecting scope. Note as many features as you can about both the testis and the ovary before proceeding with the detailed instructions. Use your micrometer to measure the dimensions of the ovary and testis. Which one is bigger? Record your measurements in your laboratory notebook.

Mammalian Spermatogenesis As you look at the testis slide under the dissecting scope, you should see a roundish structure surrounded by a tough sheath of connective tissue. This is the tunica albuginea. Within the roundish structure are numerous cross sections through tubules, the seminiferous tubules (Figure 5.2). These tubules should look like little doughnuts. It is in these seminiferous tubules that the sperm are being

Gametogenesis

5-3

produced. Between the tubules, you should see small, triangular islands of tissue containing blood vessels and the interstitial cells, or Leydig cells, which secrete the male sex hormone, testosterone. Most males are incredible sperm-producing machines. The human male, for example, can produce up to 300 million sperm per day and can pack the same amount into a single 3-ml ejaculate. Any organ that can mass-produce a cell to this extent must have an enormous surface area for the task. And this is the case. The seminiferous tubules, U-shaped structures each about 80 cm long, are packed into 250 testicular lobules, each with 1– 4 tubules. If all the seminiferous tubules from just one testis were strung end to end, they would extend about 255 meters (or 279 yards). How far is this? Go outside to the track and run as fast as you can for 30 seconds. Now measure off how far you have run. Unless you are quite extraordinary, it will be about halfway around, or about 220 yards, less than the length of the seminiferous tubules from one testis. This extensive sperm-producing capacity might seem like overkill. However, quantity appears to have its purpose. In humans, a sperm count of less than 20 million sperm per ml of ejaculate is tantamount to being sterile. Apparently, the huge numbers are necessary in helping the one successful sperm to its mark. Other causes of sterility in men can be tight jeans, hot baths, and fevers, all of which raise the temperature of the testis. Sperm development is extremely temperature-sensitive and can only occur in mammals at a temperature 1.5–2.0°C cooler than normal body temperature. This is why the testes are held in scrotal sacs outside the body cavity. In humans, sperm production starts declining at approximately age 45, but often not enough to cause sterility, as is evidenced by the many men who have fathered children in their later years.

Seminiferous tubule

Sertoli cell

Germ cell

Basement membrane

Blood vessel Leydig cell Spermatozoan

Figure 5.2 Diagram of a cross-section through seminiferous tubules within a mammalian testis. Three seminiferous tubules are shown containing a developing sperm and Sertoli cells. The interstitial region between the tubules contains Leydig cells and blood vessels.

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Stereology Now examine the section under the low power of your compound microscope. Use your micrometer to estimate the thickness of the tunica albuginea and the diameter of a seminiferous tubule. Record these in your laboratory notebook. You have already determined the dimensions of the section. Now determine the area of your section of testis (since the section is roughly circular, you can determine area using A = πr2, where π = 3.14). Use your micrometer grid to determine how many cross sections of seminiferous tubules occur in 1 square millimeter. Now, knowing the area of your section, determine how many cross sections of seminiferous tubules occur in the entire section. Since each tubule is Ushaped, half this number should give an approximation of the number of tubules within the testis. If each one were 80 cm long, and they were strung end to end, how far would they extend? How does this number compare to that in humans? You can determine the surface density of the seminiferous epithelium by making a simple calculation used in stereology. First, choose one set of lines (horizontal or vertical) on your micrometer grid and figure out their total combined length. Then count the total number of intercepts these lines make with the tubules. (Any tubule lying on a line intercepts it twice.) This number, multiplied by 2 and divided by the combined length of the lines (in µm) equals the number of square micrometers of epithelium in a cubic micrometer of tissue. Now, if you know the volume of the testis, you can calculate the total area of seminiferous epithelium within it (since the testis is roughly spherical, you can estimate volume using V = 4/3 πr3). The number will be impressive. Tunica albuginea Look at the very thick tunica albuginea. What color does it stain? This will tell you what color collagen stains in this preparation, since the tunica albuginea consists primarily of extracellular fibers of type I collagen. The cells that make this collagen, wherever it occurs in the body, are called fibroblasts. You will see them as small, spindle-shaped cells, embedded among the layers of collagen. Use polarized light to observe the tunica albuginea. Does it shine under crossed polarizers? If so, or if not, what does this tell you about the arrangement of collagen fibers here? Seminiferous tubules The seminiferous tubule is the epithelium that produces sperm. Being an epithelium, it has a free apical surface facing the lumen of the tubule, and a basal surface that sits on a basement membrane. The basement membrane for any epithelium is of major importance to the differentiation and maintenance of that epithelium. It is an acellular layer, made up primarily of type IV collagen, laminin, fibronectin, and heparan sulfate proteoglycan. Examine a seminiferous tubule with a basement membrane. It will be very thin. What color does it stain? Now look back at the tunica albuginea. What color does it stain? Since the major component in each of these is collagen, both should stain the same color. Examine the basement membrane under polarized light. Is it different from the tunica albuginea? What does this tell you? Look at the epithelium carefully. It is made up of two types of cells, the male germ cells at various stages of differentiation as they become spermatozoa, and nurse cells called Sertoli cells. Since development of the germ cells is synchronous at any one level within the tubule, but out of synchrony with other levels, you will see different stages of development in different tubules. By looking at several tubules, you will be able to see germ cells in the spermatogonial, primary spermatocyte, secondary spermatocyte, spermatid, and spermatozoan stages (Figure 5.3). As the cells progress through their

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differentiation, they move closer and closer to the lumen of the tubule. Spermatozoa sit with their tails sticking out into the lumen. The Sertoli cells are nondividing, tall, columnar cells that extend from the basement membrane to the lumen of the tubule. The nuclei of these cells are irregularly shaped and stain lightly, making them easy to identify. Though one cannot tell from light-microscopic study, the Sertoli cells are contiguous, making an unbroken ring around the tubule. The germ cells at all stages of development sit nested in indentations of the Sertoli cells, a fact implying that the Sertoli cells function to support and nourish the germ cells throughout their development. Sertoli cells are also responsible for the translocation of the developing sperm toward the lumen and their eventual release. One of their truly significant functions is to create the blood-testis barrier. The developing sperm are isolated from any immune response as long as they are within the seminiferous tubules, since no immunoglobulins can pass this blood-testis barrier. Why do you think such a thing is necessary? (Remember that the haploid sperm are genetically different from the diploid cells of the body.) If a man has a vasectomy (a tying off of the ductus deferens, the duct leading from the epididymis, as a method of contraception), an immune response is launched against sperm that sit trapped in the ducts above the level of the ligature. If the man has the vasectomy surgically reversed in order to father children, the built-up immune response against his sperm can lead to sterility. Among the Sertoli cells will be the numerous smaller, more darkly stained nuclei of the germ cells. The spermatogonia, with their dark, granular, round or slightly oval nuclei, sit against the basement membrane. They are multiplying through mitosis, so you may see mitotic figures in this layer. The primary spermatocytes sit as a layer above the spermatogonia. Remember that they are in a period of growth, so they are the largest germ cells. Their nuclei also are large and stain relatively darkly. The first meiotic division gives rise to secondary spermatocytes, but you will be lucky to see any, since they almost immediately go into the second meiotic division. You can find them by looking for cells that are half the size of primary spermatocytes. Remember, primary spermatocytes are still diploid, and secondary spermatocytes are haploid. The second meiotic division produces spermatids. These are one-quarter the size of primary spermatocytes and are closest to the lumen. Try to find meiotic figures. You can distinguish first maturation division figures as being larger than second maturation division figures. Diagram the stages of spermatogenesis as seen in tubules on your slide, labeling all structures that are in boldface type above. Spermiogenesis The spermatids in some tubules form a layer three or four cells thick. They are going through a complex process of differentiation called spermiogenesis to become spermatozoa. You should be able to see the following parts of the process on your slide: 1. Elongation of the nucleus and cytoplasm of the spermatid. 2. Condensation (shrinkage and compaction) of the nucleus in what is becoming the head of the sperm and formation of an acrosomal cap. 3. Formation of a flagellum that extends toward the lumen of the tubule. 4. An extensive shedding of cytoplasm that pinches off and appears as dark residual bodies near the lumen of the tubule. These residual bodies will be phagocytosed (eaten) by the Sertoli cells.

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When the sperm are fully formed spermatozoa, only the tip of the head remains attached to a Sertoli cell. See if you can find any fully formed spermatozoa. Eventually, the Sertoli cell will release the spermatozoa into the fluid of the lumen. The entire process of spermatogenesis (from spermatogonium to spermatozoan) takes 48 days in the rat and 64 days in humans. Draw each of the stages of spermiogenesis that you are able to find in your section. Interstitial cells The interstitial cells, or Leydig cells, lying between the seminiferous tubules constitute the mesenchyme of the testis. Look at these cells again. They are large, about 20 µm in diameter, with a spherical, lightly staining nucleus. Though they are relatively few in number, their role is extremely important. In response to leutinizing hormone, LH (yes, I know, you thought only females produced LH), the Leydig cells secrete testosterone, which is required both for spermatogenesis and for development of secondary male characteristics. (Follicle stimulating hormone, FSH, is also produced in males. Its target is the Sertoli cells, causing them to increase their cAMP levels and secrete androgen binding protein.) Spermatozoa

Spermatids Residual body

Sertoli cell

Spermatogonia Primary spermatocyte

Basement membrane Secondary spermatocyte

Figure 5.3 Schematic diagram of a section through a mammalian seminiferous tubule. All stages of sperm development are shown. Spermatogonia are undergoing mitosis. The primary spermatocytes are in a period of growth. In leaving this stage, they divide meiotically to produce secondary spermatocytes, which quickly divide in a second meiotic division to produce spermatids. The spermatids differentiate into spermatozoa, a process called spermiogenesis. Three different stages in this process are shown (although all three stages would never be seen together as drawn): an early stage where the nucleus is condensing and the tail is forming; a later stage where excess cytoplasm is being pinched off as a residual body; and a final stage where the spermatozoa are ready for release. Since sperm development is synchronized at any one level within a tubule, you must look at different tubules or at different levels along a single tubule to see the stages represented.

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Comparative Sperm Morphology Obtain a slide of guinea pig sperm. These are extremely large sperm. Examine a sperm under high power. The acrosome should be obvious (Figure 5.4). This is the bag of enzymes that will open during fertilization in the acrosome reaction. The released enzymes coat the head of the sperm and break down the various extracellular layers surrounding the egg, allowing the sperm smooth sailing toward the egg cell surface. Try to focus on the midpiece and tail of the sperm. Remember that the flagellar tail is near the limits of resolution of your microscope. Use the methods that give you the best resolution of your microscope (Koehler illumination, blue filter; look back at your microscopy lab). Now use various techniques to increase contrast. Which of the methods gives you the best results? Record this information in your laboratory notebook, and make a labeled diagram of the guinea pig sperm. Examine the various types of sperm that are on display. Notice the variety of sizes and shapes that exist among species. Make a diagram of each type that you see, being sure to indicate its species.

Acrosome

Head Nucleus Neck

Midpiece

Tail

Figure 5.4 Diagram of a guinea pig spermatozoan. The acrosome is a bag of enzymes used During fertilization. The nucleus is highly condensed.

Mammalian Oogenesis The ovary does not have the capacity for germ cell production that the testis does. A woman is born with only about 800,000 eggs, and that’s all she’ll have for her lifetime. The oogonial stage, when germ cells are increasing by mitosis, occurs prior to birth, and by the time of birth all the germ cells exist as primary oocytes. During the 30–40 years that a woman has menstrual cycles, she will ovulate about 400 eggs; the remaining 99+% of eggs undergo atresia, or degeneration. (Though this may seem like a tremendous waste, it doesn’t come close to the level of unused sperm males produce.)

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Mesovarium

Tunica albuginea

Hilum

Blood vessel Primordial follicle Early primary follicle

Atretic follicle Medulla

Secondary follicle

Primary follicle

Graafian follicle

Primary oocytes Corpus albicans

Antrum

Figure 5.5 Diagram of a longitudinal section through a mammalian ovary.

Look at your slide of mammalian (probably rat) ovary against a white background under the dissecting scope. You will see a thin connective-tissue sheath, the tunica albuginea, surrounding the teardrop-shaped ovarian tissue. Notice how thin this tunica albuginea is compared to that surrounding the testis. On the narrow end of the ovary, a region called the hilum, you may see the remnants of the mesovarium, a narrow rope of connective tissue that attached the ovary to the broad ligament within the body cavity. Within the ovary you should see two regions: an outer cortex, containing circular structures called follicles, and an inner medulla, containing connective tissue, nerves, lymph vessels, and many large blood vessels. The demarcation between these two regions is indistinct. The germ cells are all located in the follicles within the cortex. A follicle consists of a single egg surrounded by one or more layers of nurse, or follicle, cells. Now look at your slide under the low power of your compound microscope. Look again at the tunica albuginea. You might see a single layer of small, cuboidal cells just outside of the tunica albuginea. This is the mesothelium, an epithelium that is often accidentally brushed off during histological processing.

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Make a diagram of the ovary, and add to it the details of the structures as they are covered in the instructions below. Follicles Under low power, and then higher power, examine the follicles (Figures 5.5 and 5.6). Primordial follicles consist of a primary oocyte, arrested in its first meiotic division, surrounded by a single layer of squamous follicle cells. You should see many of these. Most of the follicles are at this stage. Starting at puberty, and continuing until menopause, each menstrual cycle begins with the growth of 20–50 primordial follicles, stimulated by a pituitary hormone, follicle stimulating hormone (FSH). As soon as the follicles begin to grow, they are known as primary follicles. First, the follicle cells become cuboidal and/or columnar, and they increase in numbers. The multiple layers of follicle cells are the stratum granulosum (these cells will be providing nutrients for the egg). The follicle cells are also secreting a fluid, the liquor folliculi, which gradually accumulates in spaces that develop between follicle cells. Once these spaces appear, the structure can be called a secondary follicle. The spaces constitute the antrum, which eventually becomes enormous, filling the entire central region of the follicle. The egg at this point sits in a little hummock of follicle cells called the cumulus oophorus. Such a follicle is now mature, and can be called a tertiary or Graafian follicle. (It is named after its discoverer, the Dutch anatomist Regnier de Graaf, who first described the ovarian follicle in 1672—not the staid, elderly anatomist we might picture, but a young man of 31. Sadly, he died only a year later.) It is from a Graafian follicle that an egg ovulates. The Graafian follicle contains large amounts of liquor folliculi. This clear, viscous, nutritive fluid contains high amounts of hyaluronate (a glycosaminoglycan, or GAG) as well as hypoxanthine and adenosine. The last two substances have been shown in vitro to keep the oocyte in meiotic arrest, a function they most likely also serve in vivo. Not all growing follicles get to be Graafian follicles. In humans, for example, among the 20–50 that start out on the road to maturation each cycle, usually only one—the dominant follicle—becomes a Graafian follicle. The others undergo atresia, or degeneration, at various points along the way. These atretic follicles, therefore, will be numerous in your section. The cells will look misshapen, and the nuclei will stain darkly. (In mammals with multiple births, such as the rat, more than one Graafian follicle develops each cycle, of course.) Using your dissecting scope and low power on the compound microscope, measure the diameters of each type of follicle with your micrometer. Record these measurements in your laboratory notebook. Theca As the follicles mature, cells outside the follicular cells coalesce around the follicle, forming a theca. As the theca becomes more pronounced, two layers can be distinguished, the theca interna closest to the follicle, and a more fibrous outer theca externa. The theca interna secretes nutritive fluid for the follicle cells, as well as some estrogen- and androgen-intermediary compounds that the follicle cells then convert to estrogen. Oocyte Look carefully at the oocytes in each type of follicle. Notice how much larger the oocyte becomes during follicular growth. The oocyte in a primordial follicle is about 25–30 µm in diameter. Using this as a guide, estimate how large the oocyte is in each of the other types of follicles, and record your number. Before it ovulates, the oocyte has grown considerably. This should impress upon you that the primary oocyte stage truly is a stage of growth.

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Primordial follicle

Follicle cells

Early primary follicle Mid-primary follicle

Primary oocytes Late primary follicle

Theca interna Theca externa

Secondary follicle

Antrum

Cumulus oophorus

Figure 5.6 Schematic diagram of stages in follicle development.

Graafian follicle

Corona radiata Zona pellucida

Now focus on an oocyte in a growing follicle. Notice that immediately surrounding the egg is a translucent area. This is the zona pellucida. It contains various glycoproteins, named ZP1, ZP2, and ZP3. These play a major role in fertilization, inducing the acrosome reaction in the sperm and eventually being altered by cortical granule substances released by the fertilized egg. In their altered form they prevent polyspermy. Notice that traversing the zona pellucida are many fine, filamentous structures. Use various methods to increase contrast in your microscope in order to see these better. These are cytoplasmic bridges between the follicle cells immediately surrounding the oocyte and the oocyte. Nutritive substances such as yolk proteins are being pumped through these channels into the egg during the growth phase. This layer of follicle cells immediately surrounding the egg is called the corona radiata (meaning “radiating crown”). When the oocyte is ovulated, this layer will remain attached to the egg and travel with it into the oviduct. At fertilization, the sperm will have to penetrate the corona radiata and the zona pellucida in order to get to the egg surface. Ovulation of the oocyte is triggered by a surge in luteinizing hormone secreted by the pituitary. Just prior to ovulation, the oocyte completes the first meiotic division and is ovulated as a secondary oocyte. The second meiotic division will begin in the upper end of the oviduct, only to once again enter meiotic arrest. It will not complete meiosis until after fertilization. It is unlikely that you will be fortunate enough to see a secondary oocyte. Look for an oocyte within a Graafian follicle that has a small first polar body on its surface.

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Corpus luteum Once the oocyte is ovulated, the Graafian follicle becomes an endocrine organ, secreting progesterone and estrogen, which maintain the uterine lining in preparation for implantation. The follicle is now the corpus luteum (meaning “yellow body”). It looks collapsed, and its walls are thrown into folds. This structure is huge—1.5 to 2 cm in diameter. Find a corpus luteum on a slide marked as having one. Under a dissecting scope, use your micrometer to measure it, and make a record of these measurements. If pregnancy does not occur, the corpus luteum declines. In humans, this occurs after 14 days, and its decline brings on menses. The declining corpus luteum is replaced by scar tissue and becomes a smaller corpus albicans (meaning “white body”). Find a corpus albicans, measure its dimensions, and make a record of these. How much smaller is it than the corpus luteum? If pregnancy does occur, then the corpus luteum grows. In humans it reaches as much as 5 cm in diameter. (This is really big!) The corpus luteum of pregnancy continues to secrete hormones until the ninth or tenth week of pregnancy, when the placenta takes over this job. This enormous corpus luteum then involutes, and is replaced by scar tissue, becoming a very large corpus albicans. The scar is so pronounced that it usually causes retraction of the ovarian surface. Try to find one of these in your section. Make a record of its measurements.

Histological Hitchcocking It is now time to test your sleuthing abilities. If you have learned your histology well, you will be as clever as the judge below. Little weeping and wailing was heard when the reclusive widow died. She was rich and had kept a l l her money in nylon stockings stashed between the mattress and box springs. The lawyer assigned to t h e case found that there was no will; however, two people had already stepped forward claiming rights to the inheritance. The first was a man who claimed he was the widow’s only child and should inherit a l l . The second was a woman claiming to be this man’s sister, who insisted that the inheritance be shared. The case went to court. The judge listened to the tads of evidence and made a careful study of the pathologist’s report from t h e autopsy. This judge had been a stellar student in developmental biology, and she knew her stuff. Based solely on information in the pathologist’s report, she determined that the woman was lying and awarded the entire inheritance to the man.

How did the judge know? What was that piece of evidence she found in the pathologist’s report? Record your answer in your laboratory notebook.

Accompanying Materials See Vade Mecum: “Gametogenesis.” This chapter of the CD explains meiosis and illustrates in histological sections the process of gametogenesis in mammalian gonads. It also steps you through a stereology exercise using the testis to determine the amount of surface area devoted to sperm production. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. Contains excellent discussion of meiosis and the structure of gametes.

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Fink, R. (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Sequence 4 shows meiosis in a flatworm egg, along with the impressive surface blebbing that occurs prior to each division.

Selected Bibliography Cobb, J. and M. A. Handel. 1998. Dynamics of meiotic prophase I during spermatogenesis: from pairing to division. Sem. Cell. Dev. Biol. 9: 445–450. A review of meiosis in mammalian spermatogenesis. Cormack, D. H. 1987. Ham’s Histology. 9th Ed. J. B. Lippincott, Philadelphia. This is a superb text that includes a lot of incidental information along with the straight histological descriptions. Elias, H. and D. Hyde. 1983. A Guide to Practical Stereology. S. Karger Publ., New York. Any histologist vastly expands the data he or she can squeeze out of a slide by investing a little time in stereology. This short manual is user-friendly, written for the biologist, and will prevent panic attacks even in a severe mathophobe. Fawcett, D. W. 1994. A Textbook of Histology, 12th Ed. Chapman and Hall, N.Y. Originally published with Bloom, this tome is one of the few left from the era when the cell biology, ultrastructure, and light-microscopic details of tissues were taught in a single course. It is superb in its coverage; the extent of details on the mammalian gonads is without match. Frayne, J., and L. Hall. 1999. Mammalian sperm-egg recognition: does fertilin β have a major role to play? BioEssays 21: 183–187. This explains, with colored diagrams, the role of components of the zona pellucida and the sperm cell membrane in the fertilization reaction. Hecht, N. 1998. Molecular mechanisms of male germ cell differentiation. BioEssays 20: 555–561. This review gives a detailed account of the nuclear events during mammalian spermatogenesis. Joyce, I. M., F. L. Pendola, K. Wigglesworth and J. J. Eppig. 1999. Ooctye regulation of kit ligand expression in mouse ovarian follicles. Dev. Biol., 214: 342–353. A research article that demonstrates that mammalian oocytes influence gene expression in their surrounding follicle cells. It’s an excellent example of current research on mammalian oogenesis. Moore, K. L. and T. V. Persaud. 1998. Before We Are Born. 5th Ed. W. B. Saunders, Philadelphia. The second chapter of this very readable short text is an excellent summary of human reproduction. Picton, H., D. Briggs and R. Gosden. 1998. The molecular basis of oocyte growth and development. Molecular and Cellular Endocrinology 145: 27–37. An excellent review of mammalian oogenesis. Reider, C. L. (ed.). 1999. Methods in Cell Biology, Vol. 61, Mitosis and Meiosis. Academic Press, San Diego. A wealth of valuable information on methods for studying the machinery of cell division. Ross, M. H., L. J. Romrell and G. I. Kaye. 1995. Histology: A Text and Atlas. 3rd Ed. Williams & Wilkins, Baltimore. The illustrations in this text are exquisite. In addition to copious colored diagrams and photomicrographs throughout, there are a series of full-page plates at the end of each chapter.

Suppliers Prepared microscope slides of testis, sperm, and ovary can be obtained from such suppliers as: Turtox/Cambosco, Macmillan Science Co., Inc. 8200 South Hoyne Ave. Chicago, IL 60620 1-800-621-8980

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Triarch, Inc. P.O. Box 98 Ripon, WI 54971 1-800-848-0810

Connecticut Valley Biological Supply Co., Inc. P.O. Box 326 82 Valley Road Southampton, MA 01073 1-800-628-7748

Ward’s Natural Science Establishment, Inc. P.O. Box 92912 5100 West Henrietta Road Rochester, NY 14692-9012 Tel. 1-800-962-2660 www.wardsci.com

Glossary Acrosome: A bag of enzymes that sits at the tip of a spermatozoan, above the nucleus. It opens during fertilization, releasing enzymes that coat the head of the sperm and break down the various extracellular layers surrounding the egg. Acrosome reaction: The events in the head of a sperm that opens the acrosomal vesicle, releasing its contents to the outside. The acrosome reaction occurs when the sperm hits the outer extracellular coats surrounding the egg. Components in these coats set off the acrosome reaction. AMH (anti-Müllerian duct hormone): A hormone produced by Sertoli cells within the testis during mammalian embryonic development. It causes the destruction of the Müllerian ducts, the precursors to the oviducts and uterus. Anti-Müllerian duct hormone (AMH): A hormone produced by Sertoli cells within the testis during embryonic development. It causes the destruction of the Müllerian ducts, the precursors to the oviducts and uterus. Antrum: In the mammalian ovary, the space that develops between follicle cells that fills with liquor folliculi. Atretic follicles: In a mammalian ovary, the follicles that degenerate during any cycle of follicle maturation, as the dominant follicles go on to mature. Basement membrane: The extracellular matrix (acellular) found below any epithelium. It creates a cellular barrier between the epithelium and its underlying mesenchyme and is of major importance to the differentiation and maintenance of an epithelium. It is made up primarily of type IV collagen, laminin, fibronectin, and heparan sulfate proteoglycan. Corona radiata: In the mammalian ovary, the follicle cells immediately surrounding the oocyte which will remain attached to the egg when it is ovulated. Corpus albicans: In the mammalian ovary, the scar tissue that replaces a declining corpus luteum.

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Corpus luteum: In the mammalian ovary, the follicle that is left after an egg has been ovulated. It functions as an endocrine organ, secreting progesterone and estrogen, which maintain the uterine lining in preparation for implantation. Cortex: A histological term referring to the outer region of an organ, as opposed to the medulla, which refers to the inner region. Cortical granule reaction: The reaction in the cytoplasm of an egg upon fertilization in which the cortical granules of the egg fuse with the egg cell membrane, releasing their contents to the exterior. The extracellular matrix around the egg is modified by these contents such that further sperm are unable to get to the egg cell surface. Cortical granules: Vesicles found in the cortical cytoplasm of an unfertilized egg. These contain components which are released to the exterior upon fertilization or activation of the egg. These components modify the extracellular substances such that sperm are no longer able to approach the egg cell surface. Cumulus oophorus: In the mammalian ovary, the follicle cells in a tertiary follicle that surround the developing ovum. Diploid: Having two of each type of chromosome. 2N is used to indicate the diploid number of chromosomes. Eosin: An acidic dye that has an affinity for cytoplasmic elements. Its yellowish-red color makes it an ideal counter stain to hematoxylin. Fibroblast: A cell that secretes extracellular fibers, such as collagen and elastin. It is found in mesenchymal tissues and is the most common cell type of connective tissue. Flagellum: A long locomotory organelle that extends from the surface of certain cells such as most spermatozoa. It consists of a 9+2 arrangement of microtubules, nine outer doublets and two inner singlets. Follicle: In a mammalian ovary, the structure that is made up of a developing ovum surrounded by nurse (follicle) cells. Germ cells: The generic term for cells that are, or will become, gametes (sperm or eggs). Glycoprotein: A molecule containing protein bound to carbohydrates. Glycosaminoglycan (GAG): A long, unbranched polysaccharide (formerly called mucopolysaccharide) made up of repeating units of a disaccharide in which one of the sugars is an amino sugar. Complexes with protein to make proteoglycans. Out in the extracellular matrix, they cause extensive swelling of spaces by absorbing water. Examples of GAGs are hyaluronic acid (hyaluronate), chondroitin sulfate, heparan sulfate. Gonad: The organ in which gametes form. In males, this is the testis, a sperm-forming organ, and in females this is the ovary, an ovum-forming organ. In the embryo, the gonad begins developing in the absence of germ cells, and secondarily becomes populated by the primordial germ cells, the cells that will become the gametes. Gonial stage: The first stage in germ cell development during which the germ cells increase their numbers by mitosis. Gonotid stage: The stage in germ cell development following the second meiotic division. The cell at this stage is haploid, with each of its chromosomes consisting of a single chromatid. Graafian follicle: In a mammalian ovary, a mature, or tertiary follicle. Haploid: Having only one of each type of chromosome. 1N is used to indicate the haploid number of chromosomes.

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Hematoxylin: A basic dye that is used as a histological stain. It stains acidic components, primarily nucleic acids, a dark blue and is therefore used as a nuclear stain. Cytoplasmic ribosomes are also darkly stained. Hilum: In the mammalian ovary, the narrow region at the proximal end where the ovary attaches to the broad ligament within the body cavity. Interstitial cells (Leydig cells): In the testis, these are the cells that lie between the seminiferous tubules. They secrete testosterone in response to leutinizing hormone (LH). Leydig cells: The interstitial cells of the testis. These cells lie between the seminiferous tubules. They secrete testosterone in response to leutinizing hormone (LH). Liquor folliculi: In the mammalian ovary, the fluid secreted by follicle cells which accumulates in spaces that develop between follicle cells. The fluid is clear, viscous, and nutritive, containing high amounts of hyaluronate, hypoxanthine, and adenosine. These last two substances appear to keep the oocyte in meiotic arrest. Medulla: A term referring to the inner region of an organ, as opposed to the cortex, which refers to the outer region. Meiosis: The type of cell division that halves the number of chromosomes in cells, taking them from a diploid (2N) to a haploid number (1N) of chromosomes. This occurs only in germ cells. Mesothelium: In the vertebrate, the epithelium that surrounds organs within the peritoneal cavity. Mesovarium: In the mammalian ovary, the narrow rope of connective tissue that attaches the ovary to the broad ligament within the body cavity. Micrometer: A device used to measure structures under the microscope. These are usually made as a slide or as an insert for the ocular. Müllerian duct: In the vertebrate embryo, the duct that will become the oviduct and uterus. In mammals, it also forms the upper end of the vagina. Oogonium: A developing ovum that is in the gonial stage, dividing by mitosis. Ootid: A developing ovum in the gonotid stage. An ootid is haploid, with each of its chromosomes consisting of a single chromatid. The second meiotic division divides the secondary oocyte asymmetrically, producing one ootid and one tiny polar body. Ovary: The female gonad in which germ cells form ova. Polar bodies: The small cells divided off from the developing ovum as it goes through meiosis. Each meiotic division in the egg is asymmetric, resulting in a large cell that will continue on to become the mature ovum and a small polar body, that is eventually sloughed. Polarizer: A filter on a microscope that transmits light vibrating only in one direction – all other rays are excluded. Polyspermy: Fertilization of an egg with more than one sperm. Primary follicles: In a mammalian ovary, the follicle as it begins to grow. The follicle cells become cuboidal and/or columnar and increase in number, forming multiple layers. Primary gonocyte stage: The stage in germ cell development following the gonial stage. During this stage, the germ cells leave the mitotic stage and enter a stage of growth. At the end of this stage, the cell enters the first meiotic division. Primary oocyte: A developing ovum in the primary gonocyte stage, a stage during which the cell is growing and not dividing. At the end of this stage, the cell enters the first meiotic division. Primary spermatocyte: A developing sperm in the primary gonial stage, a stage during which the cell is growing and not dividing. At the end of this stage, the cell enters the first meiotic division.

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Primordial follicle: In a mammalian ovary, a follicle that consists of a primary oocyte, arrested in its first meiotic division, surrounded by a single layer of squamous follicle cells. It is the first stage in follicular development. Primordial germ cells: The early population of cells that give rise to the germ cells. These are typically identifiable early in development. Residual bodies: In mammalian sperm development, the excess cytoplasm that is pinched off from the developing sperm during spermiogenesis. Secondary follicle: In a mammalian ovary, a follicle that has begun to develop an antrum (a space filled with fluid). Secondary gonocyte stage: The stage in germ cell development following the first meiotic division. The cell at this stage is haploid. Each of its chromosomes consists of two sister chromatids. At the end of this stage, the cell enters the second meiotic division. Secondary oocyte: A developing ovum in the secondary gonocyte stage. A secondary oocyte is haploid. Each of its chromosomes consists of two sister chromatids. The first meiotic division divides the primary oocyte asymmetrically, producing one secondary oocyte and one tiny polar body. At the end of this stage, the secondary oocyte enters the second meiotic division. Secondary spermatocyte: A developing sperm in the secondary gonocyte stage. The cell at this stage is haploid. Each of its chromosomes consists of two sister chromatids. The first meiotic division divides the primary spermatocyte symmetrically, producing two secondary spermatocytes. At the end of this stage, the secondary spermatocyte enters the second meiotic division. Seminiferous tubules: The epithelial tubules within the testis where sperm develop. Sertoli cells: The nurse cells that support the development of sperm in the mammalian testis. Also, during mammalian embryonic development, the Sertoli cells secrete AMH (anti-Müllerian duct hormone), which destroys the Müllerian duct. Spermatid: A developing sperm in the gonotid stage. The cell at this stage is haploid, with each of its chromosomes consisting of a single chromatid. The second meiotic division divides the secondary spermatocyte symmetrically, producing two spermatids. At the end of this stage, the spermatid begins the process of spermiogenesis to be come mature spermatozoa. Spermatogenesis: The entire sequence of events in forming a sperm, from the spermatogonial stage to the mature spermatozoan. Spermatogonium: A developing sperm in the gonial stage, dividing by mitosis. Spermatozoan: A mature sperm. Spermiogenesis: The differentiation process that transforms a spermatid into a mature spermatozoan. Stereology: A statistical method by which you extract three-dimensional measurements from twodimensional data, for example, volume measurements of structures in histological sections of tissue. Stratum granulosum: In a mammalian ovary, the multiple layers of follicle cells surrounding a developing ovum. Tertiary follicle (Graafian follicle): In a mammalian ovary, a follicle that has developed a huge antrum that fills the entire central region of the follicle as a single space. Testis: The male gonad in which germ cells form sperm. Theca: Literally, a case. Any sac or casing surrounding an organ or structure. In the mammalian ovary, it is the layer of cells and connective tissue that surround a follicle. Theca externa: In the mammalian ovary, the outer portion of the cellular capsule surrounding a follicle. It is more fibrous than the theca interna.

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Theca interna: In the mammalian ovary, the inner portion of the cellular capsule surrounding a follicle. Tunica albuginea: Literally, a white cloak. This is a connective tissue sheath found around various organs. Around the testis, it is a tough substantial sheath. Around the ovary, it is thin. Wolffian duct: In a vertebrate embryo, the duct that first serves the pronephric and then the mesonephric kidney. A synonym for the pronephric and mesonephric duct. In amniotes, it becomes the ductus (vas) deferens, and epididymis. Zona pellucida: The extracellular coat surrounding a mammalian oocyte. It contains various glycoproteins, ZP1, ZP2, and ZP3, which are important in the fertilization reaction. Zygote: A fertilized egg following fusion of the egg and sperm.

Gametogenesis Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. After which meiotic division in gametogenesis does the germ cell become haploid?

2. What differences are there between the meiotic divisions in a developing sperm and those in a developing ovum?

3. What hormone do Leydig cells secrete?

4. What changes must a spermatid go through to become a spermatozoan?

5. What are some of the major structures of a mature sperm?

Name: _______________________

Gametogenesis - 2

6. Compare the nurse cells (Sertoli cells) in the mammalian testis with nurse cells (follicle cells) in a mammalian ovary.

7. What is the zona pellucida?

8. What cells make up the corona radiata?

9. What is the ovum doing during the primary oocyte stage?

chapter

6

Echinoid Fertilization and Development

Sea Urchins and Sand Dollars

Collecting Have you watched seagulls in flight drop them to smash on the rocks below, pulled the prickly ones from their rocky crevices, or wriggled the fragile, disc-shaped ones from the sand with your toes? Sea urchins and sand dollars, a favored delicacy of seagulls, have also been favored organisms of study for the developmental biologist since the early 1900s. Their development, almost identical, is easily studied since their gametes are readily spawned—and spawned in large numbers. The only problem is finding the adults when they are ripe. If you need gametes in winter, sea urchins are the choice, but they’re often hard to get. On the Northeast coast, the threat of winter ice causes sea urchins to migrate out into deep water; they no longer sit conveniently in the intertidal zone to be picked up by the local kids or developmental biologists. They don’t return to the intertidal zone until their spawning season is virtually over. The common green sea urchin, for example, is ripe and ready to spawn starting in January. If you don’t have a wet suit for diving, you will need accomplices. One good accomplice is the person who sets lobster traps. Sea urchins get caught in lobster traps and are usually discarded. The lobster pound can also be a good source. If you’re willing to compete with the Japanese food trade, you can try one of the many sea urchin processing plants that now dot the coastline. In the past decade, sea urchin processing in this country has escalated into a huge industry that provides the Japanese dinner table with raw ripe gonads. Ships are collecting the urchins for the processing plants just at the time that developmental biologists need them. If you have no accomplices, you can always obtain sea urchins by the more mundane method of placing an order with a biological supply company. Always know the species being provided, the time that they are ripe, and the temperatures that they tolerate (see Table 6.1). When transporting and holding ripe urchins, it is best to pack them in wet seaweed or put them individually into small plastic containers of seawater. Keep them cold (4–8°C). If they are held all together in a single tank of seawater, one sea urchin spawning in the tank will stimulate all the others to spawn. If you don’t have access to seawater, it is easy enough to make artificial seawater or to buy instant ocean from a supply company.

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Table 6.1 Availability of gametes from common North American sea urchins and sand dollars S pe c ie s

Sea Urchins Arbacia punctulata Atlantic sea urchin Lytechinus pictus White sea urchin Lytechinus variegatus Variegated urchin Strongylocentrotus droebachiensis Green sea urchin

Strongylocentrotus franciscanus Red sea urchin Strongylocentrotus purpuratus Purple sea urchin Sand Dollars Dendraster excentricus Eccentric sand dollar Echinarachnius parma Common sand dollar

Nort h Ame ric a n dis t ribut ion

Ripe ga me t e s Re c omme nde d E gg f ound a t e mpe ra t ure dia me t e r ( µm)

East coast: Cape Cod purple Texas; Yucatán West coast: Southern California–south East coast: North Carolina to Florida East coast: Arctic to New Jersey West coast: Alaska to Puget Sound West coast: Alaska to Southern California West coast: British Columbia to Baja California

June–August to spring–fall

fall–spring

20°C–24°C 79 °C Florida;(above is lethal) 16°C–20°C 111 (below 8°C is lethal) 22°C–28°C 103

January– June

0°C–10°C

160

March–July

10°C–17°C

140

December– May

13°C–15°C

80

West coast: Alaska to Baja California

Late spring– summer

10°C–22°C

114

East Coast: Labrador to Cape Hatteras West coast: Alaska to Puget Sound

summer

4°C–16°C

145

aVaries somewhat along its north–south distribution. Source: Primarily after Czihak, 1975 and Strathmann, 1987.

Artificial seawater NaCl KCl MgCl2·6H2O MgSO4·7H2O CaCl2

28.32 gm 0.77 gm 5.41 gm 7.13 gm 1.18 gm

Dissolve in l liter distilled water. Add 0.2 gm NaHCO3 Adjust to pH 8.2, if necessary.

30

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Fertilization The events of sea urchin fertilization have been worked out in great detail. You will be studying both normal fertilization and parthenogenesis (development without fertilization). You will be able to ask some very sophisticated biochemical questions once you know some of the details of these events. We will be injecting potassium chloride (KCl) to induce spawning in both male and female sea urchins. KCl causes a contraction of smooth muscles, and immature gametes will be spawned along with ripe gametes. This can cause problems, and you must be able to recognize a mature egg from an immature one. In the sea urchin, eggs are mature when they are ootids. This means that meiosis is completed and the nucleus is relatively small. An immature egg will be a primary oocyte with a huge nucleus (called a germinal vesicle). The germinal vesicle will be easily recognizable under a dissecting microscope, if you adjust the light to obtain good contrast and focus up and down through the egg. The egg has two extracellular coats: a vitelline envelope, which before fertilization fits snugly around the egg surface and cannot be distinguished; and an outer layer of jelly. This jelly contains a chemical sperm attractant (a small polypeptide that is species-specific). It is not present in the jelly of an immature egg. From a mature egg’s jelly, this attractant diffuses outward, and sperm swim up the concentration gradient. The jelly also contains a relatively species-specific fucose-containing polysaccharide that activates the acrosome reaction in the sperm. This polysaccharide binds to glycoprotein receptors on the head of the sperm, causing the acrosomal vesicle within the head to fuse with the cell membrane and release its enzymes. These enzymes coat the head of the sperm and eat through the jelly, making a path to the egg cell surface for the swimming sperm. In the process, the acrosomal vesicle becomes inverted and greatly elongated by the assembly of actin microfilaments. This elongate structure, the acrosomal process, is what will fuse with the egg cell surface, and it can be seen under the compound microscope under oil immersion if the contrast is maximized. The sperm first binds to the vitelline envelope using a species-specific cell surface protein called bindin, which binds to a bindin-receptor protein on the vitelline envelope. The sperm then fuses with the egg cell membrane, and in so doing causes a brief influx of sodium ions. This influx raises the resting membrane potential of the egg from –70 mV to above 0 mV. Sperm can not fuse with an egg whose membrane potential is above about –10 mV, so this change in membrane potential effectively prevents any additional sperm from fusing. This is called the fast block to polyspermy. It takes only one-tenth of a second to occur, but is not permanent, lasting only about a minute. Since the fast block depends on the availability of sodium in the medium, you can circumvent it by keeping the sodium concentrations in the surrounding seawater artificially low. Binding of the sperm with the egg cell membrane also sets up a second block to polyspermy, the slow block. This block takes a minute to occur, and it is permanent. You will see it as a lifting of the vitelline envelope away from the egg cell surface. This membrane then toughens (involving a chemical process much like tanning leather) and is now called the fertilization envelope. Sperm cannot penetrate this tough layer. The point of sperm entry can be identified by a cone-shaped elevation called the fertilization cone. It represents a tangle of microvilli that have elongated and wrapped themselves around the sperm. The biochemical events that are causing the lifting of the vitelline envelope are initiated through the phosphatidylinositol bisphosphate (PIP) cycle, which is set off by the binding of the sperm to the egg cell membrane. Once the PIP cycle is activated, there is a sudden spike in calcium levels within the egg due to the release of calcium from the smooth endoplasmic reticulum. This spike in calcium levels causes hundreds of vesicles (the cortical granules) housed in the cortical cytoplasm of the egg to fuse with the egg cell membrane and to empty their contents into the perivitelline space between the egg cell

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membrane and the vitelline envelope. The released contents of the cortical granules swell, lifting the vitelline envelope away from the egg cell surface, and tan the envelope, making it tough and impenetrable by other sperm. The PIP cycle also causes a rise in internal pH by activating a sodium–hydrogen ion pump. Sodium ions are pumped into the egg while hydrogen ions are pumped out. The resulting rise in internal pH activates the egg, causing protein synthesis to start and the egg to begin its development. The details above are sophisticated but not beyond your manipulation with relatively simple reagents. Think hard about what you could do to interfere with one or more aspects of the events of fertilization. Instructions for normal fertilization The same instructions can be used for sea urchins or sand dollars, except where noted. You will not be able to sex the animals until they have begun to spawn. Invert five or six animals over dry watch glasses or petri dishes. The five gonadal openings are on the aboral side, and as the animal spawns, gametes will be shed into the dish. On the oral side, you will see the hard white mouthparts (“Aristotle’s lantern”) surrounded by a tough leathery peristomial membrane. You will be injecting through this membrane into the perivisceral cavity. The length of the needle you choose should be long enough to penetrate into the cavity but not much longer. (When spawning sand dollars, a much shorter needle is used than when spawning sea urchins.) Use a syringe to inject 1–2 ml (0.5–1 ml for sand dollars) of isotonic KCl (0.53 M = 3.9%). This will cause the smooth muscles of the gonads to contract and spawn their gametes. After 2–5 minutes, repeat the injection. This gives a heavier spawning. As soon as spawning begins (within minutes of injection if the animals are ripe), check the color of the gametes. This will tell you the sex of the animal. Sperm are creamy white; eggs are yellow, pink, or dark red (depending on the species). For males Immediately pour off the first sperm to get rid of perivisceral fluid, since this will interfere with the sperm’s ability to fertilize. Then allow the animal to shed into the watch glass or petri dish without diluting the sperm. This is called dry sperm. Dry sperm will be good for 6–10 hours at room temperature and even longer if kept in the refrigerator. For females Allow the female to shed eggs into seawater by placing her, inverted, on top of a beaker of seawater. The beaker should be small enough that the animal does not fall in. The beaker must be full enough so that seawater touches the aboral side of the animal. As the animal sheds, the eggs will drift down through the seawater and settle in the bottom of the beaker. After shedding is completed, decant off the seawater and add fresh. Do this twice. This washes the eggs of perivisceral fluid which interferes with fertilization. (N.B.: Sometimes artificially spawning sea urchins will cause parthenogenetic activation of the eggs. Examine egg batches for raised fertilization envelopes, which indicates parthenogenetic activation. These eggs could be used in the parthenogenesis section of the chapter.) To fertilize the eggs, first make a standard sperm suspension of 1 drop of dry sperm in 10 ml of seawater. This suspension must be used within 20 minutes and then discarded. Dry sperm are relatively inactive. Diluted, however, they become very active and quickly use up their energy stores. Use sterile procedure and sterile glassware if you are keeping the eggs for culturing. Add two drops of standard sperm suspension to 10 ml of seawater containing eggs. Repeat after two minutes. After 10 minutes, decant off the seawater and add fresh. This culture can be kept for observing normal development. Know the species you are using, and decide on an appropriate temperature (or range of

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temperatures) for rearing (see Table 6.1). For long-term cultures, refrigerator temperatures are usually adequate. The seawater for culturing should be sterilized by filtering it through a 0.22-µm porosity filter. Streptomycin can be added (2 mg/liter) to retard bacterial growth, but is normally not needed in cultures maintained at cold temperatures.

Watching Fertilization You can watch fertilization under the microscope by placing eggs on a slide and introducing sperm from one side. One way to do this is to place a large drop of egg suspension on a microscope slide. Put the slide on the microscope stage and have a footed coverslip close at hand. (A footed coverslip is made by nicking the corners of the coverslip against some hard paraffin, so that crumbs of paraffin remain attached at each corner. This will give just enough spacer between the slide and the coverslip to avoid crushing the eggs.) Then put a drop of sperm suspension on the slide close to but not touching the drop of eggs. Place the footed coverslip on the slide so that it covers both drops. This will cause the two drops to mix. Immediately focus on eggs that are mature. (Remember, those with large germinal vesicles are immature.) Use a clock with a second hand to time the events that you see. You should be able to determine the exact spot of sperm entry, which will be marked by the fertilization cone (Figure 6.1). Time the raising of the fertilization envelope. Do you see any eggs in which the fertilization envelope starts to rise but doesn’t finish? Record any variations you see. Once the fertilization envelope is raised, do sperm continue to be attracted to the egg? Do the sperm bounce off the fertilization envelope or stick to it? Enter your answers in your notebook, along with any other observations you make. If you can find some immature eggs, place these separately on a slide, and watch as you introduce sperm from the side. Are the sperm attracted to the egg? Compare this with sperm behavior near a mature egg. Prepare a slide of a fertilized egg, place a footed coverslip over it, and look at it under a 40× objective. Close down the iris diaphragm on your microscope to increase contrast. Focus on sperm that are caught in the jelly surrounding the egg. Can you see any acrosomal processes? Why would you expect to see them? Enter your answer in your notebook, and diagram what you see. Do not go to oil immersion, since this will not work using these wet mounts and will only mess up the microscope by getting seawater on the objective lens. Put a drop of sperm suspension on a slide, and place a coverslip over it. Observe this under a 40× objective, and close down the iris diaphragm to increase contrast. Diagram the sperm. Do you see any acrosomal processes on these? Why? Record your answers.

Interfering with Fertilization Many of the events important to fertilization depend upon the jelly surrounding the egg. What do you think would happen if you removed this jelly? You can remove it by exposing the eggs to seawater that has been adjusted to a pH of 5.0 by the addition of 0.1N HCl (added dropwise while measuring the pH with a pH meter). Swirl the eggs and return them to normal seawater after 2 minutes. When sperm are added to these eggs, how do the sperm behave? Does fertilization occur? What did you expect would happen? Enter your questions, answers, and observations in your notebook.

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Remember that certain events in fertilization rely on sodium pumps in which sodium is being pumped into the egg. If sodium were unavailable to the egg, what might the results be? Design several questions you could ask, and try the fertilization procedures using sodium-free seawater. What do you think your results will be? What are your results? How might you explain any discrepancies between the expected and the actual results? Enter your questions and answers in your notebook. Sodium-free seawater Glycerol MgCl26·H2O KCl NaHCO3 CaCl2

84.60 ml 11.20 gm 0.77 gm 0.21 gm 0.12 gm

Make up to 1 liter with distilled water.

Remember that calcium also is extremely important in fertilization. The calcium ionophore A23187 transports calcium ions across a cell membrane, causing a sudden increase in free calcium ion levels within the cell. What do you think the effect would be of adding A23187 to unfertilized eggs? Explain your reasoning. (For experiments, 1 mg A23187 is dissolved in 1 ml of dimethyl sulfoxide—DMSO—to make a stock solution. This can be stored in the freezer. The stock solution is diluted 1:100 in seawater for use. All operations should be done in a hood, wearing gloves, and using bench paper that is discarded after use. DMSO passes freely through the skin and will carry along with it anything that it is attached to.) If you are following these cultures beyond class period, do not leave them in sodium-free seawater or DMSO solution. Return the embryos to normal seawater. You can do this by swirling the dish to concentrate the embryos. Then pipette them into fresh seawater. Do this several times to wash them clean of their previous solutions. Store them at a temperature appropriate for the species being used.

Parthenogenesis Parthenogenesis, the development of an egg without fertilization, can be induced in the sea urchin in a number of ways. Remember, all that has to be achieved to activate the egg is an assault that will cause a rise in pH. (This is caused by activating the PIP cycle in normal fertilization.) In the past, numerous methods have been tried, often successfully. Hypertonic salt solutions as well as hypotonic solutions have worked, as have a pricking of the cell membrane and even violent shaking of the eggs. No one knows yet exactly how each method works, but it would be reasonable to suppose that each causes an activation of the PIP cycle. We will use aged eggs (4–6 hours) since these often work better than freshly spawned eggs. Why do you think this is the case? Enter your answer in your notebook. Try putting several milliliters of an aged-egg solution in a glass vial and shaking it in your hand. Also try exposing the eggs for 10 minutes to a hypertonic solution of seawater: 2.2 ml of 2.5 M (14.6%) NaCl in 7.8 ml of normal seawater. The hypertonic method was developed by Ernest Everett Just in the early 1900s. (E. E. Just [1883–1941] was an eminent embryologist, a pioneer in the study of fertilization, and an African-American who suffered discrimination in this country. Unable to find a supportive atmosphere at home, he fled to Europe to pursue his work. Forced to return by the outbreak of World War II, he came home a sick man, and died, too young, of a painful intestinal neuralgia and pancreatic cancer. He’s the only embryologist ever to be honored on a U.S. postage stamp [issued Feb., 1996].)

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View the treated eggs under a dissecting scope. What are you looking for that will indicate activation of the eggs? Many times, you will see the elevation of the fertilization envelope. What does this tell you? Under conditions of normal fertilization, cleavage occurs within the first hour at room temperature. Leave some of your eggs at room temperature and place some in the refrigerator. These can be checked at intervals. If cleavage does occur (congratulations!), your eggs will be among the lucky Fertilization envelope

Fertilization cone

Unfertilized egg

Fertilized egg

Fertilization envelope formed

Two-cell stage

Four-cell stage

Eight-cell stage

Blastocoel

Micromere Sixteen-cell stage

Primary mesenchyme

Early gastrula

Blastula

Morula

Gut invaginating

Mid-gastrula

Secondary mesenchyme

Late gastrula

Stomodeum Gut

Figure 6.1 Developmental sequence of the sea urchin embryo from fertilization to the pluteus larval stage.

Spicules

Prism larva stage

Pluteus larva

Anus

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1% that can be expected to be parthenogenetically activated. Even those that are activated, however, rarely make it past the blastula stage. Why? Suggest an answer and enter it in your notebook. If you plan on keeping these cultures beyond the class period, again, it is important that you remove them to normal seawater. Store them at a temperature appropriate for the species being used.

Cleavage, Gastrulation, and Larval Stages The first several cleavages can be observed during the laboratory period. But you will have to use your sterile cultures to observe the later stages of development. The stages and pattern of cleavage can be seen in Figure 6.1. Cleavage is holoblastic (the entire egg cleaves) and radial (the cleavage planes are parallel or at right angles to the animal-vegetal pole). The first cleavage takes about an hour at room temperature, and subsequent cleavages occur about every half hour. At the 16-cell stage, a small group of micromeres are cleaved at the vegetal pole. These are the primary mesenchyme cells and will be the first to show gastrulation movements. At 5–6 hours, the embryo is at the blastula stage, and by 7–8 hours, the embryo has hatched out of its fertilization envelope and is spinning around the dish using its cilia for locomotion. This is called a hatched blastula. Gastrulation begins at about the hatched blastula stage. Primary mesenchyme cells first migrate into the blastocoel and then form a necklace of cells that will secrete the skeletal supports for the larva, the spicules. The spicules are first tripartite rods, and they eventually branch to have several arms. Gastrulation continues by invagination of the vegetal plate to form the archenteron (meaning “ancient gut”). The forming gut will elongate, capped by a loose collection of cells, the secondary mesenchyme. The secondary mesenchyme aid in elongating the gut with their contractile filopodia, pulling the archenteron toward the far wall of the blastocoel and guiding it to its final destination. They later disperse to form mesodermal organs. As the gut is developing, the embryo goes through a prism larval stage (between 18–20 hours), looking like an exquisite rotating jewel, and finally becomes a pluteus (echinopluteus) larva (at about 22–24 hours). The pluteus is an ornate organism, projecting long, delicate arms supported by branched spicules. The timing of development will vary considerably with temperature. You can keep cultures in the refrigerator, at room temperature, or at any other temperatures the laboratory can supply. As soon as you have swimming blastulae, I suggest catching these in a sterile pipette and transferring them to a fresh culture dish with sterile seawater. This will avoid bacterial contamination from the decomposing embryos that didn’t make it. Make diagrams of any stages you are able to observe. If you are lucky enough to get larvae, be sure to look at them under the microscope using polarized light. As the larvae rotate, their spicules briefly align with the polarizers with each turn, flashing beautifully for you. Compare the rates of development at the different temperatures you used by presenting your data in chart and graph form. Notice also (from the cultures that don’t make it) that you are establishing viable temperature ranges for the species you are using. You can see from Table 6.2 that complete timetables of sea urchin or sand dollar development are hard to come by. If you develop a complete timetable for one or more temperatures, this is publishable work.

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Table 6.2 Developmental timetables for sea urchin and sand dollar embryos at various temperatures Strongylo- EchinaStrongylocentrotus centrotus rachnius droe ba c hie ns is purpura t us pa rma Arba c ia punc t ula t a ___________________ __________ _________ ___________________

S tage Fertilization 2-Cell stage 4-Cell stage 8-Cell stage 16-Cell stage 32-Cell stage Morula Early blastula Mid-blastula Hatching blastula Early gastrula Mid-gastrula Late gastrula Prism stage Early pluteus Late pluteus Metamorphosis

4 °C

8 °C

1 0 °C

1 5 °C

2 0 °C

2 3 °C

0 5 hrs 8 hrs 10.5 hrs 14 hrs 18 hrs

0 3 hrs 5 hrs 6.5 hrs 8.5 hrs 11 hrs

0 3.5 hrs 5 hrs 6 hrs 8.5 hrs

0 1.5 hrs 3 hrs 4 hrs 5.5 hrs

0 1 hrs 1.8 hrs 2.4 hrs

0 50 min 78 min 1.71 hrs 2.25 hrs 2.78 hrs 4 hrs 4.5 hrs 6 hrs 7–8 hrs 12–15 hrs

50 hrs

6.5 days 11 days

30 hrs

27–28 hrs

4 days 7 days

50 hrs 57 hrs 80 hrs 4.7 days

7–22 wks

63–86 days

6.5 hrs 7 hrs 8 hrs 12 hrs 25 hrs 31 hrs 48 hrs 50 hrs 72 hrs

6 hrs 10 hrs

48 hrs

17 hrs 18 hrs 20 hrs 24 hrs 5–16 wks

Sources: S. droebachiensis and S. purpuratus after Strathmann, 1987; E. parma after Karnofsky and Simmel, 1963; A. punctulata at 20°C after Costello et al., 1957; A. punctulata at 23°C after Harvey, 1956.

Accompanying Materials See Vade Mecum: “Sea Urchin.” This chapter of the CD provides video of the living embryo, with details on the raising of the fertilization envelope, color-coding of germ layers, and an artistic rendition of metamorphosis based on Ethel B. Harvey’s original photographs. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. You will find excellent discussions of fertilization, the PIP cycle, and development of the sea urchin to the pluteus larval stage. Fink, R. (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Sequence 1. This shows sea urchin development from fertilization through spicule formation.

Selected Bibliography Carroll, D. J., D. Albay, M. Terasaki, L. A. Jaffe and K. R. Foltz. 1999. Identification of PLC γdependent and -independent events during fertilization of sea urchin eggs. Dev. Biol. 206: 232–247. This paper sorts out which events of fertilization are caused by the rise in calcium within the egg due to activation of the PIP cycle. These include the cortical granule reaction, the intracellular rise in pH, MAP kinase dephosphorylation, DNA synthesis, and the onset of cleavage.

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Costello, D. P., M. E. Davidson, A. Eggers, M. H. Fox and C. Henley. 1957. Methods for Obtaining and Handling Marine Eggs and Embryos. Marine Biological Laboratory, Woods Hole, MA. This superb book, now out of print, is worth its weight in platinum to any marine invertebrate embryologist. For each species covered, descriptions are given of where and when to collect the animals, how to obtain gametes, and how to culture the embryos. Czihak, G. (ed.). 1975. The Sea Urchin Embryo: Biochemistry and Morphogenesis. Springer-Verlag, Berlin. This is now a classic, a rich collection of papers on sea urchin development. Ernst, S. G. 1997. A century of sea urchin development. Amer. Zool. 37: 250–259. A beautiful summary of the early research on sea urchins, including a short analysis of some of E. E. Just’s contributions. Epel, D. 1977. The program of fertilization. Sci. Amer. 237(5): 128–138. This article, though now somewhat dated, is a beautifully illustrated, clearly written explanation of fertilization in sea urchins. It is well worth getting from Scientific American Offprints. Foltz, K. R., 1995. Gamete recognition and egg activation in sea urchins. Amer. Zool. 35: 381–390. This is an excellent summary of fertilization in sea urchins. Harvey, E. B. 1956. The American Arbacia and Other Sea Urchins. Princeton University Press, Princeton, NJ. This tome, written by Ethel Harvey at a time when few women were allowed to succeed in science, is an extremely precise, exhaustive compilation of information on Arbacia. Her photographs of development are among the most complete to be had. Karnofsky, D. A. and E. B. Simmel. 1963. Effects of growth inhibiting chemicals on the sand dollar embryos Echinarachnius parma. Prog. Exp. Tumor Res. 3: 254 –295. An excellent description of sand dollar development, this paper, found unexpectedly in a journal on tumor research, illustrates how grant money for medical research still can be used for studying normal development. Manning, K. 1983. Black Apollo of Science: The Life of Ernest Everett Just. Oxford University Press, New York. A compassionate biography of an outstanding embryologist. It is E. E. Just who wrote, “We feel the beauty of Nature because we are part of Nature and because we know that however much in our separate domains we abstract from the unity of Nature, this unity remains.” This biography of Just reminds us that scientists should always see the beauty of the whole that binds together the parts that we dissect. Pearse, J. S. and R. A. Cameron. 1991. Echinodermata: Echinoidea. In A. C. Giese, J. S. Pearse and V. B. Pearse (eds.), Reproduction of Marine Invertebrates. Volume VI: Echinoderms and Lophophorates. Boxwood Press, Pacific Grove, CA, pp. 513–662. This series on invertebrate reproduction is an astonishing collection of information. The chapter on echinoids gives tables of spawning times for a long list of species and provides an excellent summary echinoid development. Sherwood, D. R. and D. R. McClay. 1999. LvNotch signaling mediates secondary mesenchyme specification in the sea urchin embryo. Development 126: 1703–1713. The work of David McClay and his collaborators is an excellent example of how to combine old-fashioned observation with modern molecular techniques. Their work has made significant advances in our understanding of gastrulation. This paper summarizes a portion of that work. Strathmann, M. F. 1987. Reproduction and Development of Marine Invertebrates of the Northern Pacific Coast: Data and Methods for the Study of Eggs, Embryos, and Larvae. University of Washington Press, Seattle. This extremely useful volume is the modern version for the Pacific coast of the classic volume for the Atlantic coast by Costello et al. (cited above). Stricker, S. A. 1999. Comparative biology of calcium signaling during fertilization and egg activation in animals. Dev. Bio. 211: 157–176. This gives a concise review of what is known about the role of calcium in fertilization in all groups of metazoans studied so far.

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Wessel, G. M. and A. Wikramanayake. 1999. How to grow a gut: ontogeny of the endoderm in the sea urchin embryo. BioEssays 21: 459–471. A well-written summary of gastrulation movements in the sea urchin and the mechanisms of these movements, this review also covers a great deal of the history of this field. Wilt, F. H. and N. K. Wessells (eds.). 1967. Methods in Developmental Biology. Thomas Y. Crowell, New York. This is one of my favorite techniques books in developmental biology. For every species covered, it has a wealth of information in a compact space. If you ever see it on a “for sale” list, don’t let it slip away. Wray, G. A. and D. R. McClay. 1989. Molecular heterochronies and heterotopies in early echinoid development. Evolution 43: 803–813. The study of evolution and embryology should be inexorably linked but seldom are. This concise paper shows how the study of development can advance our understanding of evolution. Vacquier V. D. and G. W. Moy. 1997. The fucose sulfate polymer of egg jelly binds to sperm REJ and is the inducer of the sea urchin sperm acrosome reaction. Dev. Biol. 192: 125–135. This paper discusses methods for dejellying sea urchin eggs in addition to reconfirming that a fucosecontaining polysaccharide activates the acrosome reaction in the sea urchin sperm.

Suppliers Connecticut Valley Biological Supply Co., Inc. P.O. Box 326 82 Valley Road Southampton, MA 01073 1-800-628-7748 Stongylocentrotus droebachiensis Instant ocean

Gulf Specimen Marine Laboratories, Inc. P.O. Box 237 Panacea, FL 32346 1-850-984-5297 www.gulfspecimen.org Lytechinus variegatus Arbacia punctulata (June-August)

Marine Biological Laboratory Woods Hole, MA 02543 1-508-548-3705, ext. 325 www.mbl.edu Stongylocentrotus droebachiensis (November–March) Arbacia punctulata (mid June–mid August) Echinarachnius parma

Echinoid Fertilization and Development

Pacific Bio-Marine Laboratories, Inc. P.O. Box 536 Venice, CA 90291 1-310-677-1056

Sea Life Supply 740 Tioga Ave. Sand City, CA 93933-3034 1-831-394-0828 www.sealifesupply.com Stongylocentrotus purpuratus (late November–June) Stongylocentrotus fransiscanus (late November– June) Lytichinus pictus (May–September)

Any good biological supply company, such as: Fisher Scientific 585 Alpha Dr. Pittsburgh, PA 15238 1-800-766-7000 www.fishersci.com Salts for artificial and Ca/Mg-free seawater Sterilizing filters (0.22-µm pore size) Sterile filter systems (disposable and reusable)

Any good chemical supply company, such as: Sigma Chemical Company P.O. Box 14508 St. Louis, MO 63178 1-800-325-3010 www.sigma-aldrich.com DMSO Calcium ionophore A23187 Streptomycin

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Glossary Acrosomal process: A process that forms in certain types of sperm, such as that of sea urchin, during the fertilization reaction. This long, thin process forms through the inversion of the acrosomal vesicle, exposing the specific molecules embedded in the acrosomal membrane that can then bind to the egg surface. Acrosomal vesicle: A membrane-bound sac of enzymes in the tip of the spermatozoan above the nucleus. The sac opens during fertilization, releasing enzymes that coat the head of the sperm and break down the various extracellular layers surrounding the egg. Acrosome reaction: The events in the head of a sperm that opens the acrosomal vesicle, releasing its contents to the outside. The acrosome reaction occurs when the sperm hits the outer extracellular coats surrounding the egg. Components in these coats set off the acrosome reaction. Actin microfilaments: The thin microfilaments found in the cytoplasm of a cell. Made up of G-actin, the globular protein subunits. The filamentous form of actin is called F-actin. F-actin forms an extensive cytoskeletal network in the cortical cytoplasm. Myosin (the thicker of the two types of microfilaments) can move along F-actin, creating a contractile force. Animal pole: The point on the surface of the egg where polar bodies are given off. Eggs usually orient to gravity with their animal pole oriented uppermost. Archenteron: Literally, ancient gut. The primitive gut formed during gastrulation. Aristotle’s lantern: The mouth structures in the sea urchin and other echinoids. The complex includes five calcareous teeth used for grazing and chopping up food. These can be seen on the oral surface of the animal. Artificial seawater: A salt solution that can be used instead of sea water. It includes the major salts found in sea water, sodium chloride, potassium chloride, magnesium chloride, magnesium sulfate, and calcium chloride. Bindin: A cell surface protein on the acrosomal process of a sea urchin sperm that allows speciesspecific binding to the surface of the sea urchin egg. Blastocoel: The cavity within a blastula. Blastula stage: The stage in embryonic development following cleavage and preceding gastrulation. The embryo may form a hollow blastula (coeloblastula) or a solid blastula (stereoblastula). The cavity formed in a coeloblastula is called the blastocoel. Calcium ionophore: A small hydrophobic molecule that dissolves in lipid bilayers and increases their permeability to calcium allowing calcium to move into the cell. (A23187 is a calcium ionophore.) Cortical granule reaction: The reaction in the cytoplasm of an egg upon fertilization in which the cortical granules of the egg fuse with the egg cell membrane, releasing their contents to the exterior. The extracellular matrix around the egg is modified by these contents such that further sperm are unable to get to the egg cell surface. Cortical granules: Vesicles found in the cortical cytoplasm of an unfertilized egg. These contain components which are released to the exterior upon fertilization or activation of the egg. These components modify the extracellular substances surrounding the egg such that sperm are no longer able to approach the egg cell surface. Dry sperm: Sperm that have been shed and are kept in this concentrated form, undiluted. In experimental procedures, it is often useful to store sperm as dry sperm, since they survive for longer periods in this form. Echinoid: The group of echinoderms that include sea urchins, sand dollars, and heart urchins.

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Echinopluteus (pluteus) larva: The type of larva typical of echinoids (sea urchins, etc). It consists of a delicate body with long arm-like extensions supported by spicules. Ectoderm: The germ layer in an embryo that gives rise to the epidermis and nervous system. Endoderm: The germ layer in an embryo that gives rise to the epithelium (lining) of the gut and gut derivatives. Fast block to polyspermy: Well studied in the sea urchin, it is a reaction set off in the egg when the sperm makes contact with the egg. The membrane potential of the egg cell membrane changes from –70 mV to above 0 mV. This prevents further sperm from binding. This happens rapidly, taking only 1/10th of a second to be complete, but lasts for only a short period (approximately 1 minute). Fertilization: Fusion of egg and sperm. Fertilization cone: In certain eggs such as the sea urchin, a structure that forms during fertilization at the point of sperm entry. Microvilli elongate and wrap themselves around the entering sperm. These appear as a small cone-shaped structure on the surface of the egg. Fertilization envelope: In the sea urchin egg, the vitelline envelope following fertilization. The vitelline envelope lifts off the surface of the egg and is toughened by components released by the cortical granulate reaction. Filopodium: A pseudopodium that is long and thin. It is filled with microfilaments and is contractile. When a filopodium contracts, it either loses its connection to the substrate, collapsing back into the cell, or it causes the cell body to detach and pulls the cell body toward the filopodium’s attachment site. Footed coverslip: A coverslip that has a small spacer on each corner; one can be made by nicking the corners of the coverslip against some hard paraffin, so that crumbs of paraffin remain attached at each corner. This can give just enough spacer between the slide and the coverslip to avoid crushing the specimen on the slide. Gastrulation: The stage in embryonic development that follows cleavage. During gastrulation, cells rearrange themselves, the endoderm and mesoderm cells moving inward and the ectoderm cells spreading around the outside, with some ectoderm invaginating to form such structures as the stomodeum. Germinal vesicle: The nucleus of a primary oocyte. At this stage, the oocyte is diploid and growing in size, transcribing genes to make mRNA that will be stored in the cytoplasm as maternal mRNA, and transcribing ribosomal genes to make numerous ribosomes. The nucleus at this stage is large. Glycoprotein: A molecule containing protein bound to carbohydrates. Hatched blastula: A term used to describe the echinoid (sea urchin, etc.) embryo when it has hatched out of its fertilization envelope. This stage occurs early in development, just after primary mesenchyme cells have started to ingress. Holoblastic cleavage: Complete cleavage. The pattern of cleavage in which the entire egg is divided into smaller cells. Hypertonic salt solution: A salt solution that is more concentrated than cytoplasm. Hypotonic salt solution: A salt solution that is more dilute than cytoplasm. Invagination: The folding inward of a sheet of cells. This is a common type of movement during gastrulation. In the sea urchin, for example, the archenteron first forms by an invagination of the vegetal plate. Mesoderm: The germ layer in an embryo that gives rise to the muscular and circulatory systems, and most of the skeletal and urogenital systems. Micromeres: Cells in a blastula that are smaller than the others. In the sea urchin embryo, micromeres form at the vegetal pole at the 16-cell stage. These will form the primary mesenchyme.

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Ootid: A developing ovum in the gonotid stage. An ootid is haploid, with each of its chromosomes consisting of a single chromatid. The second meiotic division divides the secondary oocyte asymmetrically, producing one ootid and one tiny polar body. Parthenogenesis: The development of an egg without fertilization. Peristomial membrane: The tough, leathery membrane surrounding the mouth parts of sea urchins and other echinoids. Perivisceral cavity: The coelomic cavity (or hemocoel) in which the viscera lie. Perivitelline space: The space between the egg cell membrane (or embryo) and the vitelline envelope that surrounds the egg (or embryo). Phosphatidylinositol bisphosphate (PIP) cycle: A signal transduction pathway that is initiated by the splitting of phosphatidylinositol bisphosphate (PIP2), a phospholipid within the cell membrane. PIP2 splits into inositol triphosphate (IP3) and diacylglycerol (DAG). IP3 causes the release of calcium ions from intracellular stores, raising cytoplasmic levels of calcium. DAG is involved with activating an ion pump that moves hydrogen ions out of the cell and sodium ions into the cell, raising the intracellular pH. PIP cycle: Phosphatidylinositol bisphosphate cycle. Pluteus larva: The type of larva typical of echinoderms. It consists of a delicate body with long armlike extensions supported by spicules. Polarizer: A filter on a microscope that transmits light vibrating only in one direction—all other rays are excluded. Polypeptide: A molecule made up of a chain of amino acids. Polysaccharide: A carbohydrate made up of simple sugars. Primary mesenchyme: Mesoderm cells in echinoid embryos (such as those of the sea urchin) derived from the micromeres. The primary mesenchyme cells are the first cells to enter the blastocoel during gastrulation. They form the spicules of the larva. Primary oocyte: A developing ovum in the primary gonocyte stage, a stage during which the cell is growing and not dividing. At the end of this stage, the cell enters the first meiotic division. Prism larval stage: In an echinoid (sea urchin, etc.) larva, the stage at which the larva is shaped like a prism. It is an early stage in larval development. Radial cleavage: The cleavage pattern in which each cleavage plane is either parallel or perpendicular to the animal-vegetal axis. Secondary mesenchyme: Mesoderm cells in the echinoid (sea urchin, etc.) embryo that invaginate along with the endoderm to form the archenteron. Situated at the anterior end of the invaginated archenteron, they secondarily separate off to form the coelomic sacs and mesodermal organs. Slow block to polyspermy: Well studied in the sea urchin, it is a reaction set off in the egg when the sperm makes contact with the egg. The cortical granules of the egg fuse with the egg cell membrane and empty their contents to the exterior. Components of the cortical granules change the surrounding extracellular components such that sperm are no longer able to get close to the egg cell surface. This takes about a minute to complete and is a permanent block to further sperm penetration. Sodium-free seawater: A salt solution that has the same ionic concentration as seawater but that does not contain any sodium salts (NaCl). This can be used in experiments to test the requirement for external sodium. Spawning: The release of gametes (eggs or sperm). Spicules: In the echinoid (sea urchin, etc.) larva, these are the calcium carbonate structures that serve as the skeletal supports for the larva. The are laid down by the primary mesenchyme cells.

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Standard sperm suspension: The diluted form of sperm that is used for fertilization studies. In the sea urchin, for example, this is 1 drop of dry sperm in 10 ml of seawater. Vegetal pole: The pole that is opposite the animal pole. Yolk is more concentrated at the vegetal pole. Eggs usually orient to gravity such that their vegetal pole is oriented downward. Vitelline envelope: An extracellular coat surrounding an ovum.

Sea Urchin Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. What do you inject a sea urchin with to get it to spawn, and how can you tell the spawn of male sea urchins from that of females?

2. How can you tell a mature sea urchin egg from an immature egg?

3. Why is it important to leave sea urchin sperm undiluted until used?

4. What is the fertilization envelope, and how is it formed?

5. What is the function of the fertilization envelope?

6. What is the cleavage pattern of sea urchin eggs?

Name: _______________________

7. What do the micromeres of the sea urchin embryo become?

8. What methods of parthenogenesis were explored by E. E. Just?

9. What happens during the first stage in sea urchin gastrulation?

10. What is the function of the larval spicules?

Sea Urchin - 2

chapter

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Sea Urchin Development

Effects of Ultraviolet Radiation

1

Ultraviolet Radiation in Earth’s Atmosphere What could be more enticing than a sun-drenched beach? Yet that inviting sunlight harbors great potential harm for us, as well as any organism exposed to the damaging ultraviolet rays that pass through our atmosphere. It has been known for some time that ultraviolet light is damaging, and indeed you may have already used it in the laboratory as a germicidal agent. Ultraviolet light kills—and it does so primarily by damaging DNA. So how does any organism survive exposure to ultraviolet light? And how much of a threat are the increasing levels of ultraviolet radiation that are reaching Earth’s surface because of the damage we have caused to the protective atmospheric ozone layer? The damage caused by ultraviolet (UV) radiation depends on its wavelength (Figure 7.1). UV wavelengths fall between 200 and 400 nanometers (nm). Above this is visible light, also called PAR (photosynthetically active radiation; 400 –700 nm) that is not considered harmful. Within the UV range, the longest wavelengths, which constitute UV-A (320– 400 nm), are the least harmful to biological systems. They are also the type of UV radiation that passes most easily through Earth’s atmosphere. Only about 3% of the sunlight reaching Earth’s surface is in the ultraviolet range, and most of this is in the UV-A range. UV-B (280–320 nm; though some sources use 290–320 nm) is significantly more harmful, but most of the UV-B coming from the sun (especially wavelengths below 295 nm) is blocked by the ozone in Earth’s upper atmosphere (the stratosphere). Only about 0.25% of the sun’s light reaching Earth’s surface is UV-B. UV-C (200–280 nm) is the most dangerous to organisms; however, because of its absorption by oxygen in the atmosphere, UV-C never reaches Earth’s surface. It is UV-B radiation, therefore, that is of primary concern to biological organisms, and anything that would shift the absorption of UV-B to allow more of this wavelength to reach the Earth’s surface should be of major concern to us. This is precisely what has been happening due to a reduction in the protective ozone shield of the upper atmosphere. This shield has been reduced in recent years by the production of chlorofluorocarbons (CFCs; these include freon, chloroform, carbon tetrachloride, various cleaning agents of computer circuit boards, and propellants for aerosol cans). CFCs carry chlorine into the upper atmosphere where the chlorine destroys ozone. The increase in CFCs has created such severe ozone depletion that as much as 5% of the ozone shield has disappeared in less than a 10-year period. Even if we were to completely stop the production of CFCs today, it would still take 30 years before the CFCs that are presently in the lower atmosphere finally worked their way into the upper atmosphere. And it would take 70 years beyond that before the ozone levels were restored just to

1

With thanks to Nikki Adams, Department of Biology, California Polytechnic State University, for her help in developing this chapter.

Sea Urchin-UV Effects

UV-C 200 nm

UV-B 280 nm

UV-A

320 nm

400 nm

Blocked by

Most Harmful

atmospheric O2 More Harmful

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Figure 7.1 The damage caused by ultraviolet radiation depends on its wavelength. UV-C (200-280 nm) is the most harmful, but it doesn’t reach Earth’s surface, since it is blocked by atmospheric oxygen. UV-B (280-320 nm) is more harmful than UV-A (320-400 nm).

Least Harmful

present-day levels. Major efforts toward the elimination of CFCs were made in 1987 when 24 countries signed the Montreal Protocol, and additional countries have signed since then. Even so, there are major ozone “holes” that have developed in the upper atmosphere, and these grow and shrink in size with seasonal regularity, becoming their largest in the winter months. Ozone depletion is a global problem, although it is most severe over Antarctica where one ozone “hole” at its largest is the size of the United States. The effects of this ozone depletion are likely to be most devastating to organisms that are exposed to sunlight during their developmental stages. These include many freshwater and marine species, as well as terrestrial species such as amphibians that return to water for reproduction. Although it used to be thought that UV-B couldn’t penetrate into water very far, it has since been learned that UV-B radiation can reach to depths of 20 meters, and UV-A radiation can penetrate even farther. So, many of the organisms that we used to think were protected from increased UV exposure instead are found to be threatened.

Biological Effects of UV Radiation The damage caused by UV radiation that is environmentally relevant includes that caused by UV-A and UV-B. On an organismal level, this damage results in inhibition of photosynthesis, sunburn, photoaging of the skin, skin cancer, cataracts, damage to the neural retina, and depression of the immune system. On a cellular level, the damage can cause the alteration of DNA, proteins, and lipids, either through direct or indirect effects. UV-A radiation causes damage indirectly by creating reactive oxygen species, such as superoxide -

anions (O2 ) and hydrogen peroxide (H2O2). These highly reactive molecules cause damage by oxidizing proteins, lipids, and DNA. This oxidation can cause functional changes in enzymes and structural

Sea Urchin-UV Effects

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elements of the cell, and it can result in mutations through alteration of the DNA, which can ultimately lead to cancer. UV-B radiation causes direct damage to proteins and DNA by creating the formation of pyrimidine dimers and crosslinks between DNA and protein. This form of radiation causes significant DNA damage, delayed cell divisions, and is responsible for most of the ill effects listed above as the effects of UV light on organisms. Researchers are beginning to investigate the extent of the effects of increased UV radiation caused by the thinning of our protective atmospheric ozone layer. Already, it has been shown that increased levels of UV-B radiation can be particularly damaging to gametes and embryos. Any organism whose gametes or embryos are exposed to sunlight are likely to be susceptible and at risk.

Mechanims That Protect Against Damage from UV Organisms are not completely unprotected against the damage caused by UV light, and some have greater amounts of protection than others. Protective mechanisms include natural sunscreens, such as blue pigments, which reflect short-wavelength radiation, and pigments and other compounds that absorb UV-B, such as mycosporine-like amino acids (MAAs). MAAs are found in photosynthetic algae, fungi, and bacteria, and they can accumulate in non-photosynthetic organisms that feed on the MAA-rich organisms. Physiological mechanisms also exist for repairing the DNA damage inflicted by UV light; these rely on DNA repair enzymes such as photolyase. These enzymes repair the UV-induced DNA dimers. Interestingly, these are photorepair mechanisms, being activated by light in the visible range and down into the UV-A range. Eggs and embryos can vary in the amounts of photolyase they contain, and these variations can be species-specific. In several amphibian species from the western United States, the species that showed the most deformities induced by UV-B radiation exposure were also the species with the lesser amounts of photolyase (Blaustein et al., 1994; Blaustein et al., 1998).

Using Sea Urchins as a Model Organism for UV Studies Sea urchins can be used to study damage induced by ultraviolet radiation. They are exposed in nature to UV radiation during all stages of their life cycle, and any analysis of their sensitivity can be used as a measure of the extent of the threat that increased UV-B exposure poses. Although the adults would seem to be well protected from UV radiation by their heavily calcified tests, their gametes and embryos are transparent and poorly protected. It has long been known that UV radiation is harmful to sea urchin development. As early as 1877 (Downes and Blunt), UV radiation was shown to induce mitotic delay in cells. These results were verified and and shown to apply to sea urchin embryos as well (e.g., Giese, 1946; Rustad, 1971). It has also been shown that UV radiation has adverse effects on sperm, causing sperm to agglutinate, be less motile, and have reduced fertilizing abilities. In most of these earlier studies, however, the emphasis was not on the threats of environmental exposures to UV radiation, and the wavelengths used were often in the UV-C range, a range that isn’t environmentally relevant. More recently, however, the ill effects of UV light in environmentally relevant ranges have been documented (e.g., Adams and Shick, 1996; Anderson et al., 1993). These studies show that environmental UV light can cause cleavage delay and a number of developmental abnormalities, such as blastulae that are filled with an abnormally large number of cells, exogastrulae, and bizarrely shaped pluteus larvae with extra spicules. The

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embryos have proven to be most susceptible to damage when they are exposed during early stages (up to the formation of the blastula) rather than during later stages. The embryos are not without some protection against UV radiation. Adams and Shick have documented that the accumulation of natural sunscreens (the MAAs discussed earlier) protect the eggs and embryos. These MAAs, found in the algae that is consumed by the adult animal, are stored in the eggs, providing a certain level of protection, depending upon the amounts accumulated. Therefore, by selecting algae high in MAAs, an adult animal can influence the protection from UV radiation that it provides the next generation.

Setting Up Your Experiments The questions you can ask of sea urchin gametes and embryos about the effects of UV are limited only by your imagination. • Is the sperm or the egg more susceptible to UV radiation? • What stages of development are most susceptible to damage by UV radiation? • Will exposure to differing lengths of daylight affect development? • Is UV-B or UV-A radiation more harmful to these embryos? • Is the combined effect of UV-B and UV-A radiation exposure greater than the effect of exposure to only one of these ranges? • Do antioxidants, such as vitamin C, counteract the effects of UV-A radiation and thus reduce the formation of reactive oxygen species? • Does exposure to just PAR (visible light) cause any defects? • Does exposure to PAR mitigate any of the ill effects caused by UV radiation exposure? • Do the screens for filtering out UV light that we rely on to protect us—such as sunglasses and sunscreen lotions for our skin—actually work? In getting started, you will want to refer back to Chapter 6, “Echinoid Fertilization and Development,” to review how to obtain gametes from the sea urchin, and how to store these gametes until you are ready to use them. Remember that the eggs should be washed several times in seawater to remove any perivisceral fluid and that the sperm should be stored as “dry sperm.” Eggs should be maintained at temperatures appropriate for the species. Sperm can be stored in the refrigerator. You should be using these gametes as soon after spawning as possible. Remember that aged eggs (4 hours old or older) are more susceptible to parthenogenesis. In preparing your cultures, it is best to use artificial or filtered seawater, if possible. This avoids the inclusion of dissolved organic material, which can become oxidized when exposed to UV light, forming reactive oxygen species such as hydrogen peroxide. You can set up your cultures in petri dishes. You should put no more eggs in a dish than will form a monolayer, since you want to achieve equal exposure to the light conditions. To keep sperm from using up their energy with active swimming, they can be maintained as a thin layer of dry sperm on ice or as diluted sperm in cold (4˚C) seawater. Cultures will have to be kept covered so that they don’t lose water to the environment. The cover you choose will depend on the type of light you want to let through. If you want to test the effects of UV-A and UV-B that are occurring naturally, you can set your cultures up outside to use natural sunlight. Alternatively, you can use a light that mimics the range of light in nature. To test the effects of UV-A and UV-B separately, you can use filters that block certain

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wavelengths of light, or you can use bulbs that emit only in certain wavelengths. These bulbs are discussed below. In setting up your exposures, you can use a short exposure period (such as 30 minutes) followed either by dark conditions (to prevent photorepair mechanisms from working) or by visible light conditions that will allow for photorepair. Exposures also can be set to mimic natural light-dark cycles. Using sunlight will automatically give you a natural light-dark cycle. Alternatively, you can choose constant exposure, although you should be aware that some of these exposures could be exceedingly harmful to your embryos and may kill them. Whatever your setup, remember that you should be keeping your gametes and embryos at a temperature that promotes development. You would have determined this temperature during your previous laboratory on sea urchin development. In many cases, this means that you have to keep your cultures cold. If using natural sunlight, you may need to keep your cultures sitting on ice that you replenish as needed. If using artificial light, again, maintain cultures on ice if needed, or preferably in a cold room or temperature-controlled water bath. Special lightbulbs that can be used Everyday room lights Room lighting provides light in the PAR (visible) range. An incandescent bulb emits negligible levels of UV light. Fluorescent bulbs, however, emit measurable levels of UV-B light as well as PAR, and this factor should be considered in your experiments. (You could even test to see if there were any differences between exposure to a fluorescent bulb versus an incandescent bulb.) An unfiltered halogen bulb emits a continuous spectrum that reaches all the way into the UVC range. Grow lights A grow light, a light commonly used to enhance growth of house-grown plants, provides light enhanced in the UV-A range without emitting light in the UV-B range. Sunlamps A sunlamp, such as those sold for home use (e.g., Sylvania® FS20 sunlamp), provides light enhanced in the UV-B range. Sunlamps from 20 years ago (275 watts) emit high levels of both UV-A and UV-B radiation. Black lights A black light, a common accessory to youthful parties (e.g., NEC T10, 20 watts), provides UV-A light. UV 340 lamps A UV 340 lightbulb supplies light in both the UV-A and UV-B ranges and mimics the light found in nature. These bulbs are commonly used to test paints for weathering in sunlight. Germicidal lamps A germicidal UV bulb emits light enhanced in the UV-C range. It is therefore not environmentally relevant, but it can be used to show the detrimental effects of this exceedingly harmful radiation. Filters that can be used to exclude UV-A or UV-B Whether you are using natural sunlight or a UV 340 bulb, you can test the differing effects of UV-A and UV-B light by filtering out one or the other using various plastics and glass that block specific wavelengths of light. These sheets can be placed over your culture dishes. Alternatively, if you are working out in the field with organisms in their natural environment, you can use containers or bags made from the plastic or glass. You can place the embryos in the containers or bags and suspend them in the water where they are normally found.

Sea Urchin-UV Effects Filters that block UV-B

Allows through UV-A and PAR

glass Xerox transparencies plastic milk bottle Mylar film

Filters that block UV-A and UV-B (UV opaque)

Allows through PAR

standard plexiglass “light-block” plastic milk bottle sunglasses (rated as 100% UV blocking) sunscreens (variable) UV opaque plexiglass

UV transparent filters

Allows through UV-A, UV-B, and PAR

plastic petri dish plastic sandwich bag plastic food wrap UV transparent plexiglass cellulose acetate

7-6

Filters that block UV-B These will test the effects of exposure to UV-A and PAR. Glass Plate glass blocks almost all UV-B light while allowing through most UV-A light and PAR. For example, 1/8-inch plate glass blocks all UV-B while allowing through 76% of the UV-A light and PAR. A glass slide blocks all of the UV-B while allowing through 85% of the UV-A light and PAR. A finger bowl blocks all of the UV-B while allowing through 61% of the UV-A and PAR. (N.B.: Pyrex glass petri dishes do not block UV-B effectively.) Xerox transparencies The transparency films that are used for copier machines block 95% of UV-B while allowing through 77% of UV-A and PAR. Plastic milk bottle An ordinary plastic milk bottle blocks all UV-B light while allowing through 55% of the UV-A and 80% of PAR. Mylar A special type of Mylar film (Mylar type D fluoropolymer film) blocks all UV-B radiation. Filters that block both UV-A and UV-B (UV opaque) These will test the effects of exposure only to PAR (visible light). Standard plexiglass Standard plexiglass blocks all UV-B and most UV-A radiation while allowing PAR through. “Light-block” plastic milk bottle The special “light-block” plastic bottles designed to protect the vitamins in milk from breaking down block all UV-B light and virtually all UV-A, while allowing through only 14% of PAR. Sunglasses Your own sunglasses, if bought recently, come with a rating as to how much UV-A and UV-B they block. It would be interesting to test these as a filter, both for their protective effects on the embryos as well as for the reliability of their advertised claims. Sunscreens The sunsreens you buy to shield your skin may or may not be effective in blocking UV radiation. You could test them by applying a thin film on a UV-transparent substance such as a plastic petri dish or a plastic sandwich bag.

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UV opaque plexiglass A specially formulated plexiglass can be obtained (Plexiglas ® G UF-3 acrylic) that blocks all of UV-A and UV-B but allows through PAR. Filters that do not block either UV-A or UV-B (UV transparent) These can be used as a control cover, so that all cultures are covered by a plastic sheet. Plastic petri dish A plastic petri dish allows through most UV-A (81%) and substantial amounts of UV-B (50%) radiation as well as PAR. Plastic sandwich bag The plastic bags you use for food storage allow through almost all UV-A, UV-B, and PAR. Plastic food wrap The plastic food wrap that you commonly wrap around your sandwiches allows through virtually all UV-A, UV-B, and PAR. UV transparent plexiglass A specially formulated plexiglass can be obtained (Plexiglas® G UVT acrylic) that doesn’t block PAR, UV-A, or UV-B. It does block UV-C. Cellulose acetate Cellulose acetate, like UV transparent plexiglass, doesn’t block PAR, UV-A, or UVB, but it does block UV-C. Since it is less expensive than UV transparent plexiglass, it may be a preferred material to use. Safety first Remember that any bulb emitting UV light will be damaging to you. UV light is very damaging to your eyes: it can cause cataracts by damaging your lens, and blindness by damaging your retina. You must wear protective glasses, therefore, when exposed to UV light. If possible, avoid having the UV lights on when you are present. UV light also causes photoaging of the skin (making your skin leathery and lined), as well as skin cancer. So you should also wear protective clothing whenever you risk exposure to UV light.

Monitoring Your Cultures In monitoring your cultures, you can look for a number of things: fertilization delays, cleavage delays, and developmental delays, as well as abnormalities in any of the stages. Record any differences that you see in your cultures as a percent of normal. Count 100 embryos, then record the number of abnormal/normal. Data simply on the timing of development are valuable. Realize that an alteration in the rate of development can cause embryos to emerge as juveniles when natural conditions may not be optimal for them. You can record when your cultures reach the blastula stage; the appearance of a hatched blastula, early gastrulation, and late gastrulation; the stage of spicule formation; and so on. Always compare your experimental results with a control culture. Developmental abnormalities are also very likely to occur, especially on exposure to increased levels of UV-B radiation. Look for any developmental abnormalities, such as a blastula stage that fills up with cells, exogastrulae, abnormally long cilia, and extra spicules. The abnormalities you observe may have to do with size differences. You can determine changes in size by measuring the diameters of cleavage-stage embryos and both the outer and inner diameters of blastula stages and later. This will allow you to determine if a smaller blastocoel is caused by a smaller embryo or by taller cells.

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Keeping Up with the Issues When it comes to dealing with the issue of UV radiation, it is important to avoid being part of the problem and to actively be part of the solution. Increased UV radiation levels affect all of us, and already we are experiencing the effects through increased levels of skin cancer and cataracts. One way to participate is to keep informed. Several websites are devoted to ultraviolet radiation monitoring and can keep you up to date on increased levels in your area as well as globally. For the United States, these include the National Ultraviolet Monitoring Center (http://oz.physast.uga.edu/) and the EPA’s Ultraviolet Radiation Monitoring Network (www.epa.gov/uvnet). Another EPA site deals specifically with the depletion of the ozone layer (www.epa.gov/docs/ozone). It details how you can avoid adding to the problem by explaining what substances cause ozone depletion and what substances can be used as substitutes. It also lists regulations that are in place. A site that gives information on international efforts to protect the ozone layer, run by the United Nations, can be found at www.undp.org/ montrealprotocol/ozone.htm.

Accompanying Materials See Vade Mecum: “Sea urchin–UV.” This chapter of the CD has interactive units on normal sea urchin development and on the developmental abnormalities in sea urchin embryos caused by UVB exposure. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. Contains a chapter that discusses the harmful effects of ultraviolet radiation on embryos. Fink, R. (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Sequence 1. Shows sea urchin development from fertilization through spicule formation.

Selected Bibliography Abney, J. R. and B. A. Scalettar. 1998. Saving your students’ skin. Undergraduate experiments that probe UV protection by sunscreens and sunglasses. J. Chem. Ed. 75: 757–760. This article analyzes the UV-blocking capabilities of a number of commercial sunscreens and sunglasses. Adams, N. L. 1998. Ultraviolet radiation affects development but not reproduction of green sea urchins. Amer. Zool. 38 (5): 160A. A short report examining the extent that UV radiation interferes with reproduction in the sea urchin. Adams, N. L. 1999. The green sea urchin, Strongylocentrotus droebachiensis, covers itself in response to ultraviolet radiation. Amer. Zool. 39: 113A. A beautiful study showing that sea urchins protect themselves from UV radiation in part by covering themselves with shells and seaweed, or whatever is available to them. Adams, N. L. and J. M. Shick. 1996. Mycosporine-like amino acids provide protection against ultraviolet radiation in eggs of the green sea urchin, Stongylocentrotus droebachiensis. Photochem and Photobiol., 64: 149–158. A detailed analysis of MAAs in sea urchin eggs, this study reports that MAAs do protect these eggs from UV radiation. Anderson, S., J. Hoffman, G. Wild, I. Bosch and D. Kabentz. 1993. Cytogenetic, cellular, and developmental responses in Antarctic sea urchins (Sterechinus neumayeri) following laboratory

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ultraviolet-B and ambient solar radiation exposures. Antarctic J. 28: 115–116. An analysis of the dangers that ozone holes over Antarctica pose for sea urchins. Blaustein, A. R., P. D. Hoffman, D. G. Hokit, J. M. Kiesecker, S. C. Wall, and J. B. Hays. 1994. UV repair and resistance to solar UV-B in amphibian eggs: a link to population declines? Proc. Natl. Acad. Sci, USA 91: 1791–1795. This paper discusses the harm that UV-B radiation causes in amphibian eggs and the protective effects of photolyase in these eggs. Blaustein, A. R., J. M. Kiesecker, D. P. Chivers, D. G. Hokit, A. Marco, L. K. Belden and A. Hatch. 1998. Effects of ultraviolet light on amphibians: Field experiments. Amer. Zool. 38: 799– 812. An excellent review of the effects of UV radiation on numerous amphibian species. De Gruijl, F. R. and P. D. Forbes. 1995. UV-induced skin cancer in a hairless mouse model. BioEssays 17 (7): 651–660. This is an excellent discussion of the hazards of UV radiation that humans are commonly exposed to. Downes, A. and T. P. Blunt. 1877. Researches on the effect of light upon bacteria and other organisms. Proc. Roy. Soc. Lond. 26: 488–500. A lesson in history. As early as 1877, these researchers showed that UV radiation induced mitotic delay in cells. Dunlap, W. C. and J. M. Shick. 1998. Ultraviolet radiation-absorbing mycosporine-like amino acids in coral reef organisms: a biochemical and environmental perspective. J. Phycol. 34: 418–430. A superb review of the protective value of MAAs in coral reef organisms. Epel, D., K. Hemela, J. M. Shick and C. Patton. 1999. Development in the floating world: defenses of eggs and embryos against damage from UV radiation. Amer. Zool. 39: 271–278. A well-written article that discusses many of the questions surrounding UV radiation damage to embryos while focusing of the damage and protective devices in tunicate embryos. Giese, A. C. 1946. Comparative sensitivity of sperm and eggs to ultraviolet radiations. Biol. Bull 91: 81–87. An early study that shows that both sperm and eggs are ill-affected by UV radiation. The radiation used was in the UV-C range. Gulko, D., M. P. Lesser and M. Ondrusek. 1995. Introduction to materials and methods commonly used by participants in the 1994 H.I.M.B. Summer Program on UV Radiation and Coral Reefs. Ultraviolet Radiation and Coral Reefs, 1995. (D. Gulko and P. L. Jokiel, eds.) HIMB Tech. Report #41. UNIHI-Sea Grant-CR-95-03. Pp. 19–23. This very handy paper gives the nitty-gritty details about the sources and properties of materials used in UV radiation studies. Kerr, J.B. and C. T. McElroy. 1993. Evidence for large upward trends of ultraviolet-B radiation linked to ozone depletion. Science 262: 1032–1034. An important paper verifying that depletion in the ozone of the stratosphere has caused increases in our exposure to UV-B radiation. Mead, K. S. and D. Epel. 1995. Beaker versus breakers: how fertilization in the laboratory differs from fertilization in nature. Zygote 3: 95–99. This is a concise summary of the ecology of fertilization, including a discussion of the protective devices sea urchin eggs must have against ultraviolet radiation. Milne, D. H. 1995. Marine Life and the Sea. Wadsworth Publishing Co., Belmont, CA. This textbook gives a concise description of ultraviolet radiation in our atmosphere and the destructive effects of CFCs on ozone. Peak, M. J. and J. G. Peak. 1989. Solar-ultraviolet-induced damage to DNA. Photodermatology 6: 1–15. A sophisticated review of the damage that UV radiation causes to DNA. Rustad, R. C. 1971. Radiation responses during the mitotic cycle of the sea urchin egg. In Developmental Aspects of the Cell Cycle (I. L. Cameron, G. M. Padilla and A. M. Zimmermann, eds.). Academic Press, New York, pp. 127–159. This is an excellent review of the specific damage that UV

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radiation has on eggs that results in cleavage delay. It also discusses the ill effects that UV radiation has on sperm. Shiroya, T., D. E. McElroy and B. M. Sutherland. 1984. An action spectrum of photoreactivating enzyme from sea urchin eggs. Photochem. and Photobiol. 40: 749–751. An important reference showing that the range of light that activates photolyase repair mechanisms in sea urchin eggs is 313–500 nm with a maximum effect at 365 nm.

Suppliers Cadillac Plastic 2855 Coolidge Highway, Suite 300 Troy, MI 48084 1-800-274-1000 www.cadillacplastic.com Plexiglas® G UVT Acrylic, 6 mm thick (UVT—allows through UV-B, UV-A, and PAR) Plexiglas® G UF-3 acrylic sheet, 6 mm thick (UVO— blocks all UV, allows through PAR only)

Allied Signal Plastics P. O. Box 1205 Pottsville, PA 17901 1-800-934-5679 www.honeywell.com Aclar 33c Fluoropolymer film, 127 microns thick 5 gauge (UVT—allows through UV-B, UV-A, and PAR)

DuPont Teijin Films Barley Mill Plaza, Bldg 27 Lancaster Pike and Route 141 P. O. Box 80027 Wilmington, DE 19880-0027 www.dupontteijinfilms.com

AIN Plastics 249 E. Sandford Blvd. P. O. Box 151 Mount Vernon, NY 10550 1-800-431-2451 www.ainplastics.com Mylar type D fluoropolymer film, 127 microns thick (5 mil) (Blocks UV-B—allows through UV-A and PAR)

Polycast Technology Corporation 70 Carlisle Place

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Stamford, CT 06902 1-800-243-9002 www.plasticstechnology.com

Carolina Biological Supply Company 2700 York Road Burlington, NC 27215 1-800-334-5551 or 1-910-584-0381 www.carolina.com UV 340 bulbs (mimic the UV-A and UV-B range of nature) UV-C (germicidal) lamps—not environmentally relevant

Science supply companies such as: Ward’s Natural Science Establishment, Inc. P.O. Box 92912 5100 West Henrietta Road Rochester, NY 14692-9012 1-800-962-2660 www.wardsci.com UV safety goggles

Glossary Antioxidant: A substance that neutralizes reactive oxygen species by offering their own electrons and so protect other molecules from harm. Beta-carotene, vitamins A, C and E, and the minerals selenium and zinc are antioxidants. Archenteron: Literally, ancient gut. The primitive gut formed during gastrulation. Artificial seawater: A salt solution that can be used instead of sea water. It includes the major salts found in sea water—sodium chloride, potassium chloride, magnesium chloride, magnesium sulfate, and calcium chloride. Black lights: A blacklight, common at college parties (e.g., NEC T10, 20W), provides UV-A light. Blastocoel: The cavity within a blastula. Blastula stage: The stage in embryonic development following cleavage and preceding gastrulation. The embryo may form a hollow blastula (coeloblastula) or a solid blastula (stereoblastula). The cavity formed in a coeloblastula is called the blastocoel. CFCs: Chlorofluorocarbons. Chlorofluorocarbons (CFCs): Compounds that can carry chlorine into the upper atmosphere where the chlorine destroys ozone. CFCs include freon, chloroform, carbon tetrachloride, various cleaning agents of computer circuit boards, and propellants for aerosol cans.

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Dry sperm: Sperm that have been shed and are kept in this concentrated form, undiluted. In experimental procedures, it is often useful to store sperm as dry sperm, since they survive for longer periods in this form. Echinopluteus (pluteus) larva: The type of larva typical of echinoids (sea urchins, etc). It consists of a delicate body with long arm-like extensions supported by spicules. Exogastrula: A deformity in developing embryos in which the archenteron, instead of invaginating, evaginates, forming an external archenteron. Fertilization: Fusion of egg and sperm. Gastrulation: The stage in embryonic development that follows cleavage. During gastrulation, cells rearrange themselves, the endoderm and mesoderm cells moving inward and the ectoderm cells spreading around the outside, with some ectoderm invaginating to form such structures as the stomodeum. Germicidal lamp: A UV-bulb that emits light enhanced in the UV-C range. It is used to sterilize an area. It is dangerous to the eyes and skin, and should be turned off before a person enters the work area. Grow lights: A light commonly used to enhance growth of house-grown plants. It provides light enhanced in the UV-A range without emitting light in the UV-B range. Hatched blastula: A term used to describe the echinoid (sea urchin, etc.) embryo when it has hatched out of its fertilization envelope. This occurs early in development, just after primary mesenchyme cells have started to ingress. Hydrogen peroxide: H2O2 . A highly reactive compound that can damage biological systems by oxidizing components such as proteins, lipids, and DNA. Invagination: The folding inward of a sheet of cells. This is a common type of movement during gastrulation. In the sea urchin, for example, the archenteron first forms by an invagination of the vegetal plate. MAAs: Mycosporine-like amino acids. Micromeres: Cells in a blastula that are smaller than the others. In the sea urchin embryo, micromeres form at the vegetal pole at the 16-cell stage. These will form the primary mesenchyme. Mycosporine-like amino acids (MAAs): Amino acids that absorb UV-B radiation and therefore can protect organisms against the harmful effects of UV-B radiation. Found in photosynthetic algae, fungi, and bacteria, and those organisms that accumulate MAAs through their diet. Considered a natural sunscreen. Ozone: O3. An allotropic (alternative) form of oxygen. In our upper atmosphere, it protects the Earth’s surface from UV-B radiation. In the lower atmosphere, it is a pollutant that is harmful to the health of organisms. Ozone holes: Holes in the protective ozone shield in the Earth’s stratosphere that are allowing greater amounts of the exceedingly harmful UV-B radiation to penetrate to the Earth’s surface. As much as 5% of the this ozone shield has disappeared in less than a 10-year period. PAR: Stands for photosynthetically active radiation. This is the range of wavelengths (400-700 nm) that can activate photosynthetically active pigments. It is not considered harmful. Parthenogenesis: The development of an egg without fertilization. Photoaging: Aging of tissues that is induced by UV-radiation. This can be attributed in large part to the cross-linking of collagen. Highly cross-linked collagen in the dermis causes wrinkles. Photoaging of the skin results in leathery-looking skin that is deeply lined. Photolyase: An enzyme that is activated by light and that repairs UV-induced DNA dimers. Photorepair mechanisms: Cellular repair mechanisms that are activated by light.

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Pluteus larva: The type of larva typical of echinoderms. It consists of a delicate body with long armlike extensions supported by spicules. Primary mesenchyme: Mesoderm cells in echinoid embryos (such as those of the sea urchin) derived from the micromeres. The primary mesenchyme are the first cells to enter the blastocoel during gastrulation. They form the spicules of the larva. Prism larval stage: In an echinoid (sea urchin, etc.) larva, the stage at which the larva is shaped like a prism. It is an early stage in larval development. Pyrimidine dimers: Dimers of any of the pyrimidine bases: cytosine, thymine, uracil. Reactive oxygen species: Highly reactive oxygen containing molecules that cause damage to biological systems by oxidizing components such as proteins, lipids, and DNA. Examples are -

superoxide anions (O2 ) and hydrogen peroxide (H2O2). Skin cancer: Cancer that starts in the skin. Types include melanoma, a deadly cancer if not caught early, squamous cell carcinoma, and basal cell carcinoma. UV-radiation is thought to be responsible for most incidences of basal cell and squamous cell carcinomas. A reasonable correlation exists between UV-radiation exposure and melanoma, but the relationship is not as clear as with the other two types of cancer. Spawning: The release of gametes (eggs or sperm). Spicules: In the echinoid (sea urchin, etc.) larva, these are the calcium carbonate structures that serve as the skeletal supports for the larva. The are laid down by the primary mesenchyme cells. Standard sperm suspension: The diluted form of sperm that is used for fertilization studies. In the sea urchin, for example, this is 1 drop of dry sperm in 10 ml of seawater. Stratosphere: Earth’s upper atmosphere. It contains a protective layer of ozone which prevents most of the highly harmful UV-B radiation from penetrating to the Earth’s surface. Sunlamps: A lamp that provides light enhanced in the UV-B range. Sunlamps from 20 years ago emit high levels of both UV-A and UV-B radiation. Ultraviolet radiation: Radiation of wavelengths that fall between 200 and 400 nanometers (nm). UV 340 lamps: A light bulb that supplies light in both the UV-A and UV-B ranges and mimics the light found in nature. UV-A: The longest wavelengths (320–400 nm) within the ultraviolet range. This range is harmful to biological organisms, but is less harmful than UV-B or UV-C. Most of the UV radiation reaching the Earth’s surface is in the UV-A range. UV-B: The range of wavelengths within the ultraviolet range that fall between 280–320 nm. (Some sources use 290–320 nm.) UV-B radiation is significantly more harmful to biological organisms than UV-A. Most of the UV-B radiation coming from the sun is blocked by the ozone in the Earth’s upper atmosphere. Holes developing in this ozone layer are allowing greater amounts of UV-B radiation to penetrate to the Earth’s surface. UV-B filters: Filters that block UV-B radiation, but allow UV-A radiation and PAR to pass through. Examples include glass, Xerox transparencies, plastic milk bottles, and Mylar film. UV-C: Ultraviolet radiation in the range of 200–280 nm. This is the most highly dangerous range of UV radiation for biological organisms. It is absorbed by oxygen in the atmosphere, such that it never reaches the Earth’s surface. UV-opaque filters: Filters that block UV-A and UV-B radiation, but allow PAR to pass through. Examples include standard plexiglass, sunglasses rated as 100% UV-blocking, light-block plastic milk bottles, and many cosmetic sunscreens.

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UV-transparent filters: Filters that allow UV-A, UV-B radiation and PAR to pass through. Examples include plastic petri dishes, plastic sandwich bags, plastic food wrap, UV-transparent plexiglass, and cellulose acetate.

Sea Urchin – UV Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. What is the most harmful range of UV radiation that reaches the Earth's surface?

2. Why is the hole in the ozone layer a danger to us and other organisms?

3. What damage can UV radiation cause in organisms?

4. What filters can be used to block UV-B radiation?

5. What filters can be used to block both UV-A and UV-B radiation?

Name: _______________________

Sea Urchin UV - 2

6. How might you set up an experiment to distinguish between the effects of UV-A and UV-B radiation on the development of sea urchin embryos?

7. What effect does UV radiation have on cleavage in the sea urchin embryo?

8. What abnormalities can be seen in the blastula and gastrula stages of UV-irradiated sea urchin embryos?

9. What abnormalities can be seen in the larval stages of UV-irradiated sea urchin embryos?

10. Do you think that the enlargement of the hole in the ozone layer in our stratosphere is a threat to sea urchins? To other organisms with aquatic embryonic stages?

chapter

8

Development of the Fruit Fly

Drosophila melanogaster

Life Cycle The fruit fly Drosophila melanogaster is the familiar visitor on your overripe bananas and an organism of choice in genetics laboratories. As bridges between genetics and developmental biology are both built and traveled upon, it becomes imperative that developmental biologists study Drosophila to aid in the union of these two disciplines. Some of the major questions in developmental biology can only be answered with genetics. So we must learn about the geneticists’ organisms and make them ours as well. An embryologist well-schooled in Drosophila development is both rare and valuable. Study this laboratory exercise well and it could make you some bucks in the future. Drosophila is a holometabolous insect—that is, an insect that has a larval and a pupal stage prior to the adult stage. (Hemimetabolous insects, on the other hand, have nymph stages preceding the adult.) The adult Drosophila may live for more than 10 weeks. During this time, mating takes place. Fertilization is internal, and sperm are stored within the female’s body in a seminal receptacle and the paired spermathecae. Females reach the peak of their egg production between the fourth and seventh day after their emergence. During this time, they lay eggs almost continuously at a rate of 50–70 eggs per day. The eggs are approximately one-half mm in length, white, oval, and slightly flattened in lateral view (they look much like a kernel of rice). The ovum is surrounded by an inner, very thin vitelline envelope and an outer, tough extracellular coat called a chorion. At its anterior end two small filaments, extensions of the chorion, extend from the dorsal surface. These are respiratory filaments and serve for gas exchange, as their name implies. Eggs are laid half-buried in rotten fruit or the medium in your culture jars, and the filaments protrude into the air. Eggs hatch in 22–24 hours at 25°C. The larva that emerges looks like a tiny worm and is called the first instar larva. It feeds on the substrate that the eggs were laid in and, after another 25 hours, molts into a larger wormlike form, the second instar larva. This feeds as well and, after about 24 hours, molts into the third instar larva. This is the largest of the larval forms. It feeds, but it also starts to climb upward out of its food, so that it will be in a relatively clean and dry area to undergo pupation. The third instar molts into a pupa after 30 hours. The pupa is stationary, and in its early stages is yellowish-white. As it develops, the pupa becomes progressively darker. During the pupal stage, the larva is metamorphosing into the adult fly, also called the imago. In doing so, it lyses most of the larval structures, although some larval organs are preserved. The larval nervous system, for example, is not lysed, but even it undergoes major restructuring; the Malpighian tubules (excretory structures), fat bodies, and gonads are kept as well. Most of the adult structures, however, form anew from two sets of cells that have been carried as undifferentiated, mitotic cells within the larva throughout its instar stages: these are the imaginal discs (imaginal since they are for the imago) and the histoblasts.

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Imaginal discs These are small, almost teardrop-shaped packets of epithelial cells that will form the epidermal structures of the adult, such as the wings, legs, eyes, mouthparts, and genital ducts. Imaginal discs are carried around within the larva, growing in size but not differentiating. During the pupal stage, they evert and differentiate into their adult structures. Histoblasts These cells are found in small groups (nests) within the larva. They form the abdominal epidermis and the internal organs of the adult. They, too, grow by mitosis during the larval stages and then differentiate during the pupal stage. They are recognizable within the larva as clumps of small cells nestled among the huge differentiated polytene larval cells. The pupal stage lasts for 3–4 days, after which the adult fly, or imago, emerges from the pupal case (eclosion). Adult male flies are sexually active within hours of emerging, females don’t have ripe eggs until two days after eclosion, and the cycle begins again. Pause for a minute. Think about what you’ve just read. It’s weird! What does the Drosophila do during its life cycle? It has a larval form: a fully differentiated, feeding organism, that carries around, as extra baggage, cells that will replace it—cells that will become another fully differentiated, feeding organism. The larva is only a vessel, a nurturing culture environment for these cells that become the adult. At the appointed time, the larva self-destructs as these “passenger cells” differentiate. It is an astonishing way to make an adult. Culturing Drosophila melanogaster As you have undoubtedly noticed from the fruit basket that sat too long, Drosophila thrive on fermenting soft fruits. A very suitable culture medium, therefore, is crushed banana. It provides all the necessary nutrients for both the larval and adult stages. The banana can be kept along with the flies in sterile pint jars with cotton or foam rubber plugs. Another standard medium, commonly used by laboratories that raise Drosophila, is a cornmealmolasses-agar mixture. While the batch brews, it fills the scientific hallways with the smells of Grandpa’s favorite cookies. Cornmeal-molasses-agar culture for Drosophila water 420 ml agar 4.5 gm unsulfured molasses 60 ml cornmeal brewer’s yeast cold water

49 gm 6.5 gm 145 ml

propionic acid

3.4 ml

. agar 3–5 minutes. Mix and boil water and Add unsulfured molasses and heat to boiling again Mix together cornmeal, brewer’s yeast, and cold water in a separate container until all lumps are removed. Add cornmeal-yeast mixture to molasses-agar mixture. Boil 5 minutes, stirring constantly. Cool mixture to 60°C. Add propionic acid (as mold inhibitor). Pour culture medium 1-inch deep into sterile culture jars with sterile plugs. Pint milk bottles work well, but any widemouthed jar fitted with a plug made of cotton covered with cheesecloth or foam rubber should work well. Add a sprinkle of active baker’s yeast (from a salt shaker) to each jar before adding flies.

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It is important when maintaining cultures not to overcrowd (about 100 flies per pint culture jar) and to subculture approximately every other day. This keeps the flies healthy, large, and mold-free.

Collecting Eggs Collecting fertilized eggs is easy, but it is not easy to catch the very early stages of development, since eggs can be held within the female’s uterus after fertilization for a period of time, even as late as just prior to larval hatching. When a female is laying rapidly, however, the uterus is being cleared fast, and eggs in their early stages of development can be obtained. To achieve this, it is best to use cultures of flies that are 5 days old. A female is within her peak laying period at this time and is laying eggs as quickly as one every 3 minutes. Collecting chamber A simple collecting chamber consists of an empty culture bottle (any widemouthed bottle will do) with wet toweling stuffed in the bottom (for humidity) and a cotton or foam rubber plug. Place approximately 40 pairs of flies in the chamber by inverting a culture bottle containing flies over the mouth of the empty bottle. Holding the bottles together, bang the empty bottom bottle against padding on a tabletop to cause the flies from the upper culture bottle to drop into it. Quickly replace the plugs of both bottles. Use plastic spoons whose handles have been cut so they fit in the culture chamber without touching the plug. Put culture medium on the spoon (the same culture medium that the flies have been grown in), score it to make grooves (female flies like to lay their eggs in moist grooves), paint a light coating of baker’s yeast suspension (a slurry of bread-making yeast in water) on the surface of the scored medium, and place one or two of these spoons in the collecting chamber with the flies. Since flies will outfly your quick fingers and escape from the bottle during this process, it helps to face the bottom end of the bottle toward a bright light. The flies will be attracted away from the mouth of the bottle. Place the bottle on its side so the medium won’t slip off the spoons. After about an hour, a suitable number of eggs will have been laid on the spoons, and they can be removed and replaced with fresh spoons containing medium. Again, attract the flies away from the bottle mouth with a bright light. If you prefer to have a number of different stages, including advanced stages, on a single spoon, leave the spoon in the collecting chamber for an extended period (up to 24 hours). Females produce the greatest number of eggs in the late afternoon and evening. Mating behavior of adult flies Before removing the spoons from the bottle, observe the adult flies through the glass, or place males and females in a small covered petri dish and observe them under the dissecting scope. To remove flies easily, first place the bottle in the refrigerator or keep it on ice for at least 20 minutes. This numbs the flies, and you can remove them without their escaping into the room. (Numbing with cold is an easier and safer way than anesthetizing with ether or CO2.)You can distinguish males from females by looking for the black pigmentation on the posterior abdominal segment of the males; it is absent from females. Also, males have a shorter abdomen with six segments rather than eight, and they have a sex comb (a fringe of ten or so black, stout bristles) on the end of the first segment of the front legs. Watch for courtship behavior. A female is very much in control of whether she is inseminated, being larger and stronger than a male. She must give an acceptance signal by slowing down, extruding her ovipositor, and spreading her wings, in order for mating to occur. There is no known incidence of

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rape among these organisms. A female rejects a male by kicking with her hind legs, fending with her middle legs, flicking her wings, producing a rejection buzzing sound by fluttering her wings, or moving away rapidly. If she has already mated, she also will extrude her genitalia to reject the male. A male courts anything that produces the right “taste” or “smell” (even other males if they are immature). He orients himself toward the female’s head, taps her with his forelegs, “tasting” her to make sure she is the right species, and then pursues her when she moves, extending and vibrating one wing producing a courtship song. Though the “love song” of the Drosophila is species-specific, females do respond to the songs of other species as well. Females “hear” the song through their antennae; the aristae (feathery extensions of the antennae) augment the vibrations, and they are sensed by Johnston’s organ in the second segment of the antenna. Later in the courtship, the male extends his proboscis to touch the female’s genitalia. If all the active courtship of the male has stimulated the female enough to accept the male, the two mate with the male positioned on top of the female. Sperm travel through the male penis into the female uterus and then swim into the female seminal receptacle and spermathecae, where they are stored for fertilization. During oviposition, the eggs emerge from the female’s ovipositor posterior end first. The female oviposits preferentially on a moist food surface in a humid atmosphere. If the air is too dry, the female may feed, but she won’t oviposit.

Observations of the Egg (Use Sterile Technique) After a suitable waiting period, remove a spoon from the collecting chamber, and look at the surface of its medium under a dissecting scope. You should see a number of white eggs, often with just their respiratory filaments sticking out of the culture medium. You may find the eggs clustered in patches, laid preferentially on the moister regions of the medium. Scan the entire surface of the medium. Use a microknife or fine forceps to remove the eggs to a small petri dish containing insect Ringer’s solution. (This is a balanced salt solution that will allow continued development of the egg until hatching.) Insect Ringer’s solution NaCl 7.5 gm KCl 0.35 gm CaCl2 0.21 gm

Make up to 1 liter with distilled water.

The chorion Observe the eggs under a dissecting microscope and record your observations with diagrams. Your initial observations will have to be of superficial structures until you remove the chorion from the egg. Notice that the chorion is thick and tough, and that the two respiratory filaments and a micropyle are extensions of this chorion (Figure 8.1). The micropyle is a tiny channel that leads to the ovum, and it is through this channel that a sperm must swim in order to fertilize the egg. Though there have been reports of polyspermy in Drosophila, normally only one sperm is successful in making the trip. What do you think the purpose of the micropyle is? Record your answer in your notebook. Now place several of the eggs in a drop of Ringer’s solution on a slide and observe these under a compound microscope. Focus at the surface of the chorion as well as deeper. Place a small drop of toluidine blue over the eggs. This dye will help to show the ornamental markings of the chorion. These markings are beautiful. They are the impressions (“footprints”) left by the ovarian follicle cells that deposited the chorion prior to ovulation.

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The chorion is a complex structure, consisting of inner and outer laminae bounding a layer of tiny air pockets that connect anteriorly with an extensive meshwork of airspaces in the respiratory filaments. This design maximizes gas exchange and minimizes water loss. The respiratory filaments are the major region of gas exchange. By restricting gas exchange to such a small surface area, the egg is able to minimize water loss when it finds itself stuck in the drying winds of a dry summer. In addition to a meshwork of air pockets, the respiratory filaments have a water-repellent surface network that maintains a film of gas (a plastron) around them when submerged. The plastron allows the respiratory filaments to function as a physical gill if the egg gets trapped in a rain puddle. The egg’s chorion, therefore, allows survival of the egg through the rainy season and the dry. To see some of the complexities of the chorion and respiratory filaments, put a footed coverslip over the egg and adjust your microscope to increase contrast. (A footed coverslip is made by nicking the corners of the coverslip against some hard paraffin so that crumbs of paraffin remain attached at each corner. This will give just enough spacer between the slide and the coverslip to avoid crushing the egg. Adjust the size of the crumbs to the size of the eggs. Add more Ringer’s solution underneath the coverslip as needed.) Examine the respiratory filaments under higher power. What differences do you see between them and the rest of the chorion? Include your answers in your notebook with diagrams of what you see. Focus deeper to the level of the embryo within the chorion, and try to stage your embryo using the staging series provided. You will be correcting your stagings after you remove the chorion.

Respiratory filament Micropyle Chorion

Ovum

Dorsal view

Lateral view

Figure 8.1 Diagrams of the Drosophila egg.

Dechorionating an egg Soak an egg in undiluted bleach (such as Clorox) for 5 minutes. Agitate the solution slightly with a pipette. You should see remnants of chorion as it separates from the egg. Rinse the egg in Ringer’s solution. If the chorion is not gone, repeat the operation. Note: In moving eggs from one solution to another, a micropipette can be used, but the egg often sticks to the glass and remains within the pipette. Better ways are: to transfer the egg using a hairloop; to catch the egg in the meniscus between half-closed tongs of fine forceps; or to let the egg stick to the side of a needle. Place the dechorionated egg in a drop of Ringer’s solution on a slide, and observe it under a compound microscope. Determine its stage of embryogenesis using the descriptions and staging series below.

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Embryogenesis Study the written description of embryogenesis below, and then proceed with your observations. Look at several embryos to see as many stages of embryogenesis as possible. Cleavage The eggs you collect will be in various stages of development. It is difficult to see what is going on inside the egg both because of the chorion (if you haven’t removed it) and because the egg is very yolky. It is a centrolecithal egg, meaning that the yolk is concentrated centrally and the cytoplasm is pushed to the periphery. Cleavage is unusual in that nuclear division (karyokinesis) occurs many times before the cytoplasm cleaves (cytokinesis). The nuclei of early cleavage are centrally located through the first seven divisions (Figure 8.2A), after which they start migrating to the periphery (Figure 8.2B). By the time there are about 5000 nuclei all lined up in the peripheral cytoplasm, cell membranes are laid down between them, making a peripheral layer of separate cells, and the embryo goes from being a syncytial blastoderm (Figure 8.2C) to a cellular blastoderm (Figure 8.2D). This pattern of cleavage is called superficial or peripheral cleavage. Cellularization of the blastoderm does not occur simultaneously around the egg. The cells that form first are at the posterior end. They are relatively large and are called the pole cells (Figures 8.2C, D). The pole cells form the primordial germ cells, which give rise to the gametes. It is interesting that they should be set aside so early in development. Do you have any suggestions as to why? Gastrulation Following cleavage, gastrulation proceeds primarily by the infolding of a midventral band of cells. First a ventral furrow appears as the mesoderm folds inward (Figure 8.2E). At the anterior and posterior ends of this furrow, the endoderm invaginates forming the anterior and posterior midgut (Figure 8.2F). Later the ectoderm will also invaginate to form the anterior stomodeum (foregut) and posterior proctodeum (hindgut) (Figures 8.2G, H). In addition to these ventral infoldings, there is also a lateral infolding toward the anterior end which extends around the circumference of the embryo. This is the cephalic furrow (Figure 8.2F) and roughly delineates the boundary of the future head. Gastrulation creates a multilayered band of germ layers on the ventral side of the egg that curves around the egg’s posterior tip. This band is called the germ band. It elongates along the dorsal side of the egg so that eventually, like an acrobat with her back arched and her legs gracefully curved back to touch her head, the embryo’s posterior end meets its head end (Figures 8.2G, H). The germ band then shortens and thickens, bringing the posterior end of the embryo back toward the posterior pole of the egg (Figures 8.2I, J). As the germ band shortens, definitive segmental boundaries appear, marking off head regions (mandibular, maxillary, labial), thoracic segments (t1–t3), and abdominal segments (a1–a10). (In the larva you will find only eight abdominal segments. Abdominal segments 9 and 10 have formed the telson of the larva, a tail-like structure.). During gastrulation, a peculiar thing happens. The developing head disappears from view—it turns inward, or involutes (Figure 8.2K). Meanwhile the thoracic segments expand forward, overgrowing the region that used to be “head.” Only a tiny external head will remain. So when you finally look at your Drosophila larvae, don’t be too surprised when they appear to be headless.

Development of the Fruit Fly

(G) (A) Dorsal Anterior

Cleavage nuclei

(B)

Hindgut

(H)

Stomodeum (I)

(C)

Pole cells

(D)

Foregut

(J)

Hindgut

(K)

(E)

Posterior plate Pole cells

(L)

Midgut

(F)

Salivary gland

Midgut

Ventral nervous system

Cephalic furrow

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Development of the Fruit Fly

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Figure 8.2 L Stages in embryonic development of Drosophila. (A) Early cleavage (15 minutes–1.5 hours). Nuclei cleave in the central region. (B) Migration of cleavage nuclei (1.5 hours). Nuclei migrate to the periphery. (C) Formation of syncytial blastoderm (2 hours). Pole cells form posteriorly. (D) Cellular blastoderm (2.5 hours). Cell membranes form between the nuclei. (E) Early gastrulation (3.5 hours). The ventral furrow forms. Thickening of the posterior plate below the pole cells. (F) Midgut invagination (3.5–4 hours). The midgut invagination can be seen ventrally, as can the cephalic furrow. (G) Germ band extension (4–5 hours). Invagination of the hindgut can be seen dorsally. (H) Stomodeal invagination (5–7 hours). Invagination of the stomodeum can be seen ventrally. (I) Shortening of the germ band (9–10 hours). Foregut and hindgut invaginations are deep. (J) Shortened embryo (10–11 hours). Hindgut is now fully posterior. (K) Dorsal closure (13–15 hours). The ectoderm closes dorsally. The midgut broadens. The head involutes. (L) Condensation of ventral nervous system (15 hours–hatching). The gut regions are joined. The nervous system forms ventrally.

Later development By 16 hours of development, muscular movement will be apparent (Figure 8.2L). Just before the embryo hatches as the first instar larva at 22–24 hours, you will be able to see air-filled tracheae and other internal organs. Embryonic staging series You can use the staging series shown in Table 8.1 to stage your embryos; it is one of the more widely used series for Drosophila embryonic development. (Whenever you refer to a particular stage, you must cite the source of the staging series you’ve used, since a number of different ones exist.) Table 8.1 Embryonic stages of Drosophila Stage

Time (hours)

Developmental event (at 25ºC)

1 2

0–0:25 0:25–1:05

3

1:05–1:20

4

1:20–2:10

5

2:10–2:50

6

2:50–3:00

7

3:00–3:10

First two nuclear divisions. Egg uniformly dark in center and light at periphery. Nuclear divisions 3–8. Egg cytoplasm retracts considerably from vitelline envelope, leaving empty space at anterior and posterior poles. At posterior end, three polar buds form (later to pinch off and become pole cells), and divide once. Nuclear division 9. Dividing blastoderm nuclei cause granulated appearance in wide zone in periphery. Empty space at anterior pole disappears. Blastoderm nuclei in periphery making a bright peripheral rim. Nuclear divisions 10–13, just prior to cellularization. Two more divisions in polar buds. Cellularization of the blastoderm occurs, and nuclei elongate considerably. Pole cells begin to shift dorsally. Midventral blastoderm cells look irregular and wavy, preceding their invagination. Early gastrulation: ventral furrow forms, from which mesoderm and endoderm originate; at posterior pole, cells shift dorsally to form a dorsal plate to which pole cells adhere; cephalic furrow becomes visible as a lateroventral slit. Endoderm of anterior and posterior midgut and ectoderm of hindgut invaginate; dorsal folds appear.

(continued)

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Table 8.1 (continued) Stage

Time (hours)

Developmental event (at 25ºC)

8 9

3:10–3:40 3:40–4:20

10

4:20–5:20

Amnioproctodeal invagination, rapid phase of germ band elongation. Transient segmentation of the mesodermal layer, visible in region of germ band as prominent bulges protruding into yolk sac. Stomodeum invaginates ventrally at anterior pole. Germ band continues to expand. Interior of egg occupied by yolk sac which is a dark, uniform mass. Periodic furrows in epidermis appear. Pole cells leave cavity of posterior midgut and locate themselves dorsally outside yold sac. Primordia of Malpighian tubules form. Neuroblasts divide.

11

5:20–7:20

12

7:20–9:20

13

9:20–10:20

14

10:20–11:20

15

11:20–13:00

16

13:00–16:00

17

16:00–24:00

Growth with no major morphogenetic changes. Intersegmental furrows form in epidermis; mandible, maxilla, and labium visible as protuberances. Germ band extension reaches its maximum extent. Posterior pole becomes withdrawn from vitelline envelope. Shortening of the germ band so that opening of hindgut becomes located at dorsal side of posterior pole. Width of germ band increases. Anterior and posterior midgut clearly visible and fuse. Germ band segmentation very prominent. Germ band shortening completed. Conspicuous triangular gap ventrally due to retraction of clypeolabrum. Labium moves to ventral midline, displacing opening of salivary gland and duct. Yolk sac protrudes dorsally, has characteristic convex shape. Dorsal fold (ridge) appears. Head involution begins. Head involution continues. Dorsal closure and closure of midgut. Anal plate ventrally displaced from posterior tip. Dorsal spiracles evident. Dorsal closure and dorsal epidermal segmentation. Gut forms closed tube containing yolk sac. Supraoesophageal ganglia and pharynx evident. Intersegmental grooves distinguishable mid-dorsally. Dorsal ridge overgrows tip of clypeolabrum. Constrictions appear in midgut. Shortening of ventral cord. Tracheal tree contains air. Retraction of ventral cord continues. Embryo hatches as first instar larva.

Source: After Campos-Ortega and Hartenstein, 1985.

Larval Development You will be maintaining cultures over the week. Place some of your eggs in a small petri dish containing culture medium. Place this dish in a larger petri dish containing sterile water; this will humidify the culture through the week. During the week, keep a record of the developmental stages you see. Record, for example, when you see first instar larvae, second instar, and so on. If one is available, keep a recording thermometer in the room and make a record of the daily temperatures. You may also keep embryos at other temperatures by using the refrigerator (4°C) or an incubator with variable heat settings. (At 22–25°C, development to an adult takes about 9 days. At 10°C, it is slowed down to 57 days. At 29°C, it is speeded up to 8 days. Temperatures continuously above 29°C can be lethal.) By next week, you will have a developmental timetable that you can compare with a standard timetable (Table 8.2). Make comparisons and suggest reasons for any differences you note. Why is development slowed at lower temperatures? Why are high temperatures lethal? Record your answers in your laboratory notebook. The sections that follow may be done next week.

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Table 8.2 Larval stages of Drosophila Time after fertilization _____________________

Hours 24 49 72 120 122 124 132 169 189 216

Days 1 2 3 5 5.1 5.2 5.5 7 7.9 9

Developmental event (at 25ºC) Hatching from egg; first larval instar begins First molt; second instar begins Second molt; third instar begins Puparium formation; puparium white Puparium fully colored “Prepupal” molt Pupation; cephalic complex, wings, legs everted Eye pigmentation begins Bristle pigmentation begins Adult ready to emerge from pupa case

Source: After Doane, 1967.

Anatomy of the Larva (Nonsterile Technique Can Be Used) Put your culture dish or a scoop of medium from the laboratory stocks containing larvae under a dissecting scope. You should see three different sizes of larvae, representing the three larval instars. The first two instars should be found burrowing through the medium. A late third instar will be climbing up, away from the food, getting ready to pupate. Watch the behavior of the larvae as they burrow and eat their way through their food. Make notes in your laboratory notebook. Notice that as the larva feeds, it extends a pair of mouth hooks that bring food to the mouth. Look at the head end again. What feature is particularly conspicuous by its absence? Right—the larva has no eyes. Does this mean that the larva is blind? Test its sensitivity to light in various ways, and record your results in your laboratory notebook. You can use a box with a hole at one end to test migration toward or away from light. Don’t “toast” your larva—that is, don’t place the lamp so close to the larva that you are testing a response to heat rather than a response to light. What other behavioral responses do you note? How does the larva respond when poked, for example? Record all your observations in your laboratory notebook. Note any differences in behavior among the three larval instars. A few minutes before each larval molt, the larva will stop feeding and lie motionless. The mouth hooks then start biting through the old cuticle. With active muscle contractions, the larva ruptures and leaves its old cuticle, discarding mouthparts and spiracles as well, which are replaced by new structures of the next larval instar. If you are very lucky, or very patient, you may see a larval molt. If you do, time the molt and make whatever observations you can, recording them in your laboratory notebook. (Hint: double mouth hooks and double spiracles [see definition of spiracle below] are evidence of an approaching molt.) Use a moist paintbrush or partially closed forceps to move larvae from the culture medium to a drop of Ringer’s solution on a slide. Try to find all three instar stages. Put the slide over ice in a petri dish to anesthetize the larvae, then observe them again under the dissecting scope.

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External anatomy Look for the tiny external head, three thoracic segments, eight abdominal segments, and a telson extending beyond the anus. These segments are delineated by ventral rows of tiny hooks, denticle belts, which prevent the larva from slipping backward as it moves forward with waves of extension and contraction. You should also see a number of sensory bristles in the first instar larva distributed over the cuticle. Internal anatomy The larvae are transparent, and, with the proper lighting, you should be able to distinguish a number of internal structures (Figure 8.3). Use both your dissecting scope and compound microscope, and play with the lighting until you are satisfied. Transmitted light, light coming from below, shining up through the larva, should be best. The two fat bodies—long, whitish sheets running the length of the body—are the most obvious structures. Embedded in the fat body in the fifth abdominal segment are the transparent, vesicular gonads. Since the testis is several times larger than the ovary, you can sex the larvae simply on the basis of gonad size. A tree of beautifully pearl-white tracheae (hollow tubes) start anteriorly at the two anterior spiracles (tufted openings to the outside) and end in the telson at the two posterior spiracles. This is the respiratory system for the larva. The gut starts anteriorly as a muscular pharynx and continues as a narrowed esophagus, which runs smack through the middle of the brain in the thoracic region. (There must be a joke or two you can come up with about animals with guts through their brains.) In the region of the esophagus, you should see two lateral transparent salivary glands, which you will be dissecting later. The esophagus empties into the heavy-walled, bulblike proventriculus, which in turn empties into the gastric area that has fingerlike, blind-ending pouches, the gastric caecae. The gastric region continues into the long coiled midgut or midintestine, which doubles back on itself and empties into the straighter hindgut or hind intestine. You should also be able to distinguish the two yellowish Malpighian tubules, the excretory organs that carry urinary waste from the body, emptying it into the posterior midgut. Focus dorsally and try to see a pulsating blood vessel. This is the heart (dorsal vessel), which extends anteriorly as an aorta. The circulatory system is open, and the hemolymph bathes the internal organs. Using a clock with a second hand, time the beats of the heart. How does it compare to your own? Does it speed up when the larva is warm? Is there a difference among the three instar larvae? Are there other differences that you note that distinguish the three larval stages from one another? Note these in your laboratory notebook. Focus again on the region near the brain. Concentrated in this region are the imaginal discs. They should appear as tiny teardrop-shaped packets, each one with a connection to the tracheal system. It is these that you will be dissecting. Check Figure 8.3 carefully to determine the position of each type of imaginal disc.

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Mouth hooks Pharynx Anterior spiracle

Labial disc Clypeolabrum disc

Humerus disc Esophagus Wing disc

Antennaeye disc

Leg disc Haltere disc

Brain Ventral ganglion

Proventriculus

Leg discs

Salivary gland

Gastric caeca

Trachea

Anterior midgut Malpighian tubules

Hindgut

Gonad

Posterior midgut

Genital disc Telson

Posterior spiracle

Figure 8.3 Schematic diagram of a third instar larva. The fat bodies and circulatory system are not shown for the sake of clarity.

Development of imaginal discs It is important to understand the development of imaginal discs before dissecting them, so you can appreciate more fully what you will be seeing. The imaginal discs have their origins in the embryo where they start out as epidermal thickenings that then invaginate to become vesicles. They never separate

Development of the Fruit Fly

Figure 8.4 Schematic diagram of eversion in a leg imaginal disc. (A) During development of the disc, an epithelial tube forms that becomes folded back upon itself as it is forced into the confines of the tiny disc, bounded by a basement membrane. The epithelium on the far side of the disc does not fold and forms the peripodial membrane. (B) During pupation, the disc everts. The folded epithelium pushes outward against the peripodial membrane, extending into an elongate tube. The distal tip of the tube is shown by five cross lines and the proximal end by small dots in the diagrams of the uneverted and everted discs. (After Poodry and Schneiderman, 1970, and Condic et al., 1991.)

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(A) Peripodial membrane

Proximal end

Distal tip Basement membrane

(B)

entirely from the epidermis, but maintain a narrow connection to it. Though there is variation among the types of discs, the general scheme of development is that once the vesicles form, one side of the vesicle starts growing considerably, bending inward as it grows, forming a short, wide tube. With nowhere to expand, squashed into the confines of its little vesicular package, the tube must fold back upon itself as it grows. The opposite side of the vesicle remains smooth and thin as the peripodial membrane (Figure 8.4A). The original basement membrane, on the outside of the vesicle, does not bend inward with the growing side and remains as an envelope around the imaginal disc. The final effect is a small, transparent package with what looks like a coiled structure (the tube) within. Though not actually coiled, but rather folded like a telescope, this tube will be what forms the final adult structure for which the imaginal disc is determined. There are nine pairs of discs: labial, clypeolabrum, humerus, antenna that is attached to the eye disc, wing, haltere, and three pairs of leg discs. There is also a single genital disc that will form the genital ducts, accessory glands, and external genitalia, but not the gonads (Figures 8.3 and 8.5). The imaginal discs are suspended within the larval body, looking casually placed, like Christmas ornaments dangling from the tracheal tree. Here they undergo growth but no differentiation. It is not until the pupal stage, the stage of metamorphosis, that the imaginal discs will take on their adult form. During pupation, each disc everts and its cells elongate. The result is that the folded tube within the disc pushes outward against the peripodial membrane, becoming long and extended (Figure 8.4B). (Imagine an old-fashioned top hat with its crown pushed down being snapped to its full height, and you’ll have an approximate image of evagination of an imaginal disc.) Differentiation of the discs then begins.

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Dissection of Imaginal Discs (Nonsterile Technique Can Be Used) The removal of imaginal discs from the larva requires skill and patience, and it is truly in the realm of microdissection. The imaginal discs you will be dissecting are so small that students before you have dubbed them the “imaginary” discs. Do not despair. Just take a deep breath, don’t drink a lot of caffeine before beginning, and be confident in your skills.

Antennal region Eye region

Labial disc

Wing disc

Antenna-eye disc

Leg disc

Haltere disc

Genital disc

Figure 8.5 Schematic diagram of imaginal discs from a third instar larva.

Collect late third instar larvae from either your own cultures or the stock cultures kept in the laboratory. Third instar larvae will be the largest and the ones crawling up the sides of the bottles. You could dissect imaginal discs from any of the three instar larval stages, but since the discs have been growing all through the larval stages, they will be largest in the third instar. Rinse one of the larvae in tap water, and transfer it to a drop of insect Ringer’s solution on a slide. Place the slide over ice to anesthetize the organism. Your dissection of the discs will be done directly on the slide. Everything but the discs then will be removed from the slide, and a permanent whole-mount preparation will be made of the discs that remain. Remove your slide from the ice and place it under your dissecting scope. It will be easier to dissect on a lower than higher power (trust me). Use two pairs of fine forceps. With one, hold on to the anterior end of the larva in the region of the mouthparts. Hold on firmly. With the second pair, pull on the posterior third of the larva without closing the forceps all the way. A lucky pull will break the body wall just behind the mouthparts and pull it away like a sleeve. The inner parts will be displayed for you in their correct anterior-to-posterior orientation. Use Figure 8.3 to determine what you are looking

Development of the Fruit Fly

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at. Identify the strands of trachea (tough, white, branched, stringlike structures), the various regions of the gut, the salivary glands, the fat bodies, and the many pairs of imaginal discs hanging to the tracheae. Remember, the discs will look like small, teardrop-shaped packets with a coiled inner structure. (Keep track of the salivary glands; you will be using them later for chromosome squashes.) Detach the discs using two microneedles. Draw one needle across the other in a scissorlike fashion. Keep manipulating the freed discs to a single, clean spot on your slide until you have collected as many as you can find. Remove the salivary glands to a drop of Ringer’s solution on a fresh slide using a microneedle (or pipette, but remember, pipettes have been known to swallow and never relinquish small, soft objects). Invert a petri dish over the slide containing the salivary glands so that they won’t dry out before you get to them. Return to your slide of imaginal discs and push the remaining debris off to the side. Remove this debris carefully with a Kimwipe™. Place a drop of 70% alcohol on top of the discs, and record the time in your laboratory notebook. The alcohol will start fixing the discs. Now observe the discs under the dissecting scope, identifying as many as you can (Figure 8.5). Make diagrams of the discs in your laboratory notebook. Do not let the discs dry out. Add more 70% alcohol when necessary. Eversion of imaginal discs If you have extra leg imaginal discs that you are not saving for whole mounts, and have both time and energy, you can try causing premature eversion of them by soaking them in 0.1% trypsin (an enzyme that acts on proteins) made up in Ringer’s solution and adjusted to pH 7.0. (The enzyme solution may be made ahead of time and kept frozen in small aliquots. Unfreeze only the amount you need. The enzyme quickly loses its activity at room temperatures.) You should see eversion within 5–10 minutes. Why do you think this treatment induces eversion?

Whole mount preparations of imaginal discs It is not easy to mount an entire set of discs. Be satisfied with less. Even a few discs will be adequate. The steps will involve first fixing the discs so they will not deteriorate over time, clearing them so that they are transparent, and finally mounting them in a clear, permanent mounting medium. You will be using 70% alcohol as a fixative, not because it is the best fixative, but because it will leave no toxic residues in the laboratory. If you want to stain the discs as well, a few drops of toluidine blue can be added to 10 ml of 70% alcohol; this can be used in place of straight 70% alcohol on your discs. The discs should fix for one hour starting from the time you first put alcohol on them (recorded above). Do not let them dry out during this period. Put a petri dish cover or some other suitable top over the slide to cut down on evaporation whenever you leave the slide unattended. You will be clearing the discs in glycerin. This is an excellent clearing agent and is miscible with alcohol and water. Either draw off some of the alcohol surrounding the discs (dangerous) or let some evaporate (much safer) before adding glycerin. Place a drop of glycerin over the discs, then ring the discs with glycerin jelly that has been melted in a warm-water bath. The glycerin jelly is the permanent mounting medium. Now place a footed coverslip over the discs. If there is not enough mounting medium to fill the space under the coverslip, use a pipette to add more glycerin jelly from the side. If there is too much glycerin jelly, leave it until the next lab when it will be thoroughly solid. Then you can clear away the excess by cutting it away with a razor blade.

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Kaiser’s glycerin jelly distilled water 52 ml gelatin 8 gm glycerin 50 ml 1 crystal of thymol (to retard mold)

Heat to 75°C and stir until dissolved. Do not heat above this temperature.

Observe your slide under the compound microscope, and record what you see with diagrams. You may need to amend your previous identifications, since the imaginal discs now will be much clearer then they were before. Put your slide face up in a covered box until next week. At that time, use fingernail polish to ring the edge of the coverslip, making an airtight seal. This will prevent the glycerin jelly from drying out.

Chromosome squash from salivary glands It is common to make chromosome preparations from Drosophila larval salivary glands. The reason is that these glands are soft and easily squashed and their cells are large with huge polytene chromosomes, chromosomes that replicate without separating. By the third instar, the chromosomes can have as many as 1024 chromatids. Most of the differentiated larval cells are polytene, in fact. Larval cells grow not by mitosis, but by duplicating their chromatin and increasing cell size. Only the imaginal discs, histoblasts, and gonadal cells undergo mitosis and remain nonpolytene. Use the salivary glands on the slide already set aside. Place a drop of aceto-orcein stain on the glands, and cover the slide with a petri dish to avoid evaporation. Stain for 2–5 minutes. Then, without removing any of the stain, place a 22-mm square coverslip over the glands, and with your thumb press down firmly (but not hard enough to break the coverslip). The glands must be completely squashed. Observe under a compound microscope and make a diagram of what you see. A good squash will burst open the cells and splay out the arms of the chromosomes. The banding pattern you see gives a visual map of each chromosome. Geneticists use it to precisely define the location of mapped genes. Aceto-orcein stain 1% natural orcein in a 1:1 solution of glacial acetic acid and 85% lactic acid

Warm gently (do not boil), then cool and filter.

Pupation When the third instar larva is ready to pupate, it leaves the medium, its anterior spiracles evert, its body shortens and ceases to move, and it attaches to a firm substrate (such as the side of your bottle). The cuticle then transforms into a puparium, which is initially soft and white but soon hardens, turning tan and eventually brown and brittle. Shortly after the puparium forms, the larva detaches from the inside of the puparium by molting a fourth time. Metamorphosis then takes place. Look at the sides of a culture bottle to see the white-to-brown pupal cases stuck to the side of the glass. These can be released from the glass with a needle or microknife. Examine several under a dissecting scope to see how many stages you can observe. If time allows, peel a pupa. This can be done by using superglue to glue the pupa to the bottom of a plastic petri dish. When the pupa is secure, use microknives to chip back the pupal case to uncover the tender body of the metamorphosing pupa within. It is well worth the trouble. Remember what is

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happening during pupation: the larval organs are self-destructing, and the imaginal discs and histoblasts are differentiating to form the adult. To witness this will inspire you, or at the very least change your opinion of these rough brown packages. Eclosion marks the end of pupation and the beginning of adult life. The insect cracks open the puparium anteriorly and laterally at its seams and emerges from the pupal case. It almost invariably occurs around dawn, when leaves are still damp with dew (Drosophila means “lover of dew”) and the emerging fly can unfold its new wings and harden its cuticle without the risk of desiccation. The timing of this is controlled by circadian rhythm. If the pupa misses dawn by even a few hours, eclosion will be delayed until the next morning. The dedicated among you will undoubtedly want to rise with the sun and watch this event. Use Table 8.3 to acquaint yourself with the stages of metamorphosis and stage your pupae. Table 8.3 Stages of metamorphosis in Drosophila Hoursa

Developmental event (at 25ºC)

P1 P2

0–1 1–3

P3

3–6.5

P4

6.5–12.5

P5

12.5–25

P6

25–43

P7

43–47

P8 P9 P10 P11 P12 P13 P14

47–57 57–69 69–73 73–78 73–78 78–87 87–90

P15

90–103

White puparium: wriggling stops completely Brown puparium: oral armature stops moving permanently, heart stops pumping, gas bubble becomes visible within abdomen Bubble prepupa: puparium becomes separated from underlying epidermis; bubble in abdominal region is large, causing prepupa to become positively buoyant at end of this stage (it floats) Buoyant and moving bubble: prepupa is buoyant, and bubble moves, first appearing in the posterior of the puparium, displacing pupa anteriorly, and then appearing in the anterior, displacing the pupa posteriorly. Imaginal head sac is everted and oral armature of larva is expelled Malpighian tubules migrating and white: legs and wings extend; Malpighian tubules move from thorax to abdomen and become visible as white structures in dorsal anterior abdomen Green Malpighian tubules: Malpighian tubules turn green, and dark green “yellow body” appears between the anterior ends of the two Malpighian tubules “Yellow body”: “yellow body” (actually dark green) moves back between the Malpighian tubules; transparent pupal cuticle separates from underlying epidermis; eye cup becomes yellow at its perimeter Yellow-eyed: eyes become bright yellow Amber: eyes darken to deep amber Red-eye Bald: eyes become bright red; orbital and ocellar bristles and vibrissae darken Head and thoracic bristles: head bristles, followed by thoracic bristles, darken Wings grey: wings become gray; sex comb darkens Wings black: wings become black; tarsal bristles darken and claws become black Mature bristles: green patch (the meconium–waste products of pupal metabolism) appears dorsally at posterior tip of abdomen Meconium and eclosion: tergites become tan, obscuring Malpighian tubules and “yellow body”; legs twitch; flies able to walk prematurely if puparium removed; eclosion completed

Stage

a Times start on approximately day four of larval life when the larva is still white but is no longer able to crawl. Timing is variable among individuals, and the times given are a simplification from the Bainbridge and Bownes paper.

Source: After Bainbridge and Bownes, 1981.

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Accompanying Materials Vade Mecum: “Fruit Fly.” This chapter of the CD shows the entire life cycle of Drosophila melanogaster, including mating behaviors and ways of sexing the adult and larva. The movies on development include color-codings to indicate germ layers. Tyler, M. S. and R. N. Kozlowski. 2003. FlyCycle-II. Sinauer Associates, Sunderland, MA. This CDROM is an adaptation of the 45-minute film, Fly Cycle: The Lives of a Fly, Drosophila melanogaster, 1996 by M. S. Tyler, J. W. Schnetzer and D. Tartaglia, Sinauer Associates, Sunderland, MA. This covers the life cycle of the fruit fly as well as a number of the mutants used in research. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. In several chapters you will find excellent discussions of Drosophila development, larval polytene chromosomes, imaginal discs, and metamorphosis. The diagrams and photographs throughout are extremely useful. Fink, R. (ed.). 1991. A Dozen Eggs: Time Lapse-Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Sequence 6. This shows Drosophila embryogenesis from cleavage to hatching.

Selected Bibliography Ashburner, M. et al. (eds.). 1976–1986. The Genetics and Biology of Drosophila. Academic Press, New York. This is a series of volumes of collected papers. Sophisticated and technical, it is well worth browsing through. Volumes 2a–2e, edited by Ashburner and T. R. F. Wright, concentrate on developmental and biochemical studies. Bainbridge, S. P. and M. Bownes. 1981. Staging the metamorphosis of Drosophila melanogaster. J. Embryol. Exp. Morph. 66: 57–80. This well-illustrated paper gives detailed descriptions of each stage of metamorphosis. Campos-Ortega, J.A. and V. Hartenstein. 1985. The Embryonic Development of Drosophila melanogaster. Springer-Verlag, Berlin. This is the authoritative reference on Drosophila development. It is well illustrated and explains each stage of development. Condic, M. L., D. Fristrom and J. W. Fristrom. 1991. Apical cell shape changes during Drosophila imaginal disc elongation: A novel morphogenetic mechanism. Development 111: 23–33. Though dealing primarily with the details of cell-shape changes, the paper also provides a good start on a bibliography for imaginal discs. Demerec, M. (ed.). 1994. Biology of Drosophila. Cold Spring Harbor Laboratory Press, New York. This is a facsimile edition of the original 1950 publication. It is the Drosophila bible, an excellent classic, and a comprehensive guide to the histology and development of all stages in the Drosophila life cycle. Demerec, M. and B. P. Kaufmann. 1986. Drosophila Guide. Carnegie Institute of Washington, Washington, D.C. A very inexpensive, short paperback guide to the life cycle, breeding methods, and genetic techniques for Drosophila. Doane, W. W. 1967. Drosophila. In Methods in Developmental Biology, F. H. Wilt and N. K. Wessells (eds.). Thomas Y. Crowell Co., New York, pp. 219–244. The entire text is superb, covering a number of different species and describing rearing and experimental methods for them. Hall, J. C. 1994. The mating of a fly. Science 264: 1702–1715. A lengthy review of courtship and mating in Drosophila, including details on genetics and molecular biology. Hipfner, D. R. and S. M. Cohen. 1999. New growth factors for imaginal discs. BioEssays 21: 718–720. A brief and excellent review of how imaginal discs grow during the larval stages.

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Lawrence, P. A. 1992. The Making of a Fly: The Genetics of Animal Design. Blackwell Scientific, Oxford. This book is beautifully illustrated, making it useful beyond its genetic emphasis. Leptin, M. 1994. Drosophila. In Embryos, Color Atlas of Development, J. B. L. Bard (ed.). Wolfe Publ., London, pp. 113–134. A well-written review of Drosophila development that is beautifully illustrated. Poodry, C. A. and H. A. Schneiderman. 1970. The ultrastructure of the developing leg of Drosophila melanogaster. Wilhelm Roux Arch. 166: 1– 44. This is an excellent paper on the eversion process in a leg imaginal disc. Roberts, D. B. (ed.). 1986. Drosophila: A Practical Approach. IRL Press, Oxford. This is a gold mine of well-explained techniques and general information. One chapter, “Looking at Embryos,” is particularly useful for this laboratory study. Spieth, H. T. and J. M. Ringo. 1983. Mating behavior and sexual isolation in Drosophila. In Genetics and Biology of Drosophila, Vol. 3c, M. Ashburner, H. L. Carlson and J. N. Thompson, Jr. (eds.). Academic Press, New York, pp. 223–284. This very thorough review describes each mating behavior, discusses variations among species, includes pictures of courtship behavior, and analyzes its adaptive significance. Treisman, J. E. 1999. A conserved blueprint for the eye? BioEssays 21: 843–850. A clear review of the recent evidence that the genetics of eye development in Drosophila and vertebrates share a number of common features. Wilkins, A. S. 1986. Genetic Analysis of Animal Development. Wiley-Interscience, New York. An excellent book with several descriptive chapters on the embryonic and larval development in Drosophila.

Suppliers Most biology or zoology departments have someone with a supply of Drosophila cultures “in the cupboard” (which they must constantly clean out to keep active). A kind word can often get you all the Drosophila material that you need for this lab. If the source is not in-house, however, a number of supply companies provide cultures.

Nasco 901 Janesville Ave. Fort Atkinson, WI 53538-0901 1-800-558-9595 www.nascofa.com Sells inexpensive Drosophila cultures, wild-type and mutants. Also sells Drosophila medium, though it is certainly easy enough to make your own.

Connecticut Valley Biological Supply Co., Inc. P.O. Box 326 82 Valley Road Southampton, MA 01073 1-800-628-7748. Sells stock cultures including eggs, larvae, pupae, and adults. Must order at least three weeks in advance. Also sells a number of mutant strains.

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Any good chemical supply company such as: Sigma Chemical Company P.O. Box 14508 St. Louis, MO 63178-9916 1-800-325-3010 www.sigma-aldrich.com brewer’s yeast (also from natural food stores) glacial acetic acid glycerin lactic acid natural orcein propionic acid salts for Ringer’s solution thymol

Glossary Abdomen: In an insect, the posterior-most part of the body, behind the head and thorax. Acceptance signal: In the Drosophila, the signal given by the adult female that she will accept the courtship of a male. She slows down, extrudes her ovipositor, and spreads her wings so that mating can occur. Aceto-orcein: A staining solution made of 1% natural orcein in a 1:1 solution of glacial acetic acid and 85% lactic acid. Agar: A gelatinous substance found in red algae. It can be extracted and used as a stiffening agent to produce a gel. In the laboratory, it is often used to make an agar-based culture medium. A 1.5% solution makes a solid but elastic gel when cooled to below 40ºC that will not melt again until it reaches temperatures above 85ºC. Agar is also widely used in the food industry for gelling and thickening; baking recipes may include agar instead of gelatin for thickening in making puddings, etc. Alleles: Alternative forms of a gene that occupy a specific position, or locus, on a chromosome. Anal plates: Sclerotized plates surrounding the anus. In male Drosophila, these are heavily bristled. Antenna: In an insect, the segmented appendage on the head carrying sensory receptors. Antennal lobe: Region of the brain servicing the antenna. Antennal organ: Known as Johnston's organ, this is the special sensory apparatus of the antenna found near its base. Anterior midgut: In Drosophila this is the region of the gut that is just posterior to the foregut and is formed in the embryo by an endodermal invagination. It will form the anterior midgut and gastric caeca of the larva. Arista (aristae, pl): In Drosophila, a feathery extension of the antenna through which sound vibrations can be detected. Balancer: A structure on the side of an organism that helps the organism to stabilize during flight (or swimming). In Drosophila, the balancers sit posterior to the wings and represent a modified pair of wings.

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Basement membrane: The extracellular matrix (acellular) found below any epithelium. It creates a cellular barrier between the epithelium and its underlying mesenchyme and is of major importance to the differentiation and maintenance of an epithelium. It is made up primarily of type IV collagen, laminin, fibronectin, and heparan sulfate proteoglycan. Blastoderm: The layer of cells formed in the embryo during cleavage. In Drosophila this layer is formed in the periphery of the yolky egg. In the chick, it is formed as a disc-shaped area at the animal pole of the yolky egg. Bristles: In insects, sclerotized, stout, hairlike projections from the adult cuticle, classified as two main types, larger macrochaetae and smaller microchaetae. Cellular blastoderm: In the Drosophila embryo, the stage at which the blastoderm has gone from being a syncytium to a being cellular. Cell membranes grow inward between the peripheral nuclei, separating off individual cells from the inner yolky cytoplasm. Cellularization: In the Drosophila embryo, the process during early development whereby the syncytial blastoderm is turned into a cellular blastoderm. Cell membranes grow inward between the peripheral nuclei, separating off individual cells from the inner yolky cytoplasm. Centrolecithal: Having yolk that is concentrated in the central region of the egg. Centrolecithal eggs are also macrolecithal eggs, having a large quantity of yolk. The insect egg is an example of a centrolecithal egg. Cephalic furrow: Literally, a furrow in the head region. In the Drosophila gastrula, it is a lateral infolding that occurs toward the anterior end and extends around the circumference of the embryo; it roughly delineates the boundary of the future head. Chitin: Made up primarily of sugars often complexed with proteins, it is the principle component of a cuticle. Chorion: A tough, protective eggshell. In Drosophila and other terrestrial insects, it is specially designed for maximal gas exchange with minimal water loss. In fish, it is a clear, tough enveloping layer. Chromosome: A structure in the cell nucleus made up of the genetic material, DNA, complexed with protein. Clypeolabrum: In the Drosophila, one of the head segments that is situated anterior to the mouth parts. It is broad and flat, and hangs down in front of the mandibles. Compound eye: The type of eye found in insects. It is made up of many individual facets, or ommatidia, each with its own corneal lens. Corneal lens: In the compound eye, the lens that sits at the distal end of each ommatidium. In Drosophila, it is a chitinous structure, biconvex, and approximately 5µm thick at its center. Cuticle: The hardened outer covering of flies and other arthropods, made of chitin and hardened with the tanned protein called sclerotin, forming a protective layer that prevents water loss. Cytokinesis: Division of a cell’s cytoplasm, as opposed to karyokinesis (division of a cell’s nucleus). Typically, cytokinesis follows karyokinesis. In the early cleavage stages in Drosophila, however, nuclear division (karyokinesis) occurs many times before the cytoplasm cleaves. Decamp: To walk away. A female Drosophila that is unreceptive to a courting male decamps as part of her rejection behavior. Dechorionation: Removal of the chorion. The chorion is a tough extracellular coat that surrounds the eggs of many species (e.g., Drosophila, zebrafish). A chorion can usually be removed either physically with fine forceps or chemically with an enzyme capable of dissolving the chorion (e.g., Clorox, in the case of the Drosophila chorion and pronase in the case of the zebrafish chorion).

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Denticle belt: On the ventral surface of the Drosophila larva, rows of small hooks that help the larva move by gripping the substrate. Dorsal vessel: The simple tubular heart of Drosophila. Drosophila melanogaster: The species name for the common fruit fly. Ecdysis: Molting; the process of shedding an old cuticle and forming a new one. Eclosion: The emergence of the adult fly from the puparium. Ectoderm: The germ layer in an embryo that gives rise to the epidermis and nervous system. In many cases, as in vertebrates, an anterior and posterior invagination of ectoderm gives rise to the stomodeum and proctodeum, respectively. Endoderm: The germ layer in an embryo that gives rise to the epithelium (lining) of the gut and gut derivatives. Epithelium: A major type of tissue in an organism, found lining body cavities and covering the outside of the body. The epidermis is an example of a specific type of epithelium. Esophagus: A region of the foregut, just posterior to the pharynx. Eye-antennal disc: In the fly, the imaginal disc that gives rise to both the compound eye and the antenna of the adult fly. Fat body: A sheet of fat cells that serve in site fat storage. In Drosophila these sheets are creamy white. First instar larva: In Drosophila, the larva that hatches from the egg. This feeding stage grows rapidly and soon molts into the second instar larva. Flick: A rapid movement of the wings. An unreceptive female Drosophila uses flicks when rejecting the male during courtship. Males also use wing flicks directed at other males that try to court them. Foregut: The anterior part of the gut. In vertebrates, this is formed from endoderm and includes the pharynx and esophagus-trachea regions. In insects it is formed from ectoderm and comprises the buccal (mouth) cavity, pharynx, esophagus, proventriculus and salivary glands of the larva. Gamete: An egg or sperm. Gastric caecae: In the Drosophila larva, a ring of four blind-ending fingerlike pouches that extend from the gastric region. They are secretory, as is the rest of the gastric region, and aid in digestion. Gastrulation: The stage in embryonic development that follows cleavage. During gastrulation, cells rearrange themselves, the endoderm and mesoderm cells moving inward and the ectoderm cells spreading around the outside, with some ectoderm invaginating to form such structures as the stomodeum and proctodeum. Genital disc: In the fly, the only imaginal disc that is unpaired, this disc gives rise to the genital ducts, accessory glands and external genitalia of the adult fly. Germ band: In Drosophila, created during gastrulation, this is the multilayered band of germ layers on the ventral side of the embryo that curves around the embryo's posterior tip. Germ band extension: In Drosophila, occurring during early gastrulation, this is the elongation of the germ band along the dorsal side of the embryo so that eventually the embryo's posterior end meets its head end. Germ band retraction: In Drosophila, occurring during later gastrulation, this is the shortening and thickening of the germ band which brings the posterior end of the embryo back toward the posterior pole of the egg. Glycerin jelly: A mounting medium used in making whole-mounts of specimens that have not been dehydrated. It contains glycerin, gelatin, and water. It is heated before use to make it liquid; when it cools, it becomes a firm gel.

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Gonad: The organ that forms gametes. In males, this is the testis, a sperm-forming organ, and in females this is the ovary, an ovum-forming organ. In the embryo, the gonad begins developing in the absence of germ cells, and secondarily, becomes populated by the primordial germ cells, the cells that will become the gametes. Haltere: In the fly, a fleshy, club-shaped organ found posterior to the wings and used for balance. Halteres are homologous to the hind wings of non-dipteran insects. Head eversion: In the fly, marking the transition from prepupa to pupa in Drosophila, this is the turning outward of the head capsule which had previously been turned inward. Head involution: In the fly, during gastrulation, this is the folding inward of the head region causing it to slowly disappear to the interior. Because the head involutes during development, the Drosophila larva appears superficially to be headless. Hemolymph: The oxygen-carrying and nutritive fluid, equivalent to blood, which bathes the organs of an organism such as a fly which has an open circulatory system. Heterozygous: Having two different alleles of a particular gene. Hindgut: The posterior part of the gut. In vertebrates, it refers to the endodermal gut posterior to the colon. In insects it is ectodermally derived, and forms the hindgut and Malpighian tubules of the larva. Histoblasts: In the fly larva, the population of cells that will give rise to the abdominal epidermis and internal organs of the adult. They are recognizable within the larva as clumps of small cells nestled among the large differentiated cells of the larva. Histological section: A thin slice of a tissue or organism that is typically prepared for sectioning by being fixed and embedded in a hard medium such as paraffin or plastic. Histological sections are usually stained with colored dyes. Holometabolous: A term referring to insects that have a larval and pupal stage prior to the adult stage. Drosophila is an example of a holometabolous insect. Homeotic gene: A gene whose expression determines the specification of a body region. In Drosophila, the expression of the homeotic gene Antennapedia, for example, specifies the identity of the second thoracic segment. Homozygous: Having two of the same allele of a particular gene. Humeral disc: In the fly, the imaginal disc that in the adult forms two dorsal thoracic plates, just posterior to the head, called the humerus and pronotum. In the larva this disc is found ringing the base of the anterior spiracle. Imaginal discs: In the fly, these are packets of folded epithelium that eventually differentiate into many of the structures of the adult such as the wings, legs, antennae, eyes, and proboscis. They form in the embryo as epidermal invaginations. They are carried around in the larval stages, growing in size, but do not unfold and differentiate until metamorphosis. Imago: The adult, the sexually mature stage of an insect. Instar: Any of the larval stages in an insect separated by a molt. Drosophila goes through three instar larval stages before the onset of metamorphosis. Involution: A type of gastrulation movement in which a sheet of cells folds inward and spreads over an inner surface. Kaiser’s glycerin jelly: A specific recipe for glycerin jelly, containing gelatin, glycerin, water, and a mold retardant such as thymol. It is used as a mounting medium for whole-mounts. Karyokinesis: Division of a cell’s nucleus, as opposed to cytokinesis (division of a cell’s cytoplasm). Though these two events normally both occur during cell division, there are exceptions. In the early

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cleavage stages in Drosophila, for example, nuclear division (karyokinesis) occurs many times before the cytoplasm cleaves. Labial discs: The imaginal discs that will form the proboscis of the adult fly. Labial palps: In the fly, the fleshy swollen structures on the distal end of the proboscis that make contact with the food when the adult is feeding. Larva (larvae, pl.): Early stage or stages in the life cycle, differing significantly in morphology and ecology from the adult. Love song: In the fruit fly, the courtship song, a series of sounds made by rapid movements of the male's wings which he extends and vibrates one at a time during courtship. Malpighian tubules: Tubules attached to the gut having an excretory function. They form as evaginations of the hindgut. Mandibular: Of the mandible. In insects, a mandible is one of the first pair of mouthparts, or “jaws.” In vertebrates, the mandible is the lower jaw. Maxillary: Of the maxilla. In insects, a maxilla is one of a pair of mouthparts, lying beneath the mandible. In vertebrates, the maxilla is the upper jaw. Melanin: A black pigment common in many animals. Mesoderm: The germ layer in an embryo that gives rise to the muscular and circulatory systems and most of the skeletal and urogenital systems. Metamorphosis: The transformation from the larval to the adult stage. Microknife: A knife with a small blade, suitable for microdissection under a dissecting microscope. It can be made inexpensively from chips of razor blade, a wooden dowel, and superglue. Microneedle: A needle that is very thin, suitable for microdissection under a dissecting microscope. It can be made inexpensively from inset pins, a wooden dowel, and superglue. Micropyle: Found in the eggs of certain organisms such as the fruit fly, a small channel through the chorion at the anterior end of the egg that provides an entryway for sperm at fertilization. Midgut: The region of gut between the foregut and hindgut. Molting: Or ecdysis, is the shedding of an old cuticle and formation of a new one. Mouth hooks: In the Drosophila larva, a pair of hooks that articulate with a chitinized H-shaped sclerite posteriorly. The mouth hooks are used to bring food to the mouth. Ocellus (ocelli, pl.): A simple eye containing a single light-perceiving element covered by a lens. In the adult Drosophila, there are three ocelli on top of the head. Ommatidium (ommatidia, pl.): The separate light-sensitive elements of the compound eye, each with its own corneal lens. In Drosophila each eye has approximately 750 ommatidia. Open circulatory system: A circulatory system in which blood is not confined to vessels. Blood, after leaving the heart through major vessels, enters the hemocoel, the complex of blood-filled spaces surrounding the organs. Typical of arthropods and molluscs. Operculum: Literally, a lid or cover. In Drosophila an operculum in the chorion at the anterior end of the egg provides an exit door for the larva at hatching. In the pupal stage, the operculum at the anterior end of the puparium is the door through which the adult fly will exit. Optic lobes: In the brain of the fly, lateral enlargements, flanking the central brain, that service the eyes. Ovarian follicle cells: Cells in the ovary that surround an egg, nourishing it. Ovary: The female gonad. Oviposition: The laying of an egg.

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Ovipositor: A modification at the hind end of adult female fly through which the eggs are laid. It can be extended and retracted. A female extends her ovipositor when laying an egg and also when rejecting a male during courtship. P1-P4: In Drosophila, the prepupal stages, according to the staging series of Bainbridge and Bownes (1981). P5-P15: In Drosophila, the pupal stages, according to the staging series of Bainbridge and Bownes (1981). Peripodial membrane: In an imaginal disc of the Drosophila larva, the thin, smooth wall on one side of the disc. It is the unfolded region of epithelium of the vesicle that forms an imaginal disc. Pharynx: In the Drosophila larva, the region of the digestive tract between the oral hooks of the mouth and the esophagus. Phenotype: The detectable feature in an organism that is the manifestation of a genetic trait. Plastron: A thin film of air maintained around a structure when submerged. It can act as a “physical gill.” Pole cells: In Drosophila, these are the primordial germ cells that later become the gametes. They are the first cells to form in the embryo, forming posteriorly. They are brought anteriorly by germ band extension, and then sink to the interior along with the invagination of the posterior midgut. Polyspermy: Fertilization of an egg with more than one sperm. Polytene chromosomes: Chromosomes that replicate without separating. They occur in Drosophila larvae in most of the differentiated cells, which grow throughout the larval stages, replicating their chromosomes without dividing. Posterior midgut: In the Drosophila this is the region of the gut that is posterior to the anterior midgut and is formed in the embryo by an endodermal invagination. Prepupa: In Drosophila, used to refer to the stages between pupariation and head eversion, though technically, it is the stages between pupariation and the retraction of the epidermis from the previous instar cuticle. (See Ashburner, 1989, for full discussion.) Primordial germ cells: The early population of cells that give rise to the germ cells. These are typically identifiable early in development. Proboscis: The extensible mouth parts of the adult fly. Proctodeum: The posterior-most region of the gut that is formed from an ectodermal invagination. The posterior opening created by the invagination is the anus. Proventriculus: In the Drosophila larva, a heavy-walled, muscular, bulb-like region of the gut posterior to the esophagus. It’s inner intima layer is chitinous and is used for mechanical break down of food. Ptilinum: A sac, on the front of the fly's head, which expands with blood to break the seam of the operculum when the fly exits its puparium during eclosion. Pulse song: Part of the Drosophila male's love song. It is a purring sound created by high-amplitude motions of the wing. Intervals between pulses and qualities within a pulse in the pulse song are species-specific and contribute to sexual isolation between species. Pupa: In Drosophila, used to refer to the stages between head eversion and eclosion. (See Ashburner, 1989, for full discussion.) Pupariation: Formation of the puparium. In Drosophila, this process occurs at the end of the third instar larval stage and includes shortening of the body, eversion of the anterior spiracles, and tanning of the larval cuticle. Puparium: The tanned, or hardened, larval cuticle that surrounds the organism during metamorphosis and from which the adult fly will eclose.

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Pupation: Formation of the pupa. Respiratory filaments: In the Drosophila, anterior extensions of the chorion of the egg. They have a water-repellent surface network which maintains a film of gas around them when submerged, allowing them to function as a physical gill. Salivary glands: Glands associated with the anterior end of the digestive system that secrete saliva. In the Drosophila larva, they are large and soft, and their chromosomes are polytene (as are most of the differentiated cells of the larva), making them suitable structures for creating chromosome squashes. Sclerotized plates: Hardened plates made of cuticular protein. Second instar larva: The larval stage between the first and second molts. In Drosophila it is a feeding stage that lasts 24 hours. Seminal receptacle: In the adult female Drosophila, a compactly coiled tube attached at the anterior end of the uterus that stores sperm received from the male during mating. Sex combs: Fringes of tiny black, stout bristles on the end of the first segment of the Drosophila male's front legs. They help the male pull himself up onto the female's back during mating. Sexually dimorphic: Having two sexes that are distinguishable from one another. Sine song: Part of the Drosophila male's love song. It is a humming sound created by low-amplitude vibrations of the wing. Somatic contractions: Contractions in body musculature as opposed to musculature of the gut. Sperm: The male gamete. Spermatheca (spermathecae, pl): A region of the female's reproductive tract of certain organisms where sperm can be stored. In Drosophila the pair of spermathecae are mushroom-shaped organs lying embedded in fat tissue and connected to the uterus by narrow ducts. Spiracle: An external opening to a trachea. In Drosophila larvae all instars have posterior spiracles, but only the second and third instars have anterior spiracles. In the adult the spiracles, though more numerous (there are nine pairs along the thorax and abdomen), are not as obvious. Stomodeal plate: In the embryo, the flattening and thickening of the ectoderm in the oral region in preparation for invagination to form the stomodeum. Stomodeum: In the embryo, the invaginated ectoderm in the oral region. Sucking pump: In the fruit fly, a muscular structure at the base of the proboscis that sucks food into the esophagus. Superficial cleavage: A pattern of cleavage that occurs in centrolecithal eggs, in which only the peripheral cytoplasm cleaves to form individual cells, leaving the central yolky cytoplasm uncleaved. The type of cleavage found in Drosophila embryos. Also called peripheral cleavage. Syncytium: A multinucleate tissue in which cell membranes don't completely separate the nuclei. In the Drosophila embryo, during cleavage, the blastoderm first consists of a syncytium in which many nuclei are lined up in the peripheral cytoplasm. Telson: In the Drosophila larva, a tail-like structure that is formed from the posterior-most segments, abdominal segments 9 and 10. Temperature-sensitive mutant: A mutant that exhibits a normal trait at a permissive temperature but fails to exhibit this trait if the temperature is shifted to a higher restrictive temperature. The wild-type displays the trait at both temperatures. Testis: The male gonad in which germ cells form sperm. Third instar larva: The larval stage following the second molt. In Drosophila the third instar larva feeds and then crawls out of the food source to pupariate. The stage lasts about two and a half days.

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Thorax: In the adult fly, the region between the head and the abdomen, bearing the wings, legs, and halteres. Trachea: In the fly, the tube-like invaginations of the body wall serving as air channels to allow internal tissues to exchange respiratory gases with the outside air. They open to the outside at the spiracles. In vertebrates, the tube that leads from the oral cavity to the lungs. Uterus: The portion of a female reproductive tract lying caudal to the oviducts. In the adult female Drosophila, it is a muscular unpaired structure that receives eggs from the oviducts and moves them toward the outside. Vaginal plates: In the adult female Drosophila, these are sclerotized plates surrounding the vulva. Ventral furrow: In the Drosophila embryo, this is a ventral invagination that marks the onset of gastrulation and represents the inward folding of the mesoderm. Ventral ganglion: In the Drosophila larva, it is the region of the central nervous system that is connected to the two brain hemispheres, lies ventral to them, and extends further posteriorly. Vitelline envelope: An extracellular coat surrounding an ovum. Vulva: In the adult female Drosophila, this is the opening of the reproductive tract to the outside, serving both as an entrance for sperm during mating and as an exit for eggs during egg laying. White pupa: In Drosophila this is the first prepupal stage when the larval body shortens, everts its anterior spiracles, and becomes immobile. Wing inflation: The gradual unfurling of the folded wings in the adult fly following eclosion. Blood flowing between the two epithelial layers of the wings unfurl the wings by extending and flattening them.

Fruit Fly Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. How can you distinguish an adult male from an adult female Drosophila?

2. What are the behaviors an adult male Drosophila exhibits while courting an adult female Drosophila?

3. Which is longer, a Drosophila sperm or a Drosophila egg?

4. What is unusual about the pattern of cleavage in the fruit fly egg compared to that of eggs from organisms such as sea urchin?

5. What is the first stage of gastrulation in the fruit fly?

Name: _______________________

Fruit Fly - 2

6. How does the fruit fly larva increase its size?

7. What structures do the imaginal discs form, and at what point in the life cycle do they differentiate into these structures?

8. Can you sex fruit fly larvae, and if so, how?

9. How does a fruit fly eclose from its pupal case?

10. Why are third instar larval salivary glands organs of choice for making chromosome squashes?

chapter

9

Early Development of the Chick Gallus domesticus

The common domestic chick, Gallus domesticus, is traditionally studied in embryology classes. Its development is an international language common to all students of embryology, and this alone is reason enough to learn about it. There are other reasons as well. Living chick embryos are easily obtained. Since chickens breed in any season, chick embryos often are no further away than the nearest farm. Above all, chick embryos can teach you a great deal about vertebrate development in general and will help you understand comparative vertebrate anatomy and vertebrate evolution. You will see the vertebrate body plan being laid down, learn how the organs form, and witness how they relate developmentally to one another. You will see the structures form that are diagnostic characters of the phylum Chordata and subphylum Vertebrata. And you can extrapolate from chick to human development and suddenly know a tremendous amount about what must go right for a normal baby to be born and what can go wrong that causes birth defects. Learn chick developmental anatomy well, and I promise it will serve you well. Throughout the chapter, you will find a number of questions (easy ones). Remember to record answers to these questions in your laboratory notebook.

The Chick Egg The female chicken lays an amniote egg, the same type of egg laid by other birds, reptiles, and monotreme mammals. An evolutionary invention of the early reptiles, it is one of the major changes that freed vertebrates from the water, allowing them to lay eggs on land. The design is an engineering masterpiece. An outer shell protects against physical damage and dehydration while still allowing for gas exchange. Packed within is enough food and water to survive the long journey to hatching—and a long journey it is. In the chick, it takes 21 days. Stranded out on land, the embryo cannot rely on an aquatic feeding larval stage, as do, for example, most amphibian embryos. The provisions packed within the shell must be enough to nourish all of development through to the hatching of the juvenile. In a medium-size chick egg (about 60 gm), these provisions include more protein (7.2 gm) than there is in a frankfurter; as much fat (6 gm) as is in 2 teaspoons of butter but with about seven times the cholesterol (300 mg); and about 8.5 teaspoons of water (40 gm). Despite the bad reputation of its cholesterol, the egg truly is an excellent food source. In order to permanently alter the way you look at an egg in the kitchen, you will be dissecting one in the laboratory. Obtain an egg that was collected from the wilds of a local grocery store. Examine the shell and see that one end is blunt. This marks the region of an air space that you will see upon opening the egg. Prior to opening your egg, put it briefly into hot water (water that has been brought to a boil,

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but is no longer boiling), or watch someone else do this with their egg. What happens? What does this tell you? Now open your egg very carefully into a finger bowl without breaking the yolk. Do this by giving the egg a sharp crack against the edge of the bowl, putting your two thumbs in the crack, and pulling the two halves apart, with the egg close to the bottom of the finger bowl. If you are lucky, you have kept the chick ovum (the yolk) intact. No wonder flipping a fried egg without bursting the yolk is an art—that yolk is one cell. It’s huge! Only a fragile cell membrane overlain by a vitelline envelope protects it from the slice of your spatula. Compare the yolk to a more typically sized cell—a piece of dandruff, for example, which represents a single epidermal cell. The difference is remarkable! Most of the bulk in the chick ovum is not cytoplasm, but spheres of yolk lipid suspended in a sea of yolk protein. The yolk is so disproportionately abundant compared to the cytoplasm that the cytoplasm is displaced to the animal pole and sits as a puddle on top of the yolk. The large quantity of yolk classifies the egg as macrolecithal (macro, large; lecithal, yolk). Examine the eggshell. Ninety-eight percent of it is calcite, which is calcium carbonate. Calcium is the major component of bone as well (in the form of calcium phosphate, or apatite). The calcium that goes into the shell comes both from the hen’s diet and from her bones. Weigh the shell. Realize that a truly remarkable hen can lay an egg a day. Approximately how much calcium will a one-a-day hen lose in a week? In a year, a hen can put into her eggshells over 25 times her own skeletal weight in calcium. Obviously, this must be replaced. The standard feed given to laying hens is 3 to 4% calcium, and one hen receives about a 100 gm of feed a day. Based on the weight of the shell, is this sufficient to replace the calcium lost in a superhen laying every day? Examine a piece of shell under a dissecting scope, turning it so that you can see it in cross section. Notice that it has three layers (Figure 9.1): two calcified layers, the inner mammillary layer and the outer spongy (or crystalline) layer; and outermost the thin shiny cuticular membrane. In the mineralized layers, calcium carbonate is laid down as crystals perpendicular to the surface of the egg—this gives the shell tremendous strength. The outer cuticular membrane is made of glycoprotein. Prior to laying, it is wet and slippery, which helps with oviposition, but soon after laying it dries out and serves to protect against invasion by microorganisms.

Thin albumen

Blastodisc

Thick albumen Chalaza

Figure 9.1 Schematic diagram of a chick egg.

Dark yolk Light yolk

Air space

Inner shell membrane

Mammillary layer Spongy layer

Shell membranes Cuticular membrane

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Notice the shell has numerous pores. Use food coloring to dye the shell. When the dye dries, a greater amount will have collected in the dips of the pores, and these will appear as dark spots. Notice that the dark spots are irregularly sized and spaced, and that they are more abundant in the area over the air space than anywhere else. Can air pass through these pores? How can you tell? Think back to what happened when you put the egg into hot water. If air could not pass through the shell, what would happen to the embryo? (Write your answers in your laboratory notebook.) Wherever air can pass, water vapor will follow. Balancing the need for gas exchange against the risks of dehydration while retaining strength is always a major task in shell design. On the inside of the shell you should see two shell membranes, an inner and outer one, separated from one another at the region of the air space (air cell) at the blunt end of the egg (Figure 9.1). Peel the two membranes apart from one another. Which of the two membranes is thicker? Look at a piece of each membrane under the dissecting scope, then stain them with 0.1% toluidine blue and mount each on a slide with a coverslip using water as mounting medium. You will see that the membranes are a dense, crisscrossing mat of fibers. These elastin-like fibers retard water loss, give mechanical support, and provide a defense against microbes. Where else do you find a high proportion of elastin? (In ligaments: attaching bone to bone, they must be able to s-t-r-e-t-c-h; in the cartilage of the external ear: pull on your ear—does it droop, or bounce back into shape?) What does this tell you about the mechanical properties of the shell membranes? Experiment to see how much stretch the membranes have. Look at the air space at the blunt end of the egg. This space forms between the two shell membranes after the egg is laid. As the egg cools from the internal temperature of the hen (about 41°C) to room temperature, the internal contents contract more than the shell, creating the air space. Try cooling a warm egg by putting it into the refrigerator, and see if the air space becomes even larger. Do you think the air space serves a function? Could it help to absorb shock when the egg is jarred? When the chick begins to hatch, it first breaks through into the air space using its beak and fills its lungs with air. The confined space soon becomes fouled with carbon dioxide, which triggers the chick to start breaking through its shell (a process referred to as pipping, a word most of us have heard without knowing its meaning. (In the folk song “I Gave My Love a Cherry,” one verse goes, “A chicken when it’s pipping, it has no bone.” Does this make any sense?) Now look at the egg albumen. Notice that it has varying viscosities. Around the yolk is a narrow band of thin albumen, surrounded by a layer of thick albumen, and external to this another layer of thin albumen (Figure 9.1). The albumen is 88% water and is a major source of water for the developing embryo. The rest is primarily glycoproteins, the most abundant of which is ovalbumen. Some of the proteins in the albumen serve antibacterial functions: lysozyme, for example, is an enzyme that disrupts bacterial cell walls; ovotransferrin binds iron and avidin binds the vitamin biotin, making these nutrients—which are necessary for bacterial growth—unavailable to the bacteria. Two other proteins, ovomucin and cystatin, are thought to have antiviral activity. In addition, the albumen has a pH that prevents bacterial growth. What pH is this? Measure the pH using pH paper. (Did your parents, when making lemon meringue pie, ever tell you not to lick the meringue bowl because of the raw egg whites? What they were worried about was the avidin, which can cause a vitamin deficiency called egg white disease. Explain. Why are cooked egg whites all right to eat?) Notice that part of the albumen is very dense and is coiled into two cordlike structures attached tightly to the yolk membrane. These are the chalazae (pronounced ka-lay-zee) (Figure 9.1). They suspend the yolk in the middle of the albumen and allow it to rotate. The yolk will rotate, always orienting to gravity so that its animal pole (and the embryo) face upward. Why do you think this is important? (By definition the animal pole of an egg is the pole where the polar bodies form. In the chick, this is also where the blastodisc is. The vegetal pole is opposite the animal pole.) Remove the

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chalazae using scissors and fine forceps and place them side by side on a slide, keeping track of which end was attached to the yolk. Do the fibers twist the same way? Add a drop of 0.1% toluidine blue stain to help distinguish the fibers. You probably noticed that one chalaza twists clockwise and the other counterclockwise. This twisting occurs during egg formation as the yolk is slowly rotated during its descent through the oviduct. This twists the fibers of the chalazae and draws them taut, pulling the yolk to the center of the egg. You can visualize this using a cork, two rubber bands, and thumbtacks. Attach the rubber bands with thumbtacks to opposite ends of the cork. Holding the free ends of the rubber bands, slowly rotate the cork as the yolk would rotate coming down the oviduct. Notice that the rubber bands twist in opposite directions, just as the chalazae do. Notice also that as the slack in the rubber bands is taken up, the cork is suspended between the two ends. Unlike rubber bands, however, the chalazae do not tend to unwind. Using microneedles, try to tease apart the fibers of the chalazae and notice how resistant they are to untwisting. Examine the yolk. Where does the yolk sit in relation to the albumen? Does it float, or is it submerged? Remember that bowl full of eggs before you made your last omelet. Did the yolks float or sink in that sea of whites? What does this tell you about the relative densities of yolk and albumen? About the importance of the chalazae? Notice the blastodisc, the puddle of cytoplasm that should be oriented upward. It is here that the egg nucleus is found. With forceps and fine scissors, remove the blastodisc by puncturing the vitelline envelope and cell membrane with the forceps, and holding on to both with the forceps while cutting around the blastodisc with the scissors. Carefully lift the blastodisc to a slide. A disc of white yolk will be attached to the underside of the cytoplasm. Examine the blastodisc under the dissecting and compound microscopes. Remember that the nucleus is partway through a meiotic division, so there is no nuclear membrane, and the chromosomes are aligned on a metaphase plate; they are very difficult to detect. If you see a spot of blood in the yolk, this can be either an indication of a developing embryo (it would have to be in the location of the blastodisc—look at it carefully) or simply the incorporation of a blood clot from the ovary at the time of ovulation. The yellow color of the yolk is due to carotenoid pigments—the same pigments seen in carrots and other yellow vegetables. (Your body uses carotenoids to make vitamin A, which has a long list of functions. When transformed into retinal, for example, it becomes part of an eye pigment, rhodopsin, necessary for vision; when transformed into retinol, it is needed for skin maintenance. Egg yolk has about one-ninth as much carotinoid as carrots.) The carotinoid pigments are absorbed from the hen’s food and deposited rapidly in the yolk. Consequently, greater amounts are deposited during the day when the hen is eating than at night when the hen is sleeping. This results in the yolk having concentric layers of dark and light yolk (Figure 9.1), and provides us with a daily record of the yolk deposits. You can see this by removing an unbroken yolk with a spoon to some boiling water and boiling it for 10 minutes. Then, using a razor blade or sharp scalpel, carefully bisect the yolk through the middle, starting at the animal pole where the blastodisc is. Count the rings. Each sequence of a light and dark ring equals one day. How many days did it take the hen to form your yolk? Remember when and where the yolk was being packed into the ovum (in the ovary prior to ovulation).

The Female Chick Reproductive Tract The female chick has only one fully developed ovary and oviduct (usually the left), while the other remains vestigial. Why do you think this is so? If your laboratory has a female chick reproductive tract on display, examine this, using Figure 9.2 to study each region discussed below.

Early Development of the Chick

OVARY

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Mature ovum

Cloaca Fimbria Vagina Infundibulum Uterus (shell gland)

OVIDUCT Egg moving through oviduct

Isthmus

Magnum

Figure 9.2 Reproductive tract of a female chick.

Look at the ovary. Those large lumps are oocytes developing to maturity. These are ovulated as secondary oocytes, when the egg is in metaphase of its second meiotic division. Meiosis is not completed until after fertilization. Fertilization takes place high in the oviduct, in the region called the infundibulum. Why would it be difficult for fertilization to occur farther down in the oviduct? After ovulation, the egg remains fertilizable for only about 40 minutes. This does not mean, however, that mating must have occurred near the time of ovulation. Sperm can be stored in the female tract and remain viable there for as long as 35 days, though viability is reduced after the third week. Look back at the slides of sperm from an earlier laboratory session to recall the shape of chicken sperm. At the time of fertilization, many sperm enter the egg, but only one sperm pronucleus fuses with the egg pronucleus. This unusual phenomenon is called physiological polyspermy. Notice that the mouth of the oviduct is not attached to the ovary. It is not a given that every egg that is ovulated successfully enters the oviduct. In fact, as many as 5% of eggs ovulated normally miss the opening and fall into the body cavity, where they are usually absorbed. (A hen that does this a high percentage of the time is called an “internal layer.”) Once in the oviduct, the ovum packed with yolk is moved along by muscular contractions of the oviduct. After passing through the infundibulum, albumen begins to be secreted around the yolk. Most of the albumen is secreted by cells in the region of the oviduct called the magnum. The egg passes from the magnum through the isthmus, where the chalazae and the inner and outer shell membranes are added. In the uterus, also called the shell gland, water and salts traverse the shell membranes and are added to the albumen, plumping out the shell membranes. The shell is then deposited, and the egg travels on through the vagina and into the cloaca.

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Here the egg is turned completely around. Having traveled with its small end first, its orientation is now reversed, and the egg is laid with the large blunt end first. The entire process from ovulation to laying takes about 24 hours, the major portion of which (about 19 hours) is spent in the shell gland. If the egg was fertilized, development of course has been taking place all this time, and the embryo is already undergoing gastrulation by the time it is laid.

Cleavage Cleavage stages cannot be observed in the living egg under normal conditions, since they occur while the egg is still in the oviduct. You will be studying cleavage, therefore, by examining models and diagrams of the events (Figures 9.3 and 9.4). Remember the function of cleavage. The fertilized egg already contains within its cytoplasm all of the germ-layer components. Cleavage divides this single cell into a lot of smaller building blocks that then can be rearranged and molded into the multicellular organism. Use the models and diagrams to identify all of the structures printed in boldface in the next two paragraphs. The first thing you should notice is that the entire egg does not cleave; only the puddle of cytoplasm sitting on top of the yolk cleaves. Dense yolk creates a formidable impediment to a cleavage furrow. You will see this later in the mesolecithal egg of the amphibian, in which the moderate amount of yolk concentrated in the vegetal half slows down the cleavage furrows. Here in the chick egg, however, the huge amount of yolk stops the cleavage furrows altogether. Cleavage initially, therefore, is necessarily partial, or meroblastic. Cleavage furrows, which cut down from the animal pole toward the yolk, stop when they hit the yolk. The cleavage pattern in the chick is also often called discoidal, since it is restricted to the circular disc of cytoplasm, the blastodisc. The cells produced during cleavage are called blastomeres, and together they make up what is called the blastoderm. In the center of the blastoderm, the cells become separated from the underlying yolk by a space called the subgerminal cavity. The space makes the central area of blastoderm look lighter and more translucent than the surrounding area. This central region is called the area pellucida (a pellucid stream is transparent; a pellucid explanation is clear and easily understood). The surrounding area of blastoderm, still connected to the underlying yolk, looks darker and opaque; it is called the area opaca.

Figure 9.3 Early cleavage stages in the chick. The blastoderm is shown removed from the yolk and viewed from above, starting with the 2-cell stage and showing progressively more advanced stages.

Early Development of the Chick

Subgerminal cavity

Yolk

Area pellucida

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Area opaca

Figure 9.4 Late cleavage stages in the chick. The animal pole of the egg is shown in crosssection. This stage occurs within the oviduct, 3–4 hours prior to laying.

Gastrulation By the time the egg is laid, the embryo is undergoing gastrulation. The cell movements during this stage have been traced by scientists working with great patience and steady hands. The normal procedure is to label tiny regions of the blastodisc with vital dyes and other markers, to watch the paths of movement these markers make during gastrulation, and to determine where they eventually end up, thereby determining the final fate of the cells. In this way, maps have been constructed showing the location of each germ layer prior to gastrulation and its route of travel during gastrulation. These maps, called fate maps, are a gift to any experimenter, providing a blueprint for a vast array of experiments. As you study them, think of experiments you would do using the fate maps as a ground plan. Briefly outline one of your experiments in your laboratory notebook. To begin your study of gastrulation, first remember what gastrulation achieves. It separates the germ layers so that they are in appropriate positions for the organ formation that will follow. Since endoderm forms the epithelium of the gut and gut derivatives, it must become a centrally located tube. The ectoderm, which forms the epidermis and nervous system, must cover the outside. And in between must come all the packing material provided by the mesoderm—muscle, skeleton, circulatory system, kidney, and dermis. Now look at the fate map of the chick blastoderm (Figure 9.5). Notice that it is laid out flat as a pancake, a configuration forced upon it by the huge inert yolk. This is a problem. Embryos such as those of sea urchins and amphibians don’t face this problem. Because they form a hollow sphere as blastulae, they are able to gastrulate by simply moving the mesoderm and endoderm inside the sphere and then arranging these germ layers appropriately within the sphere. But in the chick, what is a sphere in these other organisms is laid out flat, and the germ layers are forced to separate in a flat plane forming a three-layered sandwich with endoderm on the bottom, ectoderm on top, and mesoderm in between. The flat germ layers later get tucked and folded to form the gut tube and other structures. Meanwhile, the entire sandwich is growing outward over the surface of the yolk and eventually surrounds the yolk. It is only the central region of the gastrula sandwich that will form the body of the embryo. The outer regions will form extraembryonic membranes: the amnion, chorion, allantois, and yolk sac. (The amnion and chorion serve to protect the embryo; the allantois stores nitrogenous waste and where it is attached to the chorion provides a vast respiratory surface that also resorbs calcium from the eggshell for the growing embryo; the yolk sac absorbs yolk and transports it back to the embryo in its vitelline blood vessels. You will see these in later chapters.)

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Epidermal ectoderm Neural ectoderm

Notochordal mesoderm

Somite mesoderm Gut endoderm Lateral plate mesoderm

Primitive streak

Figure 9.5 Fate map of the chick gastrula. The blastoderm, at about the time of laying, is shown from above so that only the epiblast is seen. The extraembryonic endoderm has already delaminated to form the hypoblast and can not be seen in this view.

Look at the blastoderm fate map again, and imagine how you would orchestrate the gastrulation movements to create a three-tiered sandwich. Now look at Figures 9.6 and 9.7 and at the models of chick gastrulation to see how close your plan of gastrulation came to what actually happens. The first step in chick gastrulation is the splitting off of the major bulk of endoderm from the underside of the blastoderm to form a layer of endoderm below the rest of the cells. This splitting is called delamination. What molecular mechanisms must be involved in delamination? Obviously, the endoderm must lose its stickiness to the rest of the blastoderm; it must change its affinity by changing its cell surface adhesion molecules. If you were able to conduct experiments, how might you test for this? Delamination produces an upper epiblast, consisting of the presumptive ectoderm, mesoderm, and some endoderm; and a lower hypoblast, consisting of the extraembryonic endoderm. The extraembryonic endoderm will participate in forming extraembryonic membranes. The presumptive endoderm remaining in the epiblast, the embryonic endoderm, will form the endodermal derivatives of the embryo itself, but it must first join the hypoblast before doing so.

Area pellucida

Blastocoel

Yolk

Area opaca

Epiblast

Hypoblast

Figure 9.6 Early gastrula stage in the chick. The animal pole is shown in cross-section. The extraembryonic endoderm has delaminated to form the hypoblast lying below the epiblast. Compare with Figure 9.4.

Early Development of the Chick Figure 9.7 Mid-gastrula stage in the chick. A three-dimensional view of the anterior half of a gastrulating chick shows the direction of movement of the cells. Embryonic endoderm and mesoderm are ingressing through the primitive streak. The endoderm is migrating down to join the hypoblast, and the mesoderm is migrating outward between the epiblast and hypoblast, forming an intermediate layer.

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Epidermal ectoderm Hensen's node

Neural ectoderm

Primitive streak Epiblast

Mesoderm

Yolk

Embryonic endoderm

Hypoblast

The embryonic endodermal cells move to the hypoblast by turning inward along a line of ingression and then moving downward to the hypoblast. The line along which cells are moving inward from the epiblast, or ingressing, is called the primitive streak. The prospective mesoderm cells are the next to enter the primitive streak and leave the epiblast. However, they do not move down to join the hypoblast but instead make a U-turn and migrate outward between the epiblast and hypoblast to form a separate layer between the two. Compare this type of gastrulation to that of a sea urchin or amphibian. What in the sea urchin or amphibian would be analogous to the primitive streak? Yes, the blastopore. Remember the dorsal lip of the blastopore in an amphibian gastrula? It is where notochordal mesoderm cells involute to form the notochord. There is an analogous structure in the chick embryo at the top of the primitive streak called Hensen’s node (or primitive node). It is here that the notochordal cells converge, ingress, and migrate forward, rather than laterally, to form a streak of notochordal cells down the midline. Look at various stages of gastrulation, noticing that gastrulation begins in anterior regions first and progresses to more and more posterior regions. Anterior regions may have finished gastrulating and gone on to organ formation while posterior regions are still gastrulating. This anterior-to-posterior wave of progression results in the primitive streak becoming confined to ever-more posterior regions. This is often referred to as the regression of the primitive streak.

24-Hour Chick Whole Mount Molding a body out of the three-tiered sandwich of a chick gastrula is a task that takes a lot of folding and pinching. The beginnings of this process are well illustrated in a 24-hour chick embryo (by tradition, this means an embryo incubated for 24 hours—total time of development has been more than 24 hours, since development began prior to egg laying) (Figure 9.8). You will be looking at whole mounts in which the cellular region has been removed from the yolk and mounted on a slide. Because of the transparency of the embryo at this stage, all levels can be seen by focusing up and down through the embryo. Use both your dissecting and compound microscopes, experimenting with different settings for

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Proamnion Area vasculosa

Anterior neuropore

Neural tube Foregut Head fold Notochord Somites

Open neural plate

Hensen's node

Primitive streak

Figure 9.8 Schematic diagram of a 24-hour chick embryo.

Lateral plate mesoderm Unsegmented somite mesoderm Area pellucida

Area vitellina

contrast and lighting. Be able to identify any of the structures printed in boldface below, and write in your laboratory notebook answers to any questions posed. Realize that you yourself looked very much like this embryo at one time (about three weeks into your development). By 24 hours of incubation, the anterior half of the embryo is undergoing neurulation while the posterior half is still gastrulating. Notice that the area opaca has greatly expanded. In fact, it now occupies too large of an area to be mounted on your slide, so much of it already has been trimmed away. Even so, you still should be able to identify the two subdivisions of the area opaca: the outer area vitellina (vitellina means yolk), where cells contain many yolk granules; and the inner area vasculosa (referring to vascularization), where blood islands are forming. Blood islands are masses of blood-forming cells that will later anastomose to form a capillary network that will bring yolk nutrients back to the embryo proper. Look at the area pellucida, the clear area around the embryo; it looks almost like a footprint. Just in front of the embryo’s head is a region of the area pellucida that is particularly clear. This is called the proamnion (an unfortunate name whose meaning, “before the amnion,” has nothing to do with the structure—suggest a better name and we’ll vote on it). The reason the proamnion is so translucent is that it consists only of ectoderm and endoderm. The spreading mesoderm has not yet reached this area. Focus now on the embryonal area. The folding and pinching has already begun. At the anterior end you will see a head fold. This is where an anterior fold has undercut the developing head and raised it

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above the level of the blastoderm. Look at the models on display to see this more clearly. Notice how the head fold has caused the endoderm to fold into a closed tube to form the foregut. Posteriorly, where the endoderm is still flat, is the open midgut, and the transition between the closed foregut and open midgut is the anterior intestinal portal (portal means door). This is best visualized by looking at models. Now look carefully on your whole mount for the darker outline of the foregut and the crescentshaped line of the anterior intestinal portal. Look at models to see the mesoderm lying directly underneath the foregut and lateral to it. This mesoderm will be forming the heart later on and is therefore called the cardiac mesoderm. On your whole mount, it should look somewhat darker than the surrounding region. One of the most notable features of the 24-hour embryo is the invaginating neural ectoderm. Previously it was an open neural plate, but by 24 hours, the anterior region has folded upward to form a closed neural tube. A major part of this folding is caused simply by a change in shape of the neural plate cells from cuboidal to truncated-pyrimidal. Convince yourself of this by making a sheet of cells (neural plate) out of small cubes of clay. Now change the shape of each cube by pinching its apical end (a contraction due to an apical belt of actin and myosin microfilaments in the cells of the neural plate) and elongating it along its apical-basal axis (caused by an elongation of microtubules in this axis). Now fit the clay cells back together, and you will see that your sheet of cells has folded upward. Look at your whole mount again. Notice that neurulation is taking place earlier anteriorly than posteriorly. Identify regions of closed neural tube (if any), open neural plate, and in between regions where neural folds have formed but have not closed. Notice that at the anterior tip of the embryo, the neural plate does not close completely but leaves an open channel called the anterior neuropore. This will be closed by 36 hours. (Sometimes the neural folds in the spinal region fail to close during development, causing an extremely severe birth defect called spina bifida with myeloschisis. Occurring in humans as well as chicks, vertebrae do not form properly in the area, and the spinal cord is left an unprotected, flattened mass of neural tissue.) One thing you cannot see in the whole mount, which you will see later in serial sections, are the neural crest cells lying on top of the neural tube. As soon as the tube closes, they start migrating away from the neural tube, sometimes to distant regions. These are fascinating cells and will be discussed more completely in the next chapter. For now, just keep in mind that they are unique to the vertebrates and form a variety of structures including cartilage and membrane bones of the head, pigment cells of the skin, and spinal ganglia. Look at the open midgut region, and notice that the mesoderm has segmented into blocks of somites on either side of the neural tube. Count and record the number of somites your embryo has. As the embryo develops, additional somites will form from the more posterior unsegmented somite mesoderm. Recall what the somites will be forming: the axial skeleton (vertebrae and ribs), muscles of the axial skeleton and limbs, and dermis. Between the two rows of somites, in the midline underneath the neural ectoderm, you should see a streak of condensed tissue. This is the notochord. How far does it extend anteriorly and posteriorly on your embryo? At the posterior end of the embryo, notice the region of the primitive streak and Hensen’s node, showing that ingression is still occurring in this region. Can you think of any reason why an embryo would spend more time making its anterior end than its posterior end? Now that you are thoroughly familiar with your embryo, you can stage it more accurately than simply calling it a 24-hour embryo. You can use the staging series developed by Hamburger and Hamilton; a truncated version of this series is given in the next chapter. This is the most widely used staging series for the chick. Record the stage of your chick. By convention, when using this staging series, you would precede the stage number with the initials HH.

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Accompanying Materials See Vade Mecum: “Chick Early.” This chapter of the CD shows step by step how to open the egg and identify its parts. It shows early development in a series of movies developed from models and histological whole mounts. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. Contains chapters with clear descriptions and illustrations of the amniote egg, chick cleavage and gastrulation, and a discussion of the mechanisms of the gastrulation movements. Fink, R. (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Sequence 11. This shows early chick development, allowing you to see neurulation and regression of the primitive streak.

Selected Bibliography Burley, R. W. and D. V. Vadehra. 1989. The Avian Egg: Chemistry and Biology. John Wiley and Sons, New York. Full of tables and graphs, this book includes every statistic about an egg that you probably ever could use. Deeming, D. C. and M. W. J. Ferguson (eds.). 1991. Egg Incubation: Its Effects On Embryonic Development in Birds and Reptiles. Cambridge University Press, Cambridge. This is a collection of papers from an international meeting. The comparisons between bird and reptile are fascinating. For example, did you know that although bird eggs require turning during incubation, turning a reptilian egg usually kills the embryo? Diaz, C., L. Puelles, F. Marín and J. C. Glover. 1998. The relationship between rhombomeres and vestibular neuron populations as assessed in quail-chicken chimeras. Dev. Biol. 202: 14–28. This is an excellent example of combining the more modern techniques of immunohistochemistry with the older technique of using chimeras in fate mapping. Hamburger, V. and H. L. Hamilton. 1951. A series of normal stages in the development of the chick embryo. J. Morphol. 88: 49–92. This is the original printing of the Hamburger-Hamilton chick staging series. It contains an exquisite set of photographs and drawings along with the written description of each stage. North, M. O. 1972. Commercial Chicken Production Manual. Avi Publishing Co., Westport, CT. A warhorse of a manual, this will stand by you with ready answers to a host of practical questions about chick eggs and their incubation. Romanoff, A. L. and A. J. Romanoff. 1949. The Avian Egg. John Wiley and Sons, New York. This is a classic. Though some of the biochemical information is now outdated, the wealth of anatomical details is well worth the task of carrying such a heavy book around. Selleck, M. and C. Stern. 1991. Fate mapping and cell lineage analysis of Hensen’s node in the chick embryo. Development. 112: 615–626. A superb look at how to do classic fate mapping with modern techniques. Spratt, N. T., Jr. 1946. Formation of the primitive streak in the explanted chick blastoderm marked with carbon particles. J. Exp. Zool. 103: 259–304. This is a classic paper showing the original ways in which fate maps were determined. Stern, C. D., 1994. The chick. In Embryos, Color Atlas of Development. J. B. L. Bard (ed.). Wolfe Publ., London, pp. 167–182. An excellent review of early chick development that includes a discussion of fate mapping techniques.

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Tuan, R., 1987. Mechanisms and regulation of calcium transport by the chick embryonic chorioallantoic membrane. J. Exp. Zool. [Suppl.] 1: 1–13. This paper demonstrates that calcium needed for bone formation in the chick embryo is resorbed from the eggshell by the chorioallantoic membrane and brought back to the embryo by the allantoic blood vessels.

Suppliers Prepared microscope slides of chick embryos can be obtained from suppliers such as: Turtox/Cambosco, Macmillan Science Co., Inc. 8200 South Hoyne Ave. Chicago, IL 60620 1-800-621-8980

Connecticut Valley Biological Supply Co., Inc. P.O. Box 326 82 Valley Road Southampton, MA 01073 1-800-628-7748

Ward’s Natural Science Establishment, Inc. 5100 West Henrietta Road P.O. Box 92912 Rochester, NY 14692-2660 1-800-962-2660 www.wardsci.com Models of chick development are available from Turtox, listed above.

Glossary Actin microfilaments: The thin microfilaments found in the cytoplasm of a cell. Made up of G-actin, the globular protein subunits. The filamentous form of actin is called F-actin. F-actin forms an extensive cytoskeletal network in the cortical cytoplasm. Myosin (the thicker of the two types of microfilaments) can move along F-actin, creating a contractile force. Air cell: In the chick egg, the air space at the blunt end of the egg between the inner and outer shell membranes. It provides a cushion for the developing embryo as well as a the air for the chick’s first breath during hatching. Albumen: Egg white. The albumen of a chick egg is 88% water and is a major source of water for the developing embryo. The rest is primarily glycoproteins. Allantois: The extraembryonic membrane found in amniotes that forms off the hindgut of the embryo. It stores nitrogenous waste and in the chick will eventually fuse with the chorion to form the chorioallantoic membrane. It is vascularized and is formed from the splanchnopleure, a layer made up of endoderm and splanchnic lateral plate mesoderm. Amnion: The extraembryonic membrane found in amniotes that surrounds the embryo. It contains amniotic fluid, which protects the embryo by providing an isotonic fluid and buffering the embryo

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against being jarred. It forms from the somatopleure, a layer made up of epidermal ectoderm and somatic lateral plate mesoderm. Amniotes: Vertebrates that have an amnion during development: reptiles, birds, and mammals. Amniote egg: The type of egg typical of the amniotes, and an egg that can survive on land. It consists of a hard outer shell that protects the embryo from physical damage and dehydration, and a large yolk along with a watery albumen that provide the nutrients and water necessary until hatching. The embryo develops four extraembryonic membranes, the chorion, amnion, yolk sac, and allantoic membranes. Anamniote: Vertebrates that do not have an amnion during development: the fishes and amphibians. Animal pole: The point on the surface of the egg where polar bodies are given off. Eggs usually orient to gravity with their animal pole oriented uppermost. Anterior intestinal portal: In the chick embryo, the opening between the closed foregut and the open midgut. The opening continues to shift posteriorly as development proceeds. Anterior neuropore: In the chick embryo, the opening at the anterior end of the brain. It later closes. Area opaca: In the chick embryo, the outer extraembryonic region that is still connected to the underlying yolk. Area pellucida: In a chick embryo, the inner extraembryonic region that is separated from the yolk by a subgerminal cavity. This area looks lighter and more translucent than the surrounding area because of the space below it. Area vasculosa: The inner region of the area opaca where blood islands are forming. Area vitellina: The outer region of the area opaca where cells contain many yolk granules. (Vitellina refers to yolk.) Avidin: An enzyme that inhibits bacterial growth by binding the vitamin biotin, making it unavailable to the bacteria. It can be found in the albumen of a chick egg. Biotin: One of the B vitamins, it is water-soluble and sulfur-containing. It is important for a number of enzyme functions and for healthy nails, hair, digestive enzymes and sugar metabolism. It is destroyed by avidin, found in raw egg white. Blastoderm: The layer of cells formed in the embryo during cleavage. In Drosophila this layer is formed in the periphery of the yolky egg. In the chick, it is formed as a disc-shaped area at the animal pole of the yolky egg. Blastodisc: In a chick egg, the puddle of non-yolky cytoplasm that sits atop the vast quantity of yolk. The nucleus of the egg sits in the blastodisc, and it is here that development begins. Blastomeres: The cells that are created through cleavage during early embryonic development. Blastopore: In a gastrulating embryo, the point at which cells are moving inward to form the archenteron. In deuterostomes, the blastopore becomes the anus, in protostomes, the mouth forms at or near the blastopore opening. Blastula stage: The stage in embryonic development following cleavage and preceding gastrulation. The embryo may form a hollow blastula (coeloblastula) or a solid blastula (stereoblastula). The cavity formed in a coeloblastula is called the blastocoel. Blood islands: In the chick embryo, the beginnings of blood vessel formation out in the area vasculosa. These are masses of blood-forming cells that will later anastomose to form a capillary network that will bring yolk nutrients back to the embryo proper. They form from splanchnic lateral plate mesoderm. Calcite: Calcium carbonate (CaCO3). One of the commonest minerals and is found in a variety of crystalline forms. It is the form of calcium found in the egg shell of a bird. CAM graft: A chorioallantoic membrane graft.

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Cardiac mesoderm: The mesoderm that will form the heart. Carotenoid pigments: The yellow-orange pigments found in leafy-green, yellow-to-red vegetables, from which the body makes vitamin A. The yellow color of yolk is from carotenoid pigments, derived from the bird’s food. Chalazae: In a chick egg, the portion of albumen that is very dense and coiled into two cordlike structures that attach tightly to the yolk membrane. The chalazae suspend the yolk in the middle of the egg and allow it to rotate. Chordata: The phylum that includes the vertebrates and two invertebrate groups, the urochordata and the cephalochordata. Diagnostic features of the phylum are the presence of a notochord (hence its name), a dorsal hollow nerve cord, and the presence of pharyngeal pouches during development. Chorioallantoic membrane: The fused chorion and allantoic membranes. In the chick, this serves a major function in respiration. It becomes very large, and presses against the inner aspect of the shell, allowing for efficient gas exchange. Chorioallantoic membrane graft: A graft placed on the chorioallantoic membrane of an embryonic chick. The highly vascularized membrane makes an excellent and inexpensive incubator for growing embryonic tissues. Chorion: The extraembryonic membrane found in amniotes that lies outside of the amnion. It protects the embryo, and, in the chick will, eventually fuse with the allantoic membrane. It forms from the somatopleure, a layer made up of epidermal ectoderm and somatic lateral plate mesoderm. Cleavage: The stage in embryonic development, normally following fertilization, in which the egg, or portions of the egg, divide by mitosis into a number of cells. Cleavage results in the formation of the blastula. Cloaca: Literally, means sewer. In a chordate, the terminal region of the endodermal hindgut. In general, it receives the ducts and products from the gonads, and the digestive and excretory organs. Cuticular membrane: The outermost shiny membrane of a chick egg shell. It is made of glycoprotein that, prior to laying, is wet and slippery, thereby helping with egg laying. Soon after laying, it dries out and serves to protect against invasion by microorganisms. Cystatin: A protein that is thought to have antiviral activity. It can be found in the albumen of a chick egg. Delamination: A type of gastrulation movement in which a multilayered sheet of cells splits into two sheets. In the chick embryo, at the beginning of gastrulation, delamination separates the blastoderm into two layers, an upper epiblast and a lower hypoblast. Deuterostome: Among the bilaterian organisms, the phyla in which during embryonic development the blastopore becomes the anus. These include the echinoderms, hemichordates, and the chordates (urochordates, cephalochordata, and vertebrates). Discoidal cleavage: The pattern of cleavage found in certain macrolecithal eggs such as the chick egg, where cleavage is restricted to the blastodisc. Initial cleavages are meroblastic; the cleavage furrows cut down from the animal pole towards the yolk and stop when they reach the yolk. Ectoderm: The germ layer in an embryo that gives rise to the epidermis and nervous system. In many cases, as in vertebrates, an anterior and posterior invagination of ectoderm gives rise to the stomodeum and proctodeum, respectively. Endoderm: The germ layer in an embryo that gives rise to the epithelium (lining) of the gut and gut derivatives.

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Epiblast: In the chick embryo, following delamination which separates the blastoderm into two layers, the epiblast is the upper layer of cells that contains the presumptive ectoderm, mesoderm and some endoderm. It is separated from the lower hypoblast. Extraembryonic membranes: The membranes that form in association with the embryo proper. In the amniote, four extraembryonic membranes form, the amnion, chorion, yolk sac, and allantois. Fate maps: Maps of the developing embryo that have been constructed to show the location of each germ layer prior to gastrulation and its route of travel during gastrulation. Foregut: In the chick embryo, the region of anterior gut that has been closed into a tube. Gallus domesticus: The species name for the common domestic chicken. Gastrulation: The stage in embryonic development that follows cleavage. During gastrulation, cells rearrange themselves, with the endoderm and mesoderm cells moving inward, and the ectoderm cells spreading around the outside, with some ectoderm invaginating to form such structures as the stomodeum and proctodeum. Germ layers: The ectoderm, mesoderm, and endoderm of a developing embryo. Headfold: In the chick, the undercutting fold of the blastoderm in the area pellucida of the head region which separates the head from the extraembryonic regions, lifting the head up off of the remaining blastoderm. Hensen’s node (or primitive node): In the gastrulating chick embryo, the top of the primitive streak where the notochordal cells are ingressing and migrating forward. Hypoblast: In the chick embryo, following delamination which separates the blastoderm into two layers, the hypoblast is the lower layer of cells that contains the extraembryonic endoderm. Infundibulum: In the adult female chicken, the region of the oviduct where fertilization takes place. It is high up near the beginning of the oviduct. Ingression: During gastrulation, the type of cell movement that involves cells moving inward as individual cells. In the chick embryo, mesoderm and embryonic endoderm move through the primitive streak by ingression. In the sea urchin embryo, the primary mesenchyme moves inward by ingression. Involution: A type of gastrulation movement in which a sheet of cells folds inward and spreads over an inner surface. Isthmus: In the oviduct of the female chicken, it is the region where the chalazae and the inner and outer shell membranes are added to the egg as it descends. Lysozyme: An enzyme that disrupts bacterial cell walls, thereby killing the bacteria. Lysozyme can be found, for example, in the albumen of a chick egg. Macrolecithal egg: An egg with a large quantity of yolk, as in the chick, fish, and insect eggs. Magnum: In the oviduct of the female chicken, it is the region that secretes most of the albumen around the ovum as it travels down the oviduct. Mammillary layer: The inner calcified layer of the chick egg shell, lying next to the outer spongy layer. The crystals of calcium carbonate lie perpendicular to the surface of the egg, giving the shell tremendous strength. Meiosis: The type of cell division that halves the number of chromosomes in cells, taking them from a diploid (2N) to a haploid number (1N) of chromosomes. This occurs only in germ cells. Mesoderm: The germ layer in an embryo that gives rise to the muscular and circulatory systems and most of the skeletal and urogenital systems. Mesolecithal egg: An egg with a moderate amount of yolk, as in the amphibian egg. Microfilaments: The thin filaments, actin and myosin, within the cytoplasm of a cell.

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Microlecithal egg: An egg with a small amount of yolk, as in the sea urchin egg and those of many other invertebrates that have small, feeding larval stages. Microneedle: A needle that is very thin, suitable for microdissection under a dissecting microscope. It can be made inexpensively from inset pins, a wooden dowel, and superglue. Microtubules: Thin tubules found in the cytoplasm of a cell. Microtubules are made up of tubulin subunits. Microtubules function both as a cytoskeletal support and for transport within the cell. Motor-molecules, associated with the microtubules, are able to ferry such things as organelles along the microtubules. Midgut: The region of gut between the foregut and hindgut. Monotreme: A primitive type of mammal that lays eggs. Includes the duck-billed platypus and the spiny ant-eater. Myosin microfilaments: Microfilaments found in the cytoplasm of cells. Myosin microfilaments are the thicker of the two types of microfilaments; actin microfilaments are the thinner. Myosin can move along actin microfilaments in a ratchet-type mechanism that creates a contractile force within the cell. Neural crest cells: Cells found only in vertebrate embryos that start out residing on the crest of the neural tube and subsequently migrate to become a host of structures: pigment cells (except for the pigmented retina), membrane bones in the face and skull, dentine-secreting cells of the teeth, the adrenal medulla, and sympathetic ganglia of the autonomic nervous system are just some of the structures formed. Neural ectoderm: The region of ectoderm that forms the neural tube. Neural plate: The neural ectoderm prior to its invagination to form the neural tube. Neural tube: The tube formed by the neural ectoderm. The precursor to the brain and spinal cord. Neurulation: The morphogenetic movements of the neural ectoderm that form a neural tube out of the neural plate. Notochord: The axial supporting structure in a chordate embryo formed from notochordal mesoderm. Notochordal mesoderm has been called the “primary organizer” because of its inducing functions that initiate an axial organization of the embryo. The notochord is a diagnostic characteristic of the chordates. Ovalbumen: The most abundant glycoprotein found in the albumen of a chick egg. Ovary: The female gonad in which germ cells form ova. Oviduct: The duct in a female that carries eggs from the ovary. Oviposition: The laying of an egg. Ovomucin: A protein that is thought to have antiviral activity. It can be found in the albumen of a chick egg. Ovotransferrin: An enzyme that inhibits bacterial growth by binding iron, making it unavailable to the bacteria. Ovotransferrin can be found in the albumen of a chick egg. Ovulation: The release of an egg (ovum) from the ovary. Physiological polyspermy: An unusual event that occurs in some organisms such as the chick, it is the entry of multiple sperm into an egg at fertilization, but with the result that only one sperm pronucleus fuses with the egg pronucleus. Pipping: At hatching, the process of a baby bird breaking through its shell. Primitive node: Hensen’s node. Primitive streak: In the chick gastrula, the line along which embryonic endoderm and mesoderm cells ingress. Analogous to a blastopore.

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Proamnion: In the chick embryo, the region of area pellucida in front of the embryo’s head that is particularly clear, consisting only of ectoderm and endoderm. The spreading mesoderm has not yet reached this area. Pronucleus: A haploid maternal (from the egg) or paternal (from the sperm) nucleus within a fertilized egg prior to the fusion of these nuclei into a single diploid nucleus. Protostome: Among the bilaterian organisms, the phyla in which the mouth, during embryonic development, forms at or near the blastopore opening. These include phyla such as the arthropods, nematodes, annelids, molluscs, and platyhelminthes. Retinal: A form of vitamin A from which the pigment rhodopsin is made. Rhodopsin: A pigment in the eye that is necessary for vision. Retinal, derived from vitamin A, is used to make rhodopsin. Secondary oocyte: A developing ovum in the secondary gonocyte stage. A secondary oocyte is haploid. Each of its chromosomes consists of two sister chromatids. The first meiotic division divides the primary oocyte asymmetrically, producing one secondary oocyte and one tiny polar body. At the end of this stage, the secondary oocyte enters the second meiotic division. Serial sections: Histological sections of a specimen that are taken sequentially, and kept in order. Usually every section is mounted in correct order for microscopic observations. Shell gland: In the female chick, the region of the oviduct where water and salts are added to the albumen and the shell is deposited. Also called the uterus. Shell membranes: Membranes on the inner aspect of the calcareous portion of an egg shell. In the chick egg, there are two shell membranes. These protect against desiccation and invasion by microorganisms. Shell pores: Pores within an egg shell that allow for gas exchange. Somites: Blocks of mesoderm that become subdivided into a sclerotome, myotome, and dermatome. Sclerotomes form vertebrae and ribs, myotomes form skeletal muscles for the back and appendages, and dermatomes give rise to dermis. Spina bifida: A congenital defect in which the neural plate doesn’t fully close when making the neural tube, and parts of the neural tube remain open. Spongy layer: Also called crystalline layer. The outer calcified layer of the chick egg shell, lying next to the inner mammillary layer. The crystals of calcium carbonate lie perpendicular to the surface of the egg, giving the shell tremendous strength. Staging series: An embryonic timing series that identifies stages of development. A complete staging series is valuable, and once a good one for an organism has been published, this is usually used for many years. The Hamburger-Hamilton staging series for the chick, for example, has been used since its publication in 1951. Unsegmented somite mesoderm: The region of somitic mesoderm that has not yet formed blocks of somites. Uterus: The lower end of the oviduct that is modified for a specific function. In a female placental mammal, it is where the embryo implants. In the female chick, it is where water and salts are added to the albumen and the shell is deposited. In the chick, also called the shell gland. Vagina: The posteriormost region of the duct system leading from the ovaries to the outside. Vegetal pole: The pole that is opposite the animal pole. Yolk is more concentrated at the vegetal pole. Eggs usually orient to gravity such that their vegetal pole is oriented downward. Vertebrata: The class of chordates that include the fish, amphibians, reptiles, birds and mammals. Diagnostic features of the vertebrates are a vertebral column (hence its name), neural crest cells during development, and the presence of bone.

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Vitelline: Of the yolk or having to do with the yolk. Vitelline envelope: An extracellular coat surrounding an ovum. Whole mounts: A histological preparation of a specimen in which the whole specimen is mounted. The specimen is usually mounted on a microscope slide, and can be mounted either in a watermountant such as glycerin jelly or a non-water mountant such as the plastic mounting media used routinely for stained paraffin sections. Yolk: A term commonly used to indicate the ovum of an amniote egg. The yolk is a single cell and contains a large amount of yolk protein in the form of yolk platelets. In a bird egg, the yolk is denser than the albumen and would sink in the albumen if it weren’t suspended by the chalazae. Yolk sac: The extraembryonic membrane that surrounds the yolk. In the amniote, it is vascularized so that it can bring yolk nutrients from the yolk back to the embryo proper. It forms from the splanchnopleure, a layer made up of endoderm and splanchnic lateral plate mesoderm.

Chick: Early Development Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. What are some of the properties of an amniote egg?

2. At the blunt end of the chick egg is an air space. What do you think its functions might be?

3. How would you describe the cleavage pattern in the chick egg? What is this pattern of cleavage called?

4. Why do you think the initial cleavages of the chick egg are meroblastic (incomplete) rather than holoblastic (complete)?

Name: _______________________

Chick: Early - 2

5. What cavity underlies the area pellucida?

6. What is the hypoblast, and how is it formed?

7. What is the primitive streak, and how is it formed?

8. During chick gastrulation, what germ-layer regions remain external and do not move inward?

9. About how many somites can you expect to see in the 24-hour chick embryo?

10. What regions of mesoderm can be identified in the 24-hour chick embryo?

10

chapter

33-Hour Chick Embryo

Whereas the 24-hour chick embryo was roughly equivalent to a 3-week human embryo, a 33-hour chick embryo is roughly equivalent to a 4-week human embryo; what takes hours in a chick embryo takes days in a human embryo. This dramatic difference in developmental rates, in embryos with such similar developmental patterns maintained at roughly the same temperatures, teases the mind with questions. Can you suggest any reasons for the difference? Publish them (I’m serious) or discuss them with your lab colleagues. You will be using whole mounts and serial cross sections to study the 33-hour chick embryo. Be sure you are able to identify all structures printed in boldface, and record in your laboratory notebook your answers to any questions posed. You will also find it useful to keep track of germ layers as you study the cross sections. Do this by color-coding all the tissues in each cross-sectional diagram in the chapter. Remember the cardinal code of embryological colors: blue = ectoderm; red = mesoderm; yellow = endoderm; green = neural crest. The coloring is easy, and the corollaries you can derive from it will serve you well. It teaches you the answers to all those dreaded questions that start out, “What is the germ layer origin of…?” Upon looking at the 33-hour whole mount, you will immediately notice that two major advances have taken place since 24 hours of development. The anterior nerve cord is now a brain subdivided into its major regions, and the rudiments of the circulatory system are present. Understand the general outline of these advances to introduce yourself to this stage.

Central nervous system The neural tube, closed along much of its length, has become specialized anteriorly to form the brain. It has three major subdivisions—the forebrain (prosencephalon), midbrain (mesencephalon), and hindbrain (rhombencephalon)—which are visible as a series of enlargements. The forebrain is further divided into an anterior telencephalon and posterior diencephalon. The diencephalon is becoming very complex: its walls have evaginated to form optic vesicles, which later become the optic cups; its floor has evaginated to form the infundibulum, which later becomes the posterior part of the pituitary gland; its roof, at this point, has no evaginations, but by 48 hours will have evaginated to form the epiphysis, later to become the pineal gland. This is an astonishing array of derivatives. The hindbrain is also subdivided, forming a metencephalon anteriorly and a myelencephalon posteriorly. This organization of brain regions is the same as that of the adult brain, but becomes more difficult to discern as differential growth of the regions, most notably of the telencephalon and metencephalon, obscure the embryonic layout.

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Circulatory system The circulatory system has advanced so that there is now a heart in the form of a relatively straight tube with four chambers. It will not start beating until 48 hours of development. At this point, it looks very much like a fish heart (if you have ever mucked through shark oil to study the shark heart, this should sound familiar). Posteriorly, the heart has a sinus venosus. This leads forward into an undivided atrium, which opens into an undivided ventricle, swung slightly to the right, which in turn leads into the conus arteriosus (or bulbus cordis). From the conus arteriosus sprout a pair of arteries, the ventral aortae. Think about your own four-chambered heart with its two atria and two ventricles. It started out looking like this embryonic chick heart. What happened to mold it into its adult form? The answer is, very much what happens later in the embryonic chick heart. The atrium and ventricle become subdivided into right and left sides. The sinus venosus becomes reduced to the sinoatrial node (or pacemaker), and the conus arteriosus becomes subdivided to form the base of the aorta and pulmonary arteries. The embryonic blood vessels in the chick at 33 hours are still very rudimentary. There are a pair of vitelline veins that will bring nutrient (yolk)-laden blood from the yolk sac back to the embryo. These join the heart at the sinus venosus. A pair of ventral aortae leave the heart and connect to the paired dorsal aortae by way of the first pair of aortic arches. Later additional aortic arches will be added, connecting the ventral and dorsal aortae.

33-Hour Whole Mount Examine your 33-hour whole mount under a dissecting scope, and compare it to the 24-hour whole mount. Look at the two subdivisions of the area opaca: the area vitellina and area vasculosa. (Again, much of the area vitellina has had to be trimmed away for mounting.) Notice that these regions are considerably larger than at 24 hours. Record how much larger by measuring the widths and lengths of the area vasculosa with your micrometer slide. Have the number of blood islands increased as well? At the outer margin of the area vasculosa, there is now a darker band. This is the sinus terminalis, a circular blood vessel marking the terminal channel of the vitelline (yolk) circulation. Look at the area pellucida, and within the area pellucida the proamnion just in front of the head. Recall from the earlier chapter why these areas are translucent, and why the proamnion is particularly clear. Now examine your slide under the low power of your compound microscope. Do not use high power. The slide is too thick and will crack the lens ($$$). Look at the embryonic region to find the structures listed below. (Figure 10.1 will help in your identification.) Folds and tucks: Morphogenesis proceeds Head This is huge in relation to the rest of the embryo. Estimate what percentage of the embryo’s length is taken up by head as measured by the posterior limit of the brain. The head is elongating faster than the tissues around it and is lifting off the blastoderm anteriorly, forming the head fold in the process. Lateral body folds In order to create a tubular-shaped body out of a flat plane, lateral body folds tuck down around the embryonic region. You will see these best in cross section. Primitive streak This is present at the posterior end of the embryo, where gastrulation is still proceeding.

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Proamnion Telencephalon A--B--C---

Diencephalon

Notochord Mesencephalon Foregut

D---

Metencephalon E--Myelencephalon

F--G--H---

Anterior neuropore

Optic vesicle Infundibulum Ventral aorta Head fold Conus arteriosus Ventricle Sinoatrial region

Vitelline vein I--Area vasculosa Nephrogenic mesoderm Open neural plate

Lateral body fold Somites Lateral plate mesoderm Sinus terminalis

J---

Hensen's node K---

Primitive streak Area vitellina

Area pellucida

Figure 10.1 Sch ema ti c di a gra m of a wh ol emount 33-h our ch i ck embryo. Around th e edge from anteri or to posteri or i s a seri es of l etters, ea ch ma rki ng th e l evel of a cross-secti ona l di a gra m sh own i n Fi gure 10.2.

Ectodermal structures Prosencephalon This is the anterior region of the brain, subdivided into a telencephalon and diencephalon. Telencephalon This is not distinctly separate from the diencephalon, and it will suffice to label the brain region anterior to the optic vesicles as telencephalon. Later, the telencephalon becomes bilobed and is associated with the olfactory organs and the sense of smell, as well as being the center of intelligence. We more commonly call it the cerebrum. Anterior neuropore This opening at the anterior end of the brain is still present. Diencephalon This is most easily recognized by its lateral evaginations, the optic vesicles. The rest of the lateral walls become the thalamus, a region of sensory integration. The floor becomes the hypothalamus, a region that sends hormones into the posterior pituitary for storage and secretes releasing hormones that control the anterior pituitary. It should come as no surprise, therefore, to find that the posterior pituitary forms from an evagination of the diencephalon floor, the

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infundibulum. The roof of the diencephalon will form the anterior choroid plexus, a highly vascularized region that secretes cerebral spinal fluid by squeezing out an ultrafiltrate of the blood. An evagination of the diencephalon roof (the epiphysis) forms later, becoming the pineal gland. This is the mysterious “third eye” of some reptiles. (Is this surprising, since the first two eyes are also derived from the diencephalon?) In addition, the pineal gland secretes melatonin and is a seat of control for circadian and seasonal rhythms. In the early 1600s, the famous French philosopher Descartes considered the pineal gland to be the seat of the soul. Optic vesicles These are evaginations of the lateral walls of the diencephalon. They grow close to the head epidermis and will induce it to form the lens of the eye. The optic vesicles invaginate to form optic cups, the inner side of which differentiate into the neural retina and the outer layer into the pigmented retina. The optic nerve grows back from the neural retina along the connection of the optic cup to the diencephalon. The optic nerves from opposite sides then cross underneath the diencephalon and enter the mesencephalon. (Think about this: most of your eye is an outpocketing of your embryonic brain.) Infundibulum The floor of the diencephalon evaginates, forming the infundibulum. It later differentiates into the posterior pituitary (or neurohypophysis), which stores two hormones from the hypothalamus: antidiuretic hormone (which causes the kidney to resorb water and produce more concentrated urine), and oxytocin (which causes uterine contractions and milk release in mammals). Mesencephalon This is the middle, oval-shaped region of the brain that will serve primarily for processing data from the eyes and ears. The dorsal region will become the optic lobes, visual centers associated with the optic nerves. Rhombencephalon This region becomes associated with the system of hearing and balance and is divided into an anterior metencephalon and posterior myelencephalon. Metencephalon This will form the cerebellum dorsally and the pons ventrally. The cerebellum primarily coordinates stimuli concerning body position and movement; the pons is a bundle of connective pathways shunting information between the cerebrum and cerebellum. Myelencephalon Also called the medulla oblongata, this region is characterized by a series of enlargements called neuromeres, each of which will become associated with a specific set of motor and sensory nerves. The myelencephalon becomes the great linking highway between the brain and spinal cord. Spinal cord This is the posterior continuation of the neural tube. Notice how far back on your embryo the closed neural tube extends. Otic placodes These are epidermal thickenings on the side of the head in the region of the myelencephalon. “Otic” stands for ear, and these placodes, seen best in cross section, later invaginate to form the inner ear. This is the organ of both hearing (the cochlea) and balance (the semicircular canals). A category unto themselves: Color them green Neural crest cells These are some of the most phenomenal cells in the embryo. The neural crest cells are an evolutionary invention of the vertebrates. First residing on the crest of the neural tube, they later migrate to become a host of structures: pigment cells (except for the pigmented retina), membrane bones in the face and skull, dentine-secreting cells of the teeth (not in birds, of course—why is it, do you think, that birds don’t have teeth?), the adrenal medulla, and sympathetic ganglia of the autonomic nervous system are just some of the structures formed. The complete list is much longer. (I’ve always suspected that if all parts of the vertebrate body disappeared except those derived from neural crest, you’d still have the ghostly image of a body remaining.)

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Endodermal structures Remember that the endoderm forms only epithelial structures. During the formation of the gut, the endoderm contributes only the epithelial tube that forms the inner epithelial lining of the organs of the gut. It is the mesoderm surrounding this endodermal tube that forms the muscle and connective tissue layers of these organs. Foregut This is the region of anterior gut that has been closed into a tube. This happens when the head fold and lateral body folds undercut the embryo, closing off the gut and bringing the cardiac mesoderm together beneath (ventral to) the gut. The anterior region of the foregut is called the pharynx and forms the esophagus as well as a number of outpocketings, or evaginations. Two evaginations will form ventrally. The anterior one will become the thyroid gland and the posterior one the lungs. A series of lateral evaginations are the pharyngeal pouches that are later transformed into the eustachian tube and cavity of the middle ear, and the epithelium of the tonsils, thymus, parathyroid, and ultimobranchial bodies. The posterior region of the foregut forms the stomach. Anterior intestinal portal This is the opening of the foregut into the open midgut. The opening continues to shift posteriorly as closure of the gut continues. It can be seen more readily by turning your slide over and viewing the embryo from the ventral side. Mesodermal structures Heart This lies ventral to the foregut. At this point in development, like the brain, it is huge compared to the rest of the body. (Think where your heart is in relation to your foregut [esophagus]. Is it ventral or dorsal?) Notice that the phrase “having one’s heart in one’s throat” truly applies to a vertebrate embryo at this stage of development. Obviously, the heart must shift posteriorly to the thoracic region. This will occur primarily through an elongation of the head and neck regions. Look for the large vitelline veins coming in from the extraembryonic regions and entering the posterior chamber of the heart, the sinus venosus. At this point in development, the sinus venousus cannot be readily distinguished from the next chamber, the atrium. Therefore, you can refer to them both as the sinoatrial region. The ventricle is the next chamber and is easily distinguishable since it bends to the right. Where the heart narrows again anteriorly marks the conus arteriosus. If you focus very carefully while viewing the embryo from the ventral side (turn your slide upside down), you may be able to see the paired ventral aortae leading away from the conus arteriosus. Notochord This is visible as a dark line in the midline ventral to the neural tube. Determine how far anteriorly and posteriorly it extends. Pause to reflect on its importance. Remember, chordates wouldn’t exist without the notochord. The notochordal mesoderm has been called the “primary organizer” because of its many inducing functions that initiate an axial organization of the organism. It also provides axial support for the embryo until the vertebrae engulf it, taking over this function. In the adult, notochordal tissue between the vertebrae expands to form the gelatinous center (nucleus pulposus) of the intervertebral disc. When a person suffers from a “slipped disc” or herniated disc, stress on the back has caused the nucleus pulposus to bulge outward. The bulging disc puts pressure on the spinal cord and causes back pain. Often, rest and proper back exercises can remedy this condition without the need of surgery. There is also a core of notochord left in the centrum of each vertebra. Sometimes this remnant of notochord gives rise to tumors called chordomas. These tumors cause extreme back pain by narrowing the spinal canal and pinching the spinal cord and must be removed surgically. Somites These blocks of mesoderm are found on either side of the neural tube. Each somite becomes subdivided into a sclerotome that forms the vertebrae and ribs, a myotome that forms skeletal

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muscles for the back and limbs, and a dermatome that gives rise to dermis. Count the number of pairs of somites your embryo has and use this to stage your embryo using the Hamburger-Hamilton staging series (Table 10.1). Record the stage. Nephrogenic (intermediate) mesoderm This is a streak of mesoderm lying lateral to the somites. It will form the kidneys, the urogenital ducts, and the gonads. Lateral plate mesoderm Lateral to the nephrogenic mesoderm is the lateral plate mesoderm, which delaminates to form two layers: the splanchnic and somatic layers. The splitting creates a cavity, the body coelom. Somatic lateral plate mesoderm becomes associated with the epidermal ectoderm, and together they form the somatopleure. This later will form the body wall and, in the extraembryonic regions, the amnion and chorion (the extraembryonic membranes that surround and protect the embryo). The splanchnic lateral plate mesoderm associates with the endoderm, which together form the splanchnopleure. This forms the gut wall and, in the extraembryonic regions, the vascularized yolk sac and allantois. Because of its extensive vascularization, the yolk sac can transport yolk nutrients back to the embryo, and the allantois can receive nitrogenous waste from the embryo for storage as uric acid. Table 10.1 Staging series of embryonic chick development. (After Hamburger and Hamilton, 1951.) Stage and length of incubation (at 38°C) Description of embryo Stage 1: Prestreak

After laying, cleavage is complete and gastrulation has begun, but the primitive streak has not yet formed.

Stage 2: Initial streak (6–7 hours)

First formation of the primitive streak.

Stage 3: Intermediate streak (12–13 hours)

A broad streak has formed that extends from the posterior margin to the center of the area pellucida.

Stage 4: Definitive streak (18–19 hours)

The primitive streak extends about three–quarters the length of the area pellucida, which is now pear-shaped. Hensen’s node is present.

Stage 5: Head process (19–22 hours)

The notochord is visible, but the head fold has not yet formed.

Stage 6: Head fold ( 23–25 hours)

Anteriorly, the head fold is distinct. No somites have formed yet.

Stage 7: 1 Somite ( 23–26 hours)

One pair of somites can be seen, and the neural folds are visible anteriorly.

Stage 8: 4 Somites

( 26–29 hours)

Four pairs of somites are present. Neural folds meet at the level of the midbrain, and blood islands are seen in the area vasculosa.

Stage 9: 7 Somites

( 29–33 hours)

Seven pairs of somites. The optic vesicles are forming, and the heart is beginning to form.

Stage 10: 10 Somites (33–38 hours)

10 pairs of somites. Three major subdivisions of the brain are visible. Heart is bent slightly to the right.

Stage 11: 13 Somites (40–45 hours)

13 pairs of somites. Five neuromeres of the myelencephalon can be seen. Anterior neuropore is closing.

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Table 10.1 (Continued) Stage and length of incubation (at 38°C) Description of embryo ( 45–49 hours)

16 pairs of somites. Head is turning to the left. Optic placode is invaginating.

Stage 13: 19 Somites (48–52 hours)

19 pairs of somites. Head is turned almost fully to the side.

Stage 14: 22 Somites

22 pairs of somites. Body is rotated to the side as far back as 7–9 somites First two pharyngeal pouches and grooves are distinct. Optic vesicles invaginating.

Stage 12: 16 Somites

( 50–53 hours)

Stage 15: Optic cup (50–55 hours)

24–27 pairs of somites. The third pharyngeal pouch and groove have formed. The optic cup is formed.

Stage 16: Pre-limb bud (51–56 hours)

26–28 pairs of somites. Tail bud is beginning to form.

Stage 17: Early limb bud (52–64 hours)

29–32 pairs of somites. Both wing and leg buds are distinct swellings. Tail is bent ventrally. Allantois not yet present.

Stage 18: Early allantois (3 days)

30–36 pairs of somites. Limb buds have enlarged. Tail bud has turned to the right. Allantois is a thick-walled pocket.

Stage 19: Somites in tail (3–3 1/2 days)

37–40 pairs of somites, which extend into the tail. Leg bud is slightly larger than wing bud. Eyes are unpigmented.

Stage 20: Vesicular allantois (3–3 1/2 days)

40–43 pairs of somites. Leg bud slightly asymmetrical. Allantois vesicular, about the size of the midbrain.

Stage 21: Early eye pigmentation ( 3 1/2 days)

43–44 pairs of somites. Both wing and leg buds slightly asymmetrical. Allantois enlarged. Eye pigmentation faint.

Stage 22: Distinct eye pigmentation ( 3 1/2 –4 days)

Somites extend to tip of tail. Eye pigmentation is distinct.

Stage 23: Limb buds as wide as long (4 days)

Both wing and leg buds are as wide as they are long.

Stage 24: Limb buds longer than wide ( 4 1/2 days)

Both wing and leg buds are distinctly longer than they are wide. Toe plate is distinct, but toes are not yet demarcated.

Stage 25: Early elbow and knee (4 1/2 –5 days)

The elbow and knee joints are distinct. The digital plate in the wing is distinct but with no demarcation of digits. Third toe is demarcated by a faint groove.

Stage 26: Three toes (5 days)

Limbs are considerably longer. The demarcation of three toes is distinct. There is a faint groove between the second and third digit of the wing.

S tag e 2 7 : Earl y beak (5–5 1/2 days)

The three digits of the wing are demarcated by grooves. The beak is just recognizable.

Stage 28: Beak distinct (5 1/2 –6 days)

The second digit of the wing and the third toe are longer than the rest. The three wing digits and four toes are distinct. (Chickens only have three “fingers” and four toes.) The beak is a distinct outgrowth.

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33-Hour Serial Sections The ability to read serial sections and to construct a three-dimensional mental image of the whole is a skill worth acquiring. Be patient with yourself. Look at your slides; there will probably be two of them. The embryo has been sliced into (usually) 10-µm-thick sections, and the sections have been placed in order, anterior to posterior. Not a single section is lost. There will be a number of rows of sections on each slide. By convention, these were laid down starting at the upper left-hand corner of the first slide. In scanning the slide, you will read it just as you would a book, from left to right. Now put the slide on your compound microscope and move the controls of the stage so that the slide moves from left to right along the top row. Notice that when you look through the microscope, your movements look reversed, and the slide appears to be moving from right to left. To read the next row, you must move the slide down and all the way back to the left. All this magnified motion whizzing past your eyes can make you feel sick in short order. Some hints from a master of queasiness: never look through the microscope as you are changing rows, and always move slowly from one section to the next, relaxing your eyes as you do to avoid trying to focus on the blurred image as it passes. Look at the diagrams of cross sections (Figures 10.2A–K). The letter of each cross section A–K matches a letter on the whole-mount diagram (Figure 10.1), showing the level from which each cross section was taken. Your own sections will probably look slightly different from the diagrams, since the angle of cutting can vary. As you read through your serial sections, it is extremely important that you constantly refer back to the whole mount to know where you are in relation to the rest of the embryo. Remember to color-code the cross-sectional diagrams. The letters below correspond to the letters of the cross-sectional diagrams in Figure 10.2. Section A Start reading your sections from the anterior end. The first few sections through the prosencephalon will show a narrow ventral cleft through the neural tube. This is the anterior neuropore. Identify the two-layered proamnion just below the head. What are the two germ layers represented here? There is a space between the head and the proamnion called the subcephalic (“below the head”) pocket. Remember why this space exists: the head fold has undercut the embryo, lifting the head up from the level of the blastoderm. Lateral to the proamnion, the somatopleure and splanchnopleure can be seen, with the large coelomic space between them. Notice that there are capillaries in the mesoderm of the splanchopleure. This layer becomes highly vascularized. The capillaries you see are part of the vitelline circulation. Section B Sections through the diencephalon will show the lateral optic vesicles and ventral infundibulum. Look carefully at the epidermal ectoderm that is in contact with the optic vesicles to see if there are any areas of thickening. If there are, you have found the lens placodes. These later will invaginate to form the lens vesicles. The loose tissue surrounding the brain is head mesenchyme, derived from mesoderm and neural crest cells, which will later form such structures as the skull, head musculature, blood vessels, and connective tissue. Section C Further posteriorly you will see the rounded cross section of the mesencephalon. At this point, the foregut, looking like a smile, should be obvious below the notochord. The foregut in this region comes in contact with the epidermal ectoderm to form the oral membrane. Fortunately, this membrane is removed by programmed cell death; the opening produced is the mouth opening. When cell death fails to occur, an anomaly results called astomia (“no mouth”). Above the foregut you should see the paired dorsal aortae, and below the foregut the paired ventral aortae. A pair of first aortic arches connects the ventral aortae with the dorsal aortae. Which way will the blood flow through these vessels? Show the direction by drawing arrows on the diagram.

33-Hour Chick Embryo (A)

Telencephalon

Neural ectoderm Epidermal ectoderm

Somatic mesoderm Somatopleure

Proamnion

Subcephalic pocket Coelom Anterior neuropore Endoderm Splanchnic mesoderm Splanchnopleure

Diencephalon (B)

Head mesenchyme

Dorsal aorta

Optic vesicle Future lens ectoderm

Somatopleure Infundibulum

Proamnion Splanchnopleure

Foregut

Mesencephalon

(C)

Subcephalic pocket

Coelom

Epidermal ectoderm

Dorsal aorta Notochord First aortic arch Ventral aorta

Somatopleure

Proamnion Splanchnopleure

Foregut Oral membrane

Subcephalic pocket Coelom

Figure 10.2 Cross-sectional diagrams of the 33-hour chick embryo. Diagrams are arranged from anterior to posterior. The letter beside each diagram corresponds to the level shown on Figure 10.1 from which the section is taken.

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33-Hour Chick Embryo (D)

Metencephalon

Head mesenchyme Epidermal ectoderm

Dorsal aorta Lateral body fold

Coelom

Foregut Endoderm

(E)

Proamnion Subcephalic pocket

Myelencephalon

Conus arteriosus

Head mesenchyme Epidermal ectoderm

Dorsal aorta

Coelom Endoderm Foregut

(F)

Vitelline vein Ventricle Endocardium Myocardium

Myelencephalon Dorsal aorta

Epidermal ectoderm Otic placode

Coelom Endoderm Foregut

Vitelline vein Atrium Endocardium Myocardium

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33-Hour Chick Embryo

(G)

Myelencephalon

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Epidermal ectoderm Otic placode

Dorsal aorta

Coelom

Sinus venosus Foregut

Vitelline vein

Extraembryonic endoderm

(H)

Spinal cord

Epidermal ectoderm

Somatopleure

Splanchnopleure

Neural crest

Endoderm Coelom

Dorsal aorta

Foregut

Vitelline vein

Anterior intestinal portal

Section D Trace the ventral aortae posteriorly and see that they fuse to form the conus arteriosus. The head is now attached to the blastoderm, but the indentations of the lateral body folds can still be seen. These eventually will undercut the embryo and lift it off the blastoderm. Notice the narrowed rhombencephalon. The anterior portion of this is the metencephalon. Section E If you are very observant as you move posteriorly, you will be able to make out the intermittent expansions of the neuromeres as you go through the myelencephalon. At the level of the myelencephalon, you also should see that the heart appears larger and swings out to the right. This region of the heart is the ventricle.

33-Hour Chick Embryo (I)

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Neural crest Spinal cord

Somite

Nephrogenic mesoderm

Epidermal ectoderm

Coelom Somatic mesoderm Dorsal aorta

(J)

Unsegmented somite mesoderm

Splanchnic mesoderm Endoderm Notochord

Open neural plate Epidermal ectoderm

Coelom

Notochord

Lateral plate mesoderm

Endoderm

Primitive groove

(K)

Ectoderm

Endoderm

Primitive streak

Mesoderm

Section F As you move posteriorly through the heart, it swings back to the midline. You are now in the region of the atrium. Look carefully at the dorsal epidermal ectoderm in this region. On either side of the myelencephalon, you should see epidermal thickenings. These are the otic placodes, which will later invaginate to form the inner ear. Section G Further posteriorly, where the heart suddenly flattens out, you are in the region of the sinus venosus. The vitelline veins, feeding into the sinus venosus, can be seen in cross section throughout the mesoderm of the splanchnopleure in this region.

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Section H Just posterior to the region of the sinus venosus, the foregut opens at the anterior intestinal portal to become the open midgut. Look at the neural tube and for the tightly packed cells dorsally, on either side of the neural tube. These are the neural crest cells. Prior to their migration, they can be seen all along the dorsal side of the neural tube. Section I Once you are in the region of the open midgut, you are past the brain and in the area of the spinal cord. Here you should see clearly the subdivisions of the mesoderm: the somites; more laterally, the nephrogenic mesoderm; and finally, the lateral plate mesoderm, which is split into somatic and splanchnic layers. Go back and forth through several somites to see that somites are indeed blocks of tissue and not a continuous rod. Count the number of somites as you travel posteriorly through them and use this to stage your embryo. How close is it to the age of your wholemount embryo? Record the stage in your laboratory notebook. Section J As you proceed posteriorly, the neural tube will become an open neural plate, and the somite mesoderm will be unsegmented. Notice that the lateral plate mesoderm is just beginning its delamination to form the coelom. Section K Still further posteriorly, you will reach the region of the primitive streak, where ingression is still occurring. Because differentiation in the chick embryo proceeds in an anterior-to-posterior wave, you will find that traveling posteriorly through the sections gives the impression of traveling backward in time. If you want to get the impression of traveling forward through time, study your sections from posterior to anterior.

Experiment with Evolution Make a diagram of a cross section through the chick embryo in the region of the somites. Now cut it out, and keeping it flat on a table, bend the two ends around until they meet ventrally. Compare this to a diagram of a cross section through an amphibian neurula. What does this tell you about patterns of gastrulation and the impediments of yolk?

Create a Puppet Visualizing the tucks and folds that create the chick embryo out of the three-tiered gastrula sandwich is difficult without models. You can greatly enhance your understanding by making a puppet out of three layers of cloth, an outer blue, middle red, and lower yellow (of course). Put your arm under the layers of cloth. Lift your hand slightly, and tuck the cloth in around your hand. You have just created a head fold. Look at the underside of your puppet, and see how the folding has created an anterior tube lined in yellow (the foregut). Now tuck the cloth in along the sides of your arm to create lateral body folds. Wherever you tuck it in completely so that right side meets left side, you have closed off more regions of gut and lifted the embryo up above the level of the rest of the cloth (the extraembryonic regions).

Accompanying Materials See Vade Mecum: “Chick Mid.” This chapter of the CD includes whole mounts, color-coded to show germ layers, and labeled, with definitions of anatomical terms. Also a complete set of serial cross sections through a 33-hour chick embryo includes labels and color-coding to show germ layers. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. Contains excellent diagrams and descriptions of neurulation, and somite, heart and gut formation, as well as

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limb and gonad development. There is a diagram of an amphibian neurula that can be used for the “Experiment With Evolution” section above. Fink, R. (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Sequence 11. This shows head fold, brain, somite, and heart formation, along with closure of the foregut and regression of Hensen’s node.

Selected Bibliography Hall, B. K. and S. Hörstadius. 1988. The Neural Crest. Oxford University Press, London. This is a detailed, thought-provoking volume on the evolutionary origins, migrations, and differentiation of neural crest cells. It includes a facsimile reprint of the classic book by S. Hörstadius from 1950. Hamburger, V., and H. L. Hamilton, 1951. A series of normal stages in the development of the chick embryo. J. Morphol. 88: 49–92. This is the original publication of the Hamburger-Hamilton chick staging series. Hamilton, H. L. 1952. Lillie’s Development of the Chick. An Introduction to Embryology. Henry Holt and Co., New York. This revised version of F. R. Lillie’s original, much longer, book is a classic. It contains a full reprint of the Hamburger-Hamilton chick staging series. Kil, S. H., C. E. Krull, G. Cann, D. Clegg and M. Bronner-Fraser. 1998. The α4subunit of integrin is important for neural crest cell migration. Dev. Biol. 202: 29–42. This is an excellent example of how modern techniques in immunohistochemistry are used to track neural crest cells and determine the components important to their migration. LeLièvre, C. and N. M. LeDouarin. 1975. Mesenchymal derivates of the neural crest: Analysis of chimaeric quail and chick embryos. J. Embryol. Exp. Morph. 34: 125–154. These two authors have contributed impressively to our knowledge of neural crest migration and differentiation. This study shows how cell migration can be mapped using embryonic cells from the quail grafted into a chick embryo. The quail cells can be distinguished since their nuclei stain more darkly than those of the chick. Lehman, H. E. 1987. Chordate Development, 3rd Ed. Hunter Textbooks, Winston-Salem, NC. This tome on vertebrate development is filled with interesting detail. In addition to describing developmental anatomy, it discusses development from an evolutionary perspective. Mathews, W. W. and G. C. Schoenwolf. 1998. Atlas of Descriptive Embryology, 5th Ed. Prentice-Hall, Inc., Upper Saddle River, NJ. This traditional manual is one of the best for its photographs of embryos and includes a glossary of terms. Noden, D. M. 1980. The migration and cytodifferentiation of cranial neural crest cells. In Current Research Trends in Prenatal Craniofacial Development, R. M. Pratt and R. L. Christiansen (eds.). Elsevier/North-Holland, New York, pp. 3–26. This author has done painstaking, precise fatemapping, revolutionizing our understanding of the skull. That fate mapping is still so important to embryology is evidenced by Noden’s work and is clearly encapsulated in this review paper. Romanoff, A. L. 1960. The Avian Embryo. Macmillan, New York. This is the biggest book you’ll ever want to read on the bird embryo. The diagrams and charts are extensive. Simply keep in mind that some ideas concerning germ layer derivatives (those of the neural crest, for example) have changed in recent years.

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Suppliers Prepared microscope slides of chick embryos can be obtained from suppliers such as: Turtox/Cambosco, Macmillan Science Co., Inc. 8200 South Hoyne Ave. Chicago, IL 60620 1-800-621-8980

Connecticut Valley Biological Supply Co., Inc. P.O. Box 326 82 Valley Road Southampton, MA 01073 1-800-628-7748

Ward’s Natural Science Establishment, Inc. 5100 West Henrietta Road P.O. Box 92912 Rochester, NY 14692-2660 1-800-962-2660 www.wardsci.com Models of chick development are available from Turtox, listed above.

Glossary Allantois: The extraembryonic membrane found in amniotes that forms off the hindgut of the embryo. It stores nitrogenous waste and in the chick will eventually fuse with the chorion to form the chorioallantoic membrane. It is vascularized and is formed from the splanchnopleure, a layer made up of endoderm and splanchnic lateral plate mesoderm. Amnion: The extraembryonic membrane found in amniotes that surrounds the embryo. It contains amniotic fluid, which protects the embryo by providing an isotonic fluid and buffering the embryo against being jarred. It forms from the somatopleure, a layer made up of epidermal ectoderm and somatic lateral plate mesoderm. Anterior choroid plexus: A thin, highly vascularized region of the roof of the diencephalon, which secretes cerebral spinal fluid by squeezing out an ultrafiltrate of the blood. Anterior intestinal portal: In the chick embryo, the opening between the closed foregut and the open midgut. The opening continues to shift posteriorly as closure of the gut continues. Anterior neuropore: The opening at the anterior end of the brain. It will close later. Aorta: In a vertebrate, the major blood vessel leaving the heart (left ventricle) and going to the body. Aortic arches: The paired arteries that connect the ventral aorta to the dorsal aorta. They are the blood vessels of the visceral (pharyngeal) arches, one pair per visceral arch. Area opaca: In the chick embryo, the outer extraembryonic region that is still connected to the underlying yolk.

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Area pellucida: In a chick embryo, the inner extraembryonic region that is separated from the yolk by a subgerminal cavity. This area looks lighter and more translucent than the surrounding area because of the space below it. Area vasculosa: The inner region of the area opaca where blood islands are forming. Area vitellina: The outer region of the area opaca where cells contain many yolk granules. (Vitellina refers to yolk.) Atrium: The region of the heart that receives blood from the sinus venosus and delivers it to the ventricle. Blood islands: In the chick embryo, the beginnings of blood vessel formation out in the area vasculosa. These are masses of blood-forming cells that will later anastomose to form a capillary network that will bring yolk nutrients back to the embryo proper. They form from splanchnic lateral plate mesoderm. Bulbus cordis: Another name for the conus arteriosus. Cerebrum: In a vertebrate, the anterior-most region of the brain, also called the telencephalon. It is associated with olfactory organs and the sense of smell, as well as being the center of intelligence. Chorioallantoic membrane: The fused chorion and allantoic membranes. In the chick, this serves a major function in respiration. It becomes very large, and presses against the inner aspect of the shell, allowing for efficient gas exchange. Chorioallantoic membrane graft: A graft placed on the chorioallantoic membrane of an embryonic chick. The highly vascularized membrane makes an excellent and inexpensive incubator for growing embryonic tissues. Chorion: The extraembryonic membrane found in amniotes that lies outside of the amnion. It protects the embryo, and in the chick will eventually fuse with the allantoic membrane. It forms from the somatopleure, a layer made up of epidermal ectoderm and somatic lateral plate mesoderm. Coelom: The body cavity in which the gut is suspended. It is lined by mesoderm. Compound microscope: A microscope that uses multiple lenses to increase total magnification. Conus arteriosus (also called conotruncus or bulbus cordis): The region of the heart that receives blood from the ventricle and delivers blood to the ventral aortae. Dermatome: In a vertebrate embryo, a subdivision of the somite. It gives rise to dermis. Diencephalon: The region of the forebrain posterior to the telencephalon. Its lateral evaginations form the optic vesicles, a ventral evagination forms the infundibulum, and a dorsal evagination forms the epiphysis. Dorsal aortae: In the vertebrate embryo, the blood vessels taking blood to the body. They are connected to the ventral aortae by way of paired aortic arches. Blood coming from the heart goes through the ventral aortae, the paired aortic arches, and into the dorsal aortae. Ectoderm: In an embryo, the germ layer that gives rise to the epidermis and nervous system. Embryonal area: The region that will form the embryo proper as opposed to extraembryonic regions. Endocardium: The endothelial lining of the embryonic heart tube. Endoderm: The germ layer in an embryo that gives rise to the epithelium (lining) of the gut and gut derivatives. Endothelium: The specific epithelium of the blood vascular system. It is a simple squamous epithelium, designed for rapid gas exchange. Epidermal ectoderm: The region of ectoderm that forms the epidermis (the epithelial layer of the skin). Epidermis: The epithelium of the skin. It is typically a stratified squamous epithelium, with outer keratinized layers of cells that protect against dehydration and physical abrasion.

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An evagination of the roof of the diencephalon that becomes the pineal gland.

outside of the body. The epidermis is an example of a specific type of epithelium. Evagination: An outpocketing of a sheet of cells. The opposite of invagination. Extraembryonic membranes: The membranes that form in association with the embryo proper. In the amniote, four extraembryonic membranes form, the amnion, chorion, yolk sac, and allantois. Extraembryonic regions: Embryonic regions that do not contribute to forming the body of the embryo proper. These regions form the extraembryonic membranes. First aortic arch: The aortic arch associated with the first visceral arch. Forebrain: The anterior-most region of the brain. In the embryonic vertebrate, this is the prosencephalon, which gives rise to the telencephalon and diencephalon. Foregut: In the chick embryo, the region of anterior gut that has been closed into a tube. Germ layers: The ectoderm, mesoderm, and endoderm of a developing embryo. Head mesenchyme: In the vertebrate embryo, the loose tissue surrounding the brain that is derived from mesoderm and neural crest cells, and which later will form such structures as the skull, head musculature, blood vessels, and connective tissue. Hensen’s node (or primitive node): In the gastrulating chick embryo, the top of the primitive streak where the notochordal cells are ingressing and migrating forward. Hindbrain: The posterior-most region of the brain. In the embryonic vertebrate, this is the rhombencephalon, which gives rise to the metencephalon and myelencephalon. Infundibulum: Ventral evagination of the diencephalon. It becomes the posterior pituitary (or neurohypophysis). Ingression: During gastrulation, the type of cell movement that involves cells moving inward as individual cells. In the chick embryo, mesoderm and embryonic endoderm move through the primitive streak by ingression. Intermediate mesoderm: Nephrogenic mesoderm. Invagination: The folding inward of a sheet of cells. Lateral body folds: In the chick embryo, undercutting folds in the area pellucida along the lateral sides of the embryo proper. They lift the embryo up, separating it from the extraembryonic regions. Lateral plate mesoderm: Mesoderm lying lateral to the intermediate mesoderm. It delaminates into two layers: the splanchnic and somatic layers. The splitting creates a cavity, the body coelom. Lens placode: In the vertebrate embryo, a thickened disc of epidermal ectoderm cells that forms next to the optic vesicle. The lens placode later invaginates to form the lens vesicle, and ultimately forms the lens of the eye. Medulla oblongata: In a vertebrate, the posterior-most region of the brain, also called the myelencephalon. In embryonic development, it is characterized by a series of neuromeres, each of which will become associated with a specific set of motor and sensory nerves. Mesencephalon: The midbrain, the middle region of the brain that will serve primarily for processing data from the eyes and ears. Mesoderm: The germ layer in an embryo that gives rise to the muscular and circulatory systems and most of the skeletal and urogenital systems. Metencephalon: Forms the cerebellum dorsally and the pons ventrally. The cerebellum primarily coordinates stimuli concerning body position and movement; the pons is a bundle of connective pathways shunting information between the cerebrum and cerebellum. Midbrain: The middle region of the brain. In the vertebrate embryo, this is the mesencephalon which serves primarily for processing data from the eyes and ears.

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Myelencephalon: Also called the medulla oblongata, it is associated with a series of enlargements called neuromeres, each of which becomes associated with a specific set of motor or sensory nerves. Myocardium: The muscular wall of the heart. Myotome: In a vertebrate embryo, a subdivision of the somite. It will form skeletal muscles of the back and limbs. Nephrogenic mesoderm: Also called intermediate mesoderm. It will form the kidneys, urogenital ducts and gonads. The germ cells of the gonads are not of nephrogenic mesoderm origin, but migrate into the gonad once the gonad has formed. Neural crest cells: Cells found only in vertebrate embryos that start out residing on the crest of the neural tube and subsequently migrate to become a host of structures. Pigment cells (except for the pigmented retina), membrane bones in the face and skull, dentine-secreting cells of the teeth, the adrenal medulla, and sympathetic ganglia of the autonomic nervous system are just some of the structures formed. Neural fold: The neural ectoderm that is folding upward to form a neural tube. Neural plate: The neural ectoderm prior to its invagination to form the neural tube. Neural tube: The tube formed by the neural ectoderm. The precursor to the brain and spinal cord. Neuromeres: In a vertebrate embryo, the series of enlargements found along the myelencephalon. Each of these will become associated with a specific set of motor and sensory nerves. Neurula: The stage in embryonic development when neurulation is occurring. Notochord: The axial supporting structure in a chordate embryo formed from notochordal mesoderm. Notochordal mesoderm has been called the “primary organizer” because of its inducing functions that initiate an axial organization of the embryo. The notochord is a diagnostic characteristic of the chordates. Optic cup: In a vertebrate embryo, the precursor to the neural and pigmented retinas of the eye. During eye development, optic vesicles that form as outpocketings of the diencephalon invaginate to form the optic cups. Optic vesicles: Lateral evaginations of the diencephalon. They later form the optic cups that will become the neural and pigmented retinas of the eyes. Oral membrane: In the vertebrate embryo, the membrane that forms where the foregut comes in contact with the epidermal ectoderm of the invaginated stomodeum. The fate of this membrane is to undergo programmed cell death so that the oral cavity is continuous with the rest of the gut tube. Otic placode: Epidermal thickening that will invaginate to form the inner ear. Pharyngeal pouches: Typical of chordates. These form embryonically. In organisms with gills, they will form the gill slits. In higher vertebrates, they are transformed into structures such as the eustachian tube and cavity of the middle ear, the epithelium of the tonsils, thymus, parathyroid, and ultimobranchial bodies. Pharynx: In the vertebrate, the anterior region of the foregut. During embryogenesis, it consists of the esophagus and a number of outpocketings—paired lateral outpocketings called the pharyngeal pouches, and two unpaired ventral ones, the anterior one forming the thyroid gland and the posterior one forming the lungs. Pineal gland: A gland associated with the diencephalon that secretes melatonin, a hormone associated with daily rhythms. It also forms the “third eye” found in some reptiles. Pituitary gland: A composite gland found in vertebrates, made up of a neurohypophysis and an adenohypophysis. The neurohypophysis (secreting oxytocin and the antidiuretic hormone,

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vasopressin) is derived from the infundibulum, an outpocketing of the floor of the diencephalon. The adenohypophysis [secreting hormones such as thyroid-stimulating hormone (TSH), adrenocorticotropin hormone (ACTH), luteinizing hormone (LH), follicle-stimulating hormone (FSH), prolactin, and growth hormone] is derived from an evagination of the stomodeum called Rathke’s pocket. Primitive groove: In the chick embryo, the depression in the primitive streak where cells are ingressing. Primitive streak: In the chick gastrula, the line along which embryonic endoderm and mesoderm cells ingress. Analogous to a blastopore. Proamnion: In the chick embryo, the region of area pellucida in front of the embryo’s head that is particularly clear, consisting only of ectoderm and endoderm. The spreading mesoderm has not yet reached this area. Prosencephalon: The forebrain. In the vertebrates, this consists of an anterior telencephalon and a posterior diencephalon. Pulmonary arteries: In a vertebrate with lungs, the major blood vessels leaving the heart (right ventricle) and taking blood to the lungs. Rhombencephalon: The hindbrain. In the vertebrates, this consists of a metencephalon anteriorly and a myelencephalon posteriorly. Sclerotome: In a vertebrate embryo, a subdivision of the somite. It will form the vertebrae and ribs. Serial sections: Histological sections of a specimen that are taken sequentially, and kept in order. Usually every section is mounted in correct order for microscopic observations. Sinoatrial node: The pacemaker of a vertebrate heart. It represents the reduced sinus venosus. Sinoatrial region: The region of the heart that includes the sinus venosus and atrium. It receives blood from the vitelline veins and delivers it to the ventricle. Sinus terminalis: In the chick embryo, a circular blood vessel marking the terminal channel of the vitelline (yolk) circulation. It is found marking the boundary between the area vasculosa and area vitellina. Sinus venosus: The region of the heart that receives blood from the vitelline veins and delivers it to the atrium. Somatic lateral plate mesoderm: The layer of lateral plate mesoderm lining the coelom and that is associated with the epidermal ectoderm to form the somatopleure. This layer of mesoderm is not able to form blood vessels. Somatic: Of the body. Somatopleure: In an amniote embryo, the layer formed by the somatic lateral plate mesoderm and the epidermal ectoderm. It forms the body wall as well as the amnion and chorion. Somites: Blocks of mesoderm that become subdivided into a sclerotome, myotome, and dermatome. Sclerotomes form vertebrae and ribs, myotomes form skeletal muscles for the back and limbs, and dermatomes give rise to dermis. Somitic: Of the somite. Spinal cord: The posterior continuation of the neural tube. Splanchnic: Of the gut. Splanchnic lateral plate mesoderm: The layer of lateral plate mesoderm lining the coelom and that is associated with the endoderm to form the splanchnopleure. This layer of mesoderm is able to form blood vessels. Splanchnopleure: In an amniote embryo, the layer formed by the splanchnic lateral plate mesoderm and the endoderm. It forms the gut wall as well as the yolk sac and allantois.

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Subcephalic pocket: Literally, a pocket, or space, below the head. In a chick embryo, the space between the head and the proamnion. It is created by the head fold which undercuts the embryo, lifting the head up from the level of the blastoderm. Telencephalon: The anterior region of the forebrain. It is associated with olfaction and is the center of intelligence. More commonly called the cerebrum. Unsegmented somite mesoderm: The region of somitic mesoderm that has not yet formed blocks of somites. Uric acid: The least toxic form of nitrogenous waste. Almost insoluble in water, it is an adaptation for environments where water is scarce. The white pasty material from birds that ends up on your car’s windshield is uric acid. Ventral aortae: In the vertebrate embryo, the blood vessels leaving the heart and connecting to the dorsal aortae by way of paired aortic arches. Ventricle: The region of the heart that receives blood from the atrium and delivers it to the conus arteriosus. Vitelline veins: The veins coming from the extraembryonic regions that enter the posterior chamber of the heart. Whole mounts: A histological preparation of a specimen in which the whole specimen is mounted. The specimen is usually mounted on a microscope slide either in a water-mountant such as glycerin jelly or a non-water mountant such as the plastic mounting media used routinely for stained paraffin sections. Yolk sac: The extraembryonic membrane that surrounds the yolk. In the amniote, it is vascularized so that it can bring yolk nutrients from the yolk back to the embryo proper. It forms from the splanchnopleure, a layer made up of endoderm and splanchnic lateral plate mesoderm.

Chick: 33-Hour Embryo Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. By 33 hours of incubation, what subdivisions of the brain are recognizable in the chick embryo?

2. What three different structures form as outpocketings from the diencephalon?

3. By 33 hours of incubation, what are the major subdivisions of the heart in the chick embryo?

4. What is the blood vessel that takes blood from the yolk sac into the heart?

5. What is the notochord, and what are some of its functions in the embryo?

Name: _______________________ 6. What are somites, and what do they form?

7. What will the nephrogenic (or intermediate) mesoderm form?

8. What are the layers that make up the walls of the body coelom?

9. In the chick embryo, what blood vessel leads blood out of the heart?

10. In the 33-hour chick embryo, how far anteriorly does the notochord extend?

Chick: Mid - 2

11

chapter

The Living Embryo and Making of W hole Mounts

72- and 96-hour Chick Embryo

So far you have studied mounted specimens of 24- and 33-hour chick embryos. You now will be observing living chick embryos at 72 and 96 hours of incubation (comparable to the fifth week of development in humans) and learning how to make whole mounts of your own. Your whole mounts can be every bit as good as, and in some cases better than, those bought from laboratory supply houses. You will notice that the procedures for making whole mounts cannot be completed in a single day. Break-points, where you can let your specimens sit until the following laboratory period, are clearly indicated. It will take three laboratory sessions to complete your slides. In the second and third sessions, however, the manipulations are brief and will not interfere with your other laboratory assignment for that week. Just don’t forget to do them. In proceeding through the exercises, it is advisable to complete observations on one embryo and then fix it before going on to your second embryo. Leave the second embryo in the incubator until you are ready to use it.

Incubation of Eggs Most likely, eggs already have been incubated for you. However, as a scientist, you must be able to repeat all aspects of a study, so here are the basics of egg incubation. Eggs are normally incubated in a forced-draft (it has a fan), humidified (57% relative humidity) incubator at 37.5–38°C. These incubators come in all levels of sophistication. The old-fashioned, cedarbox incubators, still used by many small chicken farmers, consist of a thermostated heating coil with a small fan that circulates air within the box. Humidity is controlled very simply by placing pans of water in the incubator. These incubators must be checked daily to add water and manually turn the eggs. Turning prevents the embryo from becoming stuck to the shell. Modern mechanized incubators don’t require daily checking, since they include an automatic water supply and turning device. Very simple models, however, can also work well. I have even seen boxes rigged with lightbulbs on a rheostat that work perfectly adequately, especially if the eggs do not have to be maintained until hatching. However simple or complex, the incubator must have good temperature control. Though 37.5°C (99.5°F) is ideal, eggs will tolerate temperatures between 35°C and 40.5°C (95–105°F). Within this range, the cooler temperatures significantly slow the rate of development, and the warmer temperatures accelerate it. A humid atmosphere is also essential in order to avoid dehydration of the egg. Though 57% relative humidity is considered ideal, eggs will tolerate a range of 50–60%. Relative humidity can be determined by reading the difference between a wet-bulb and a dry-bulb thermometer. The dry-bulb is a normal thermometer. The wet-bulb is simply a thermometer whose bulb is covered by a wick kept moist with water. This records a cooler temperature due to evaporation from the wick. The less humid

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the air, the greater the cooling. When the incubator is 37.5°C (99.5°F) and the wet-bulb thermometer reads 30°C (86°F), the relative humidity is approximately 57%. Normally, just keeping a large pan full of water in the incubator is sufficient to maintain the appropriate humidity. Check this pan daily. It should never go dry. Seldom is it convenient to start incubation on the day that an egg is laid. Luckily, eggs can be stored prior to incubation, though hatchability diminishes as the storage time increases. (By 22 days of storage at 10.5°C, hatchability is down to 26%, and by 25 days, it is 0%.) If the eggs are to be stored for less than 14 days, the temperature of the holding room should be 18.3°C (65°F), and the humidity should be as high as 80%. If they are to be stored for longer than 14 days, the holding room should be 10.5°C (51°F). When the eggs are removed from the holding room, they should warm to room temperature before being placed in the incubator. Eggs should be incubated with their blunt side uppermost. This can be done most easily by leaving the eggs in the cardboard flats that they come in. The eggshell should be labeled in pencil to indicate the day the egg is put into the incubator. For proper development, eggs should be turned several times a day—as many as six. It is best to turn them to a 45° angle from their vertical position. If incubating for only 3 or 4 days, as in this study, turning is unnecessary. During incubation, eggs can be candled to determine whether they are developing or not. A candler is no more than a light in a box shining through a hole the diameter of an egg. You can make a candler by putting a light in a wooden box that has a hole the size of an egg cut in one end. Candling is best done in a dark room. Move the egg around, holding it against the lighted hole of the candler. In the living embryo, vascularized regions will be particularly obvious. A dead embryo will look dull and dense. If the eggs are being kept to hatching, the temperature of the incubator ideally should be reduced a degree or two during the last 2 days of incubation (it takes 21 days to make a chick). The chicks will be breaking through the blunt end of the shell. After the chick has “pipped” the shell (made the first, small hole), it will rest for hours or even days before finishing the job, which may take as little as an hour.

Preparing Whole Mounts The eggs you will be using for whole mounts should not be turned while incubating and should have been positioned with their blunt end up. Because the yolk orients to gravity with the embryo uppermost, the embryo should now be sitting just below the air space at the blunt end of the egg. This makes it easily accessible for observation and dissection. Get one of your eggs that has been incubated for 3 or 4 days from the incubator and place it in a “professional egg-dissection holder” (i.e., the bottom half of an egg carton cut to contain three or four depressions for eggs). Make sure that the blunt end of your egg is still facing upward. (You may candle it first to determine exactly where your embryo lies.) You will be opening the egg simply by attacking it as you would a really soft boiled egg at breakfast. With the handle of your forceps, tap the blunt end of the egg five or six times, cracking a nickel-shaped region of shell into a number of small pieces. Remove the broken bits of shell with the forceps. The outer shell membrane will be removed with the shell. Now with fine forceps, very carefully remove the tough white inner shell membrane overlying the embryo. The embryo is now exposed. Observe the embryo under the dissecting scope. The heart should be beating. Notice the jerkiness of blood movement in the arteries and the smooth flow in the veins. Use Figures 11.1 and 11.2 to identify embryonic structures and extraembryonic regions. You can make a number of observations before proceeding further. Ask as many questions as you can think of. Record both your questions and answers in your notebook. For example, what is the heart rate? Use a watch

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with a second hand to record rates. Is there both an atrial and ventricular beat? Do you see skeletal muscular movements? Does the embryo respond to touch? Does the amnion or chorion respond to touch? Does warming the egg under a lamp increase or decrease muscle or heart movement? Does cold Ringer’s solution? Record any rate changes. Does alcohol affect the heartbeat? You could test ranges that are found in the blood of a person considered legally drunk (0.1% or greater; at levels of 0.05–0.09%, a person is considered “under the influence”). Remember that alcohol is a fixative—don’t use higher percentages that might kill the embryo. Does caffeine or theophylline alter the heartbeat or muscle movements? You can test this using dilutions of the pure chemicals or with drinking strengths of coffee (containing caffeine) and tea (containing caffeine and theophylline). A good cup of coffee contains approximately 0.5–1 mg of caffeine per ml, with instant having the least and drip-prepared having the most. A cup of tea has about 0.3 mg of caffeine and 0.04 mg of theophylline per ml. Remember what these chemicals do. They increase cyclic adenosine monophosphate (cAMP) levels in cells by inhibiting the enzyme phosphodiesterase that breaks down cAMP. Increasing or decreasing cAMP levels can have profound effects on a cell. Cyclic AMP levels increase in a cell that is no longer dividing, for example, and such increases precede programmed cell death. Low levels of cAMP occur in dividing cells and are maintained at these low levels in cancer cells. What do you imagine the effect is on an embryo (in which cell division is so essential) of raising cAMP levels over long periods of time?

Metencephalon Myelencephalon Otic vesicle

Mesencephalon

Aortic arch

Optic cup

Pharyngeal pouch

Lens Diencephalon

Atrium

Epiphysis

Somite

Telencephalon Wing bud

Olfactory placode

Ventricle Vitelline artery

Spinal cord

Vitelline vein

Leg bud

Tail bud

Figure 11.1 Schematic diagram of a 72-hour chick embryo.

The Living Embryo and Making of Whole Mounts

Otic vesicle

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Myelencephalon Metencephalon

Pharyngeal pouch

Optic cup

Aortic arch

Lens Mesencephalon

Atrium

Diencephalon Wing bud Vitelline blood vessels

Ventricle

Epiphysis Somite Allantois

Figure 11.2 Schematic diagram of a 96-hour chick embryo.

Telencephalon

Leg bud Amnion

What is your advice to pregnant women on drinking alcohol, coffee, or tea? (N.B.: None of the substances listed should be injested by a pregnant woman. In particular, no level of alcohol is considered safe for the embryo or fetus.) Think of other interventions you might do to determine more about the embryo. Record all your observations in your laboratory notebook. Organize your data into graphs wherever possible. What conclusions do you draw from your data? You are now ready to remove the embryo from the yolk. This often requires three hands, especially on the first try. You will need to cut the embryo free from its extraembryonic membranes while holding on to it, so that it doesn’t slip into the murky depths of the yolk once it is freed. If the embryo is large enough to have an identifiable neck region, you can slip your forceps underneath the neck. With your other hand and a pair of very sharp scissors, cut the extraembryonic membranes around the embryo. Now slip your embryo spoon under the embryo. This is where a third hand standing by often saves the day. Do not let go of the embryo with the forceps, even though the embryo seems safe on the spoon. Lift the embryo carefully from the yolk. Often as you lift the embryo away, you will see that it isn’t completely free from the membranes. Have scissors ready to cut any remaining strands. Transfer the embryo to a petri dish containing Ringer’s solution (a balanced salt solution). Chick embryo Ringer’s solution NaCl 7.20 gm KCl 0.37 gm CaCl2 0.17 gm

Make up to 1 liter in glass using distilled water.

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If your embryo is too young to have an identifiable neck, an easy way to remove it is first, to use your forceps to grab on to the extraembryonic blood vessels that feed into the heart—and don’t let go. Then, use sharp scissors to cut around the embryo, cutting the extraembryonic membranes so that the embryo is freed from the yolk. Without moving the embryo, put the scissors down and pick up your embryo spoon (or use a partner to provide a third hand). Slip the embryo spoon under the embryo as you gently pull the embryo onto the spoon using the forceps (that are still holding onto those blood vessels). Now pull the whole thing away (still holding on with the forceps), having scissors ready to snip any strands of membrane that are still attached. Place the embryo into a petri dish containing Ringer’s solution. Make observations on the embryo removed from its yolk. You should be able to distinguish more features. Stage your embryo using the Hamburger-Hamilton staging series (see Table 10.1). In your laboratory notebook, record the stage and the number of somites your embryo has. Observe the heartbeat again. Has it changed? If you warm the embryo under a lamp, does this change the heartbeat? This is your last chance to make as many observations as you’d like on the living embryo. Be creative. When you have completed your observations, go on to the fixation step, then repeat the above steps on your second embryo.

Fixation Fixation is the first step in any procedure in which tissue is to be preserved for histological study. Fixatives kill. They kill the tissue as well as any bacteria that are present that otherwise would cause the tissue to rot. They also coagulate proteins, making them insoluble, and strengthen protein linkages. All fixatives distort the tissue to a certain extent, but in general, protein and cell structure are preserved. You will normally choose a fixative containing several ingredients that balance out the illeffects of each other. For example, alcohol shrinks tissue and causes excessive hardening. This can be countered by adding acetic acid, which swells tissue and prevents overhardening. Carnoy’s fixative glacial acetic acid 100% ethanol

100 ml 300 ml

Glacial means 100% acetic acid; vinegar is 5% acetic acid. Hard liquor such as whiskey is about 40%–50% ethanol. Rubbing alcohol is 70% isopropyl alcohol— don’t drink it, it’s toxic. If you were out in the boondocks with no scientific supplies and found the perfect specimen you needed to preserve, what makeshift fixative would you devise?

Carnoy’s fixative, a mixture of alcohol and acetic acid, is a widely used fixative. It is not ideal; the addition of formaldehyde, for example, would give better preservation of cytological detail. But Carnoy’s fixative will not leave any toxic residues in the lab. Formaldehyde and most other fixing agents leave highly toxic residues that are virtually impossible to remove from instruments and glassware. In making whole mounts, where cytological detail is not critical, Carnoy’s becomes ideal for our use because there is no risk of contaminating our lab for future live material. (You may be using one

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of your embryos for the histological sectioning procedures discussed in the next chapter. If so, you might be fixing in something other than Carnoy’s. Check with your instructor before proceeding.) With a pipette, drop fixative on top of the embryo until all regions have been covered. This will allow any extraembryonic membranes to fix flat before flooding the specimen with fixative. After 10 minutes, flood the embryo with fixative. Wait another 10 minutes, then transfer the embryo to a screwcap vial. For fixation, and for all steps that follow that require a fluid, the rule of thumb is that the amount of fluid used should be 10 times the volume of the tissue. Label your vial with your name and the stage of your embryo. Always use pencil for labeling—it doesn’t come off in the reagents. Your embryos should fix for a minimum of 2 hours. They can be left in the fixative a maximum of 2–3 days. Following fixation, your embryos will be washed and stained, then dehydrated and cleared before mounting. All of these operations can be done in the vial. To make a transfer of solution, you can either pour off the old solution (always over a finger bowl—never over the sink; guess why), or you can pipette off the old solution. The new solution is added directly to the vial. This avoids unnecessary handling that can damage the embryo.

Washing You will be washing out the fixative using 70% ethanol. Washing is often done in water. If you have used Carnoy’s as a fixative, washing in 70% alcohol is both adequate and fast. You also can use it as a holding solution. The embryos can remain in 70% alcohol indefinitely without harm. Remove the fixative and add 70% ethanol to your vials. Your embryos should be in this for a minimum of 8 hours. Keep the embryos in 70% ethanol until next week’s lab. You should change the solution at least once during the week to ensure that all fixative is washed out of the tissue. Under current OSHA guidelines, any waste alcohol solutions containing over 24% alcohol must be handled as hazardous waste and may not be poured down the sink.

Staining (To Be Done the Week Following Fixation) You will be using a whole-mount staining procedure that is exceedingly stable and will not overstain your embryos. Before staining, pour off the 70% alcohol and add distilled water to your vials. Let the embryos soak for at least 10 minutes before transferring them to the stain. Then pour off the water and add Mayer’s carmalum stain. This is a general nuclear stain that is excellent for whole mounts. Stain for about 48 hours. The timing is not especially critical. The embryos should be a fairly deep red. Once the desired level of staining is reached, pour off the stain and add 70% ethanol to the vials. You will be storing the embryos in 70% ethanol until next week’s lab. Mayer’s carmalum Stock solution: carmine ammonium alum (aluminum ammonium sulfate) distilled water Working solution: carmalum stock solution glacial acetic acid distilled water

1 gm 10 gm

200 ml

Boil 20 minutes.

5 ml 0.4 ml 100 ml

Mix. When dissolved, filter. Add 1 ml formalin to inhibit mold growth. Will keep indefinitely.

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Dehydration and Clearing (To Be Done the Week Following Staining) It is necessary to dehydrate the embryos because the mounting medium used here will not mix with water. Rather than removing the water from your specimens in a single step, which could damage the tissue, you will use a graded series of alcohols to slowly replace the water with alcohol. This avoids severe convection currents that are set up as one fluid replaces the other. Following dehydration, the embryos must be cleared in a clearing agent. You could use xylene or toluene. Toluene is gentler on the tissue, but is more expensive, and in making whole mounts the gentleness of the clearing agent is not critical. Clearing will make the embryos transparent. Clearing is also necessary because the mounting medium will not mix either with water or with alcohol. Xylene or toluene, which are immiscible with water, will mix with both 100% alcohol and the mounting medium. These clearing agents therefore provide a bridge for us between the dehydration steps and mounting. Dehydrate and clear your specimens according to the following schedule. For each transfer of fluid, pour off the old fluid and add the new directly to the vials. Used alcohol at concentrations above 24% must be discarded as hazardous waste according to OSHA guidelines. Xylene and toluene are both very toxic and also must be disposed of as hazardous waste. These agents dissolve plastic and will dissolve plastic pipes if thrown down the sink; they will also dissolve plastic petri dishes or vials, so make sure the vials you are using are glass. Because of their toxicity (they rot your liver), the clearing step must be done under a fume hood, or do it outside. If, on your first transfer into xylene, you notice any white clouding of the fluid, this means that there is still water in the tissues. You must immediately go back to another 15-minute change of 100% ethanol. If you don’t, there will be water droplets in your preparation that will make your slide look like salad dressing. Dehydration and clearing schedule 70% ethanol 10 minutes 95% ethanol two changes, 10 minutes each 100% ethanol two changes, 15 minutes each xylene two changes, 10 minutes each

Mounting The mounting medium you use should be a neutral mounting medium, such as Histoclad. Cheaper mounting media are acidic. They cause the stain to fade and oxidize very quickly, making a cloudy ring around the edge of the slide that gradually works its way to the middle. Any slide that needs to be permanent is worth the extra cost of a neutral mounting medium. The hardest part of mounting is devising a method to raise the coverslip enough so that it isn’t resting directly on the specimen. If your specimen is thin, you can use chips of coverslip as a spacer between the slide and coverslip. If your specimen is thick, pieces of thicker glass such as a slide could be used. I have had wonderful success using slices of thin polypropylene tubing cut into doughnuts that are the thickness of the specimen. I have even used metal washers, though plastic washers would be better. Devise a method that will work for you.

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Once you start the mounting procedure, work with speed. Do not let your specimen dry out. It will do so very quickly. First, with a diamond marking pen (a drill-bit with a carbide tip works as well), scratch your initials and the stage of your embryo onto one end of a slide. Then lift your embryo from its vial of xylene and place it on the slide. Position the spacer around the embryo. Put a generous amount of mounting medium over the specimen and carefully lower a square coverslip into place, letting one side hit the mountant first and then slowly lowering the rest of the coverslip. If you see that there isn’t enough mounting medium, more can be introduced from the side by placing extra mountant alongside the coverslip and letting it wick underneath. If you have too much mounting medium and it is oozing out, don’t touch it. After it dries for a week, excess mountant can be removed with a razor blade (carefully). Allow your slides to dry flat in a dust-free environment (such as a small cardboard box) at room temperature for at least a week before observing them under the microscope. When you do observe them, restage the embryos to see if your staging of the living embryos was correct. It is far easier to stage an embryo in a whole mount than one that is unstained and uncleared. You now should have some very beautiful, permanent slides.

Accompanying Materials See Vade Mecum: “Chick Late.” This chapter of the CD shows movies of the technique for removing the embryo from the yolk, as well as labeled whole mounts of 72- and 96-hour chick embryos, and a series of movies about the living embryo.

Selected Bibliograpy Hamburger, V. and H. L. Hamilton. 1951. A series of normal stages in the development of the chick embryo. J. Morphol. 88: 49–92. This is the original publication of the Hamburger-Hamilton chick staging series. Humason, G. L. 1972. Animal Tissue Techniques, 3rd Ed. W. H. Freeman, San Francisco. This is one of the best sources on histological techniques—it is out of print, but has been reissued by John Hopkin’s University Press (see reference below). Presnell, Janice K., Gretchen L. Humason and Martin Schreibman. 1997. Humason’s Animal Tissue Techniques, 5th Ed. Johns Hopkins Univ. Press, Baltimore. The reissuing of Humason’s classic. Romanoff, A. L. 1960. The Avian Embryo. Macmillan, New York. This book has an extensive chapter on the embryonic heart, including experimental results on changes in heart rate under conditions of altered temperature and chemical environment.

Suppliers Fertile chick eggs are most easily obtained from local hatcheries. If this is not possible, eggs can be shipped from supply companies such as those listed below. These companies also have inexpensive egg incubators. More expensive, cedar-box incubators can be bought from farm suppliers.

Connecticut Valley Biological Supply Co., Inc. P.O. Box 326 82 Valley Road Southampton, MA 01073 1-800-628-7748

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Nasco 901 Janesville Ave. Fort Atkinson, WI 53538- 0901 1-800-558-9595 www.nascofa.com

Ward’s Natural Science Establishment, Inc. 5100 West Henrietta Road P.O. Box 92912 Rochester, NY 14692-2660 1-800-962-2660 www.wardsci.com

Chemicals such as caffeine and theophylline can be obtained from most biochemical suppliers, such as: United States Biochemical Corp. P.O. Box 22400 Cleveland, OH 44122 1-800-321-9322

A pH-neutral mounting medium is difficult to find. Do stay away from Permount, which is acidic. Neutral or near-neutral mounting media are supplied by: Ward’s Natural Science Establishment, Inc. 5100 West Henrietta Road P.O. Box 92912 Rochester, NY 14692-2660 1-800-962-2660 www.wardsci.com

Glossary Amnion: The extraembryonic membrane found in amniotes that surrounds the embryo. It contains amniotic fluid, which protects the embryo by providing an isotonic fluid and buffering the embryo against being jarred. It forms from the somatopleure, a layer made up of epidermal ectoderm and somatic lateral plate mesoderm. Atrium: The region of the heart that receives blood from the sinus venosus and delivers it to the ventricle. Caffeine: A drug found in foods such as coffee, tea, and chocolate. In its purified form, it is white and crystalline. A bitter alkaloid, C8H10N4O2. One of its biological effects in higher organisms is to raise cAMP levels by inhibiting phosphodiesterase, the enzyme that breaks down cAMP. cAMP: Cyclic AMP.

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Candler: A devise that allows one to examine the contents of a chick egg to determine whether the embryo inside is alive or not. It consists of a box containing a light, and a hole in the box roughly shape of an egg. By holding the egg against the hole, one can see the condition of the embryo within. Candling: Using a candler to check the condition of a chick egg. Carnoy’s fixative: A fixative that contains glacial acetic acid and ethanol. Though a harsh fixative on tissues, it has the advantage of not leaving any toxic residues behind once it has evaporated. Chick embryo Ringer’s: A physiological salt solution that is balanced for a chick embryo so that it can maintain the embryo in vitro. Chorion: The extraembryonic membrane found in amniotes that lies outside of the amnion. It protects the embryo, and in the chick will eventually fuse with the allantoic membrane. It forms from the somatopleure, a layer made up of epidermal ectoderm and somatic lateral plate mesoderm. Clearing: The process in a histological preparation in which tissues become clear and translucent. Common clearing agents include xylene and toluene, which are immiscible with water and require that the tissues be dehydrated prior to clearing. Glycerin is a clearing agent that is miscible with water and can be used without dehydrating the specimen. Coverslip: A thin transparent cover that is placed over a specimen on a microscope slide. They are usually made of glass and are of the same refractive index as the mounting medium used for making histological slides. Cyclic adenosine monophosphate (cAMP): Cyclic AMP. Cyclic AMP: Cyclic adenosine 3´,5´-monophosphate, made from ATP. Within a cell, it is used as a second messenger. Rarely, as in the aggregation of Dictyostelium amoebae, it is pulsed to the outside of the cell and in this circumstance causes directional cell migration up a gradient of cAMP. When cAMP occurs outside of a cell, it does not normally enter a cell. Experimental forms of cAMP, such as dibutyryl cAMP, are capable of entering the cell. Dehydration: The removal of water. In histological procedures, a specimen is usually dehydrated through a graded series of alcohol prior to embedding or mounting. Diamond marking pencil: An instrument in the shape of a pen, that has a chip of industrial-grade diamond in its marking tip. It is used to scratch marks into glass, in particular, a glass slide that must be labeled in a permanent manner. Embryo spoon: A small perforated spoon that allows embryos to be picked up without their floating off the spoon. It can be made inexpensively from a plastic spoon and a heated needle, used to perforate the bowl of the spoon. Extraembryonic membranes: The membranes that form in association with the embryo proper. In the amniote, four extraembryonic membranes form, the amnion, chorion, yolk sac, and allantois. Extraembryonic regions: Embryonic regions that do not contribute to forming the body of the embryo proper. These regions form the extraembryonic membranes. Fixation: The process of fixing a tissue. Fixation kills the tissue, as well as any bacteria present, and coagulates (cross-links) proteins. Fixative: Anything that fixes a living organism so that it can’t be degraded by bacteria or other microorganisms. Fume hood: A chamber that contains a ventilation fan and is vented to the outside, so that fumes of substances within the chamber won’t enter the room. Incubator: A chamber with a controlled environment where organisms or tissues can be grown. In vitro: Literally, in glass (Latin). It refers to the condition of being outside of the living organism in a laboratory culture condition, such as in a petri dish.

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In vivo: Literally, in life (Latin). It refers to the condition of being within the living organism. Mayer’s carmalum: A histological stain containing carmine. It is a general nuclear stain that is excellent for whole mounts. Mounting medium: The medium in which a specimen is mounted for observation under the microscope. A mounting medium can be water soluble, such as glycerin jelly, or immiscible with water, such as a plastic polymer dissolved in a solvent. When a mounting medium dries, it is firm and transparent, and typically has the same refractive index as glass. OSHA: Occupational Safety and Health Administration, a division of the U.S. Department of Labor. Pipping: At hatching, the process of a baby bird breaking through its shell. Shell membranes: Membranes on the inner aspect of the calcareous portion of an egg shell. In the chick egg, there are two shell membranes. These protect against desiccation and invasion by microorganisms. Stained: In histological procedures, this means dyed with histological dyes to contrast various components of cells and tissues. Theophylline: A drug found in tea. In its purified form, it is white, and slightly water soluble. It is an alkaloid, C7H8N4O2. One of its biological effects in higher organisms is to raise cAMP levels by inhibiting phosphodiesterase, the enzyme that breaks down cAMP. Ventricle: The region of the heart that receives blood from the atrium and delivers it to the conus arteriosus. Washing: In histological procedures, this is the step following fixation in which the fixative is washed out of the tissue. Whole mounts: A histological preparation of a specimen in which the whole specimen is mounted. The specimen is usually mounted on a microscope slide either in a water-mountant such as glycerin jelly or a non-water mountant such as the plastic mounting media used routinely for stained paraffin sections.

Chick: Late Development Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. Do you think that it could be dangerous to the fetus for a pregnant woman to take a very hot bath? Why? What have you observed in chick embryos that makes you draw this conclusion?

2. What changes do you see in the 72-hour chick embryo that make it look different from a 33hour or 48-hour chick embryo?

3. The ganglia of what cranial nerves can be seen in a 72-hour chick embryo?

4. What sensory structures are visible in the 72-hour chick embryo?

Name: _______________________

Chick: Late - 2

5. What changes do you see in the 96-hour chick embryo that make it look different from a 72hour chick embryo?

6. The ganglia of what cranial nerves can be seen in the 96-hour chick embryo?

7. At what temperature and what humidity do you incubate chick eggs?

8. How do you candle an egg, and what does it show you?

9. How would you go about making a homemade candler? How much do you think the raw materials for this would cost?

10. What is the best way to orient eggs when incubating them if you will be opening them later for observation? Why is this orientation advantageous?

12

chapter

Histological Techniques

Have you ever been awed by the histologist’s ability to create a complete set of serial sections through an embryo without loosing a single section? This may seem an impossible feat until you learn the relatively easy methods of paraffin sectioning. In this chapter you will have the opportunity to make your own set of serial sections, and in the process, you will be learning a technique that has recently regained the spotlight in embryological studies. With the advent of immunohistochemistry and fluorescence microscopy, developmental biologists have a valuable set of tools, and they need to know the art of tissue preparation to apply them. Regardless of the staining technique used, the basic process is the same: Tissues are first fixed to avoid degradation, embedded in paraffin, cut into serial sections with a rotary microtome, and stained to contrast cellular elements. The paraffin technique for tissue preparation is the most commonly used for routine study of tissue. You will be using either chick embryos that you obtained from your previous laboratory exercise (Chapter 11) or other tissue provided by your instructor.

Fixation Fixation is the first step in any procedure in which tissue is to be preserved for histological study. Fixatives kill. They kill the tissue, as well as any bacteria that are present that otherwise would cause the tissue to rot. They also coagulate or cross-link proteins, making them insoluble. All fixatives distort tissue to a certain extent, but in general, proteins and cellular structure are preserved. Normally you choose a fixative containing several ingredients that balance out each other’s ill-effects. For example, alcohol shrinks tissue and causes excessive hardening. You can counter these effects by adding an acid such as acetic acid, which swells tissue and prevents overhardening. One of the safest fixatives to use, which will not leave any toxic residues behind, is Carnoy’s fixative, a mixture of alcohol and acetic acid. It is not an ideal fixative. The addition of formaldehyde, for example, would give better preservation of cytological detail. Formaldehyde, however, and most other fixing agents leave highly toxic residues that are virtually impossible to remove from instruments and glassware. If you were making whole mounts, where cytological detail is not critical, or you cannot risk contaminating your work area for future live material, Carnoy’s is ideal for use. Carnoy’s fixative* glacial acetic acid 100% ethanol

100 ml 300 ml

Some Carnoy's recipes also include chloroform along with glacial acetic acid and absolute ethanol in the proportions 3:1:6. Chloroform increases the speed of fixation, making it useful for fixing tissues that are difficult to penetrate. Chloroform, however, is extremely hazardous and must be used in a ventilated hood while wearing gloves.

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Glacial means 100% acetic acid; vinegar is 5% acetic acid. Ethanol is the type of alcohol that people get arrested for driving under the influence of; hard liquor such as whiskey is about 40%–50% ethanol. Rubbing alcohol is 70% isopropyl alcohol—don’t drink it, it’s toxic. If you were out in the boondocks with no scientific supplies and found the perfect specimen you needed to preserve, what makeshift fixative would you devise? A commonly used fixative that is better at preserving structure than Carnoy’s is FAA, made of formaldehyde, alcohol, and acetic acid. (Formaldehyde is a gas, which is sold in solution as formalin, which is about 40% formaldehyde in water.) Dietrich’s FAA** 95% ethyl alcohol formalin glacial acetic acid distilled water

30.0 ml 10.0 ml 2.0 ml 60.0 ml

A fixative that is widely used for embryological material, since it will not overharden yolky material, is Bouin’s fluid. It has the advantage that tissues may be stored indefinitely in it. Its major disadvantage is that the picric acid, though an excellent fixative for lipid-rich tissues that doesn’t harden the tissues, also stains the tissue yellow. The yellow color must be removed before other stains can be applied. Usually, this is accomplished after sectioning as the slides descend through the alcohol series. Just leave the slides in 70% alcohol for a longer period of time if the yellow color doesn’t disappear using the normal dehydration schedule. Bouin’s Fluid** picric acid, saturated 75.0 ml aqueous*** formalin 25.0 ml glacial acetic acid 5.0 ml To fix your embryo or sample of tissue, place the freshly dissected specimen (it should ideally be no more than 1–2 cm on a side) into a screw-cap vial containing your fixative of choice. For fixation and all steps that follow that require a fluid, the rule of thumb is that the amount of fluid used should be 10× the volume of the tissue. Label your vial with your name and what the specimen is, or use a piece of cardstock as your label and place it in the vial with your specimen. Always use pencil for labeling—it doesn’t come off in the reagents. The amount of time your specimen is left in fixative should depend on the size of the specimen and the type of fixative. Most specimens should be fixed 6–8 hours, or overnight. If the specimen is smaller than 5 mm on a side, then several hours will suffice. If the fixative severely hardens and shrinks the * Note that under current OSHA guidelines, solutions containing concentrations of alcohol above 24% must be discarded as hazardous waste. ** Under current OSHA guidelines, used fixatives containing formalin must be discarded as hazardous waste, as must solutions containing concentrations of alcohol above 24% . ***Picric acid is sold saturated in water and must be stored that way, since in dry form it is explosive. (The exceedingly destructive explosion in Halifax harbor during World War I was caused by sparks from a careless collision into a munitions ship loaded with picric acid.)

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specimen, as does Carnoy’s, then the minimum fixation time should be used. Specimens should not be left in Carnoy’s fixative more than 2–3 days. If the fixative is like Bouin’s, which is not harsh on the tissue, the tissue may be stored in the fixative for extended periods (months or more) without severe alteration. Tissues should be fixed and stored in the refrigerator at 4ºC.

Washing Following fixation, your tissue sample must be washed. This, as well as many of the operations that follow, can be done in the vial. To make a transfer of solution, you can either pour off the old solution into a finger bowl, or you can pipette off the old solution. The new solution is added directly to the vial. This avoids unnecessary handling that can damage the tissue. Washing is usually done in water. For most fixatives, washing specimens in distilled water for 6–8 hours, or overnight, following fixation is sufficient. If you have chosen Carnoy’s as your fixative, then washing in 70% alcohol instead of water is both adequate and faster. You also can use the 70% alcohol as a holding solution. Following washing, the specimen should be transferred to 70% alcohol and stored at 4ºC until it can be processed further. It can remain here for a number of weeks.

Dehydration and Clearing To prepare the specimen for paraffin embedding, the specimen must be dehydrated through a series of alcohols up to absolute alcohol (ethanol is preferable to methanol, since it is less harsh on the tissues). This removes all the water, which is immiscible with paraffin. By going through a graded series of alcohols, the convection currents that are set up by each transfer are minimized, thereby minimizing harm to the specimen. After the water has been removed, a clearing agent, such as xylene or toluene, which is miscible with both 100% alcohol and paraffin, makes a bridge between the alcohol and paraffin. Toluene is less harsh on tissue than xylene, causing less shrinkage and hardening, and so should be used instead of xylene if possible. In transferring the specimen through any alcohol solution, the old solution can be poured off (into a finger bowl) and the new solution added. Remember that under current OSHA guidelines, solutions containing concentrations of alcohol above 24% must be discarded as hazardous waste. Xylene and toluene are toxic. They should only be used in a ventilated hood (or outside, if necessary) and also must be put into special waste jars and disposed of according to OSHA regulations as a hazardous material. Making a graded series of alcohol: Always use 95% ethanol (ETOH) to dilute from, rather than 100% ETOH. Absolute (or 100%) ethanol is exceedingly expensive compared to 95%. Here is an easy method for making any percentage of alcohol below 95% that you wish. To make a 70% solution of ETOH, for example, pour 95% ETOH into a graduated cylinder up to the 70 ml line, then add distilled water until the solution reaches the 95-ml line. This gives you 95 ml of 70% ETOH. If you are making a 50% solution, pour 95% ETOH up to the 50-ml line, and add distilled water up to the 95-ml line. To make larger amounts, just use quantities that are multiples of the numbers above. The following schedule can be used. Times are sufficient for specimens that are no more than 1–2 cm/side. If the sample is larger, longer time periods must be used.

Histological Techniques 70% alcohol (if starting from water) 95% alcohol 100% alcohol (first time) 100% alcohol (second time) Toluene or xylene (first time)* Toluene or xylene (second time)

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1–2 hours 1–2 hours 1/2–1 hour 1/2–1 hour 1/2–1 hour 1/2–1 hour

* Warning: If there is any water left in the specimen at this point, it will show as a milkiness around the specimen when it is placed in the clearing agent. The specimen must be brought back to 100% alcohol in this case to remove the remaining water.

Special microwaving techniques that save time Microwaving the specimen during fixation and dehydration cuts down considerably on the time needed in each solution and improves infiltration of the solutions into the tissue. A number of precautions must be taken, however, to make sure that the specimen doesn’t overheat, that penetration is uniform, and that toxic exhaust doesn’t enter the room. A large beaker of water should be placed in the oven to absorb excess microwave energy. The specimen should not be placed in a “hot” spot of the oven. You can figure out where the hot spots are by placing wet paper towels in the oven and turning it on. Just as the towels begin to dry, turn off the oven. Wherever wet spots remain are good places for placing your specimen vial. Alternatively, the specimen vial can be placed on a rotating table in the oven. To avoid overheating, the oven can be put through cycles of being run on “high” for short periods (7–10 seconds) alternated with being off for 20 seconds. The specimen should not be heated over 70ºC—this causes overdenaturation of regions that are relatively impermeable and stain nonuniformly. Putting the vial on a petri dish of ice during processing can prevent overheating. After 5 minutes of microwaving, a specimen 5 mm × 5 mm × 2 mm can go on to the next step in the fixation or dehydration series. Since microwave ovens vary, you will need to experiment to determine what times and settings work best for your particular oven. It is exceedingly important that the seals on the oven be checked to make sure that microwave radiation and toxic fumes are not leaking out into the room. It is safest not to use microwave processing on steps that involve higher alcohols, or clearing agents such as xylene or toluene. Microwaving can also be used in place of chemical fixation in order to avoid the toxicity of chemical fixatives. Specimens are placed in a saline solution, such as chick Ringer’s solution, and microwaved for a period of time, maintaining a temperature of 68–70ºC. Small specimens can be microwaved for as little as 6 minutes. Again, you will need to experiment to determine the times and settings that work best for your particular oven.

Paraffin Infiltration From the second change of toluene, the specimen is to be moved through several changes of melted paraffin. The paraffin you use will have a specified melting point. A melting point of 54ºC is typical and is suitable for sections that will be cut at 8 µm or thicker. Higher melting points indicate harder paraffin that can be cut into thinner sections. Typically, the melted paraffin is kept in a paraffin oven or paraffin bath that will maintain the paraffin at a constant temperature. The specimen can be transferred to a cage specifically designed for infiltration. Be sure to include a label in the cage, along with the specimen, that distinguishes your sample from other samples in the oven. The cage can then be transferred from one bath of melted paraffin to another using forceps or a string tied to the cage. Alternatively, deep-welled spot dishes containing melted paraffin can be used in a paraffin oven and the specimen transferred from one well

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to the next. In this method, use a narrow piece of cardstock that you hold with forceps to lift the specimen for each transfer—this avoids the harm to the specimen that would occur if a metal instrument such as forceps were used to pick up the specimen. The paraffin should be kept above its melting point during this process, but not hotter than necessary, since heat severely shrinks and hardens tissue. Use two to three changes of paraffin, 1/2–1 hour each. Under ideal circumstances, you would be able to infiltrate in a vacuum oven—this gives good infiltration and shortens the time needed in hot paraffin.

Paraffin Molds One of the most successful molds for embedding is a paper box made such that the seams are watertight. Such a box can be made to any size appropriate for your specimen. It also has the advantage, over molds with rigid sides, of being pulled inward along with the paraffin as the paraffin cools and shrinks. This prevents a deep indentation from forming in the center of the block. More expensive manufactured molds made of plastic or metal can also be used, and these can be bought from supply houses. I much prefer paper boxes to any manufactured mold. A paper embedding box should be made of heavyweight paper. It is made most easily by folding the paper over a block of wood that has one face that matches the size that you need. For a specimen approximately 5 mm3 , use a rectangular sheet approximately 6 cm × 10 cm. Place this on a block of wood with a face approximately 2 cm × 2 cm (Figure 12.1A). First fold the long sides of the paper down along the wood (Figure 12.1B). Holding these in place with one hand, use the other hand to fold one of the remaining sides down (Figure 12.1C). Crease the two corners at a 45º angle (Figure 12.1D). This creates flaps, which are then folded inward, and a tab that is folded down (Figure 12.1E). Do the same to the remaining side (Figure 12.1F).

Figure 12.1 Making a paper embedding box. (A) Place a block of wood in the center of a rectangular piece of paper, approximately 6 cm × 10 cm. (B) Fold the long sides of the paper down alongside the wood. (C) Hold the two long sides against the wood with one hand, and use the other hand to fold one of the remaining sides down alongside the wood. (D) Crease the two corners at a 45° angle and fold each flap inward. This creates a tab. (E) Fold the tab downward. (F) Repeat steps C–E on the remaining side.

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Paraffin Embedding Fill the embedding mold with melted paraffin and transfer the specimen from melted paraffin immediately to the mold. The specimen can be transferred using a small strip of cardstock as a spatula. This prevents damage to the specimen. Position the specimen in the mold using a hot needle, heated in the flame of an alcohol lamp. The orientation of the specimen should be appropriate for later sectioning of the block. If using a paper box, this orientation can be marked on the tab of the box, along with information on what the specimen is and any other information necessary to identify it later. After a film of solidifying paraffin has formed over the surface of the paraffin, the mold should be transferred to a bowl of cool (but not ice-cold) water (water that is too cold can cause the paraffin to crack). The water causes the paraffin to cool more quickly, making for smaller crystal formation within the paraffin as it solidifies and hence less distortion of the specimen. Once the block is hard, remove it from the water and store in the refrigerator. You should not try mounting and cutting the block until it has cooled for 6–8 hours, or overnight. This improves the cutting process and is one of those magic tricks one learns but can’t explain.

Mounting and Trimming Paraffin Blocks The paraffin block is to be mounted onto a piece of metal, synthetic fiber, or wood, all of which can be referred to as “chucks,” which fit into the chuck holder of the microtome. When mounting the paraffin block onto the chuck, the chuck must be held securely. A heavy piece of wood with holes cut into it or even a chunk of clay into which you press the chuck are suitable holding devices. Rough trimming Remove the paraffin block from its mold, keeping track of the orientation of the specimen within it. Trim away excess paraffin using a safety razor blade. This should be done on a protective piece of wood or glass. Do not trim too close to the specimen. Leave at least 5 millimeters of paraffin on all sides and more than this on the bottom of the block. Paraffin scraps from trimming can be remelted, filtered, and reused. Paraffin improves with reheating, since impurities are volatilized off with each heating. Mounting the block Using the flame from an alcohol lamp, melt some shavings of paraffin on the blade of a metal spatula and pour the melted paraffin onto the surface of the chuck. In this way, you are to build up a layer (at least 1 mm thick) of paraffin that is firmly attached to the chuck. Heat the spatula again and use it to melt the top of this layer of paraffin. Then heat both sides of the spatula blade, hold it between the bottom of the paraffin block and the paraffin on the chuck so that it melts both surfaces of paraffin, and slide it away as you press the paraffin block against the chuck. Heat the spatula blade again, and use it to melt some paraffin on the four vertical sides of the paraffin block to ensure that it is attached firmly to the chuck. Set aside until the paraffin is firm. Fine trimming You now need to trim the face of the block so that it is in the shape of a trapezoid, with the top and bottom sides being parallel. The slanted sides help the sections that are cut to stick together into a ribbon rather than sticking to the knife. If the top and bottom sides are parallel, then the ribbon that is cut will be straight. There should be about 2 millimeters of paraffin left around the specimen on all sides.

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Sectioning You must be familiar with the controls on the rotary microtome before you can proceed with sectioning (Figure 12.2). Be sure you have been checked out by an instructor and received permission to go on to the next step. The wheel of the microtome should be in the locked position. Clamp the chuck containing your paraffin block into the chuck holder on the microtome. Using the control screws, align the chuck holder so that the face of the block is absolutely vertical. The block, when you look at it straight on, should look like the picture shown above, with the short side of the trapezoid uppermost. Insert the microtome knife into the knife holder, and tighten the clamping screws. This should always be done with great care. These knives are exceedingly sharp and dangerous. If you should drop one, let it go—never try to catch it. (You could lose a finger by attempting such a catch.) These knives are also very expensive, so it is worth taking care of them. Advance the knife toward the block. This is done by unlocking the wheel, lowering the paraffin block until it is on a level with the knife-edge, and then carefully moving the knife carriage forward until the knife is just in front of the paraffin block. Lock the knife carriage in place. Before cutting sections, check the thickness setting. Paraffin sections are typically cut at between 4 and 12 µm in thickness. For routine sectioning, set the thickness to between 8 and 10 µm. To section, turn the handle of the microtome, which advances the block toward the knife. On each rotation of the wheel, the block is moved forward the number of microns that have been set for sectioning thickness. Turn the wheel in a smooth slow motion to ensure getting the best sections. When sections begin to come off the block, the first several may be incomplete. These can be brushed away with a camel’s-hair paintbrush—no other instrument should ever be used, since these would damage the knife-edge. And never allow your fingers to get anywhere near the knife-edge.

Figure 12.2 A rotary microtome used for cutting paraffin sections.

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Once a ribbon of sections begins to form, the end of the ribbon can be lifted with a moistened camel’s-hair paintbrush. Do not pull on the ribbon; simply support its end with the brush. When you have a suitable length of ribbon (no longer than the length of your ribbon box), take a second moistened camel’s-hair paintbrush, and use it to lift the attached end of the ribbon away from the knife-edge. Place the ribbon carefully into your ribbon box (a flat stationery box does fine). Cover the box so that air currents don’t disturb your ribbon. Keep sectioning until you have all the sections you need. If you are making a set of serial sections, keep each ribbon you cut in order, laying them down so that the last section of each ribbon is on the right, and each successive ribbon is below the previous ribbon. Retrimming the block As you section, you may need to retrim the paraffin block if the block face becomes too large or if the ribbon is not straight. Remember, the ribbon will not be straight unless the top and bottom sides of the block are precisely parallel to one another. When retrimming the block, the knife must first be removed from the knife carriage. Do this without changing the position of the carriage. Retrimming with a safety razor blade may now be done on the paraffin block in place without removing the chuck from the chuck holder. After retrimming the block, replace the knife, and back the paraffin block off a slight distance (an eighth of a turn) from the knife using the handle for this—it is on the opposite side from the sectioning wheel. This will avoid taking too thick of a section on the first rotation of the wheel. Cleaning the knife When you have finished sectioning, the knife should be cleaned of any paraffin that is sticking to it. First attach the knife handle to the knife. This will allow you to hold the knife without danger. Place a few drops of xylene or toluene on the edge of the knife, and with a Kimwipe®, wipe very carefully, moving from the flat of the knife off the edge of the knife, as you would to brush crumbs off a table. Never wipe along the edge of the knife or into the edge. Remove the knife handle and place the knife in its holder (sharp edge down).

Mounting Sections No matter how carefully you cut your sections, the cutting process compresses the sections, and part of the mounting procedure is to expand the sections before they adhere to the slide. This can be done by floating the sections on warm water (5–10ºC below the melting point of the paraffin). Some histologists float the sections in a water bath and then pick the expanded sections up on a slide. I find it easier to put water on the slide itself, float the sections on the water, warm the slide on a warming tray, and then pour off the water after the sections are expanded. This second procedure is the one we will use.

Figure 12.3 Ribbons of paraffin sections floating on water on a slide.

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Select the number of slides you think you will need. Even though these have already been factorycleaned, they should be cleaned again by at least rubbing them with a dry Kimwipe®. Then, using a diamond marker, etch into one end of the slide the information that is necessary to identify it (usually, specimen or experiment number and the number of the slide in the series that you are making). An adhesive must now be applied to the slide so that the specimen sections will adhere properly and won’t fall off during further processing. If you are not trying to identify specific proteins in your staining procedure, then the best adhesive to use is Mayer’s albumen. This can be made easily and stored indefinitely. Mayer’s albumen can even be left out at room temperature, but it is better to store it in the refrigerator. Mayer’s albumen adhesive: Equal parts: glycerin egg white (shaken first to break it up and make it slightly frothy)

Add a few crystals of thymol as a preservative. Store in a dropper bottle, in the refrigerator.

For use Add a drop of adhesive to each slide. Using a Kimwipe®, smear the drop across the area of the slide where you will be placing sections. You should see a shiny film of adhesive when you tilt the slide into the light. It is important not to apply too much adhesive, since it will stain during the staining procedure and will cause too much background staining if applied too thickly. Now add a dropper full of boiled distilled water from a dropper bottle. This water should have been recently boiled and cooled. The boiling removes air from the water and helps to prevent air bubbles from forming underneath your sections. You can now place lengths of paraffin sections onto the slide. Using a scalpel or safety razor blade, cut your paraffin ribbons into lengths. These should be no longer than half the length of the coverslip that you’ll be using. Mounting the ribbons Using two camel’s-hair paintbrushes to hold the two ends of each piece of ribbon, transfer pieces of ribbon to the water on the slide. Ribbons of serial sections must be placed in order, one under the other, until the width of the slide is filled (Figure 12.3). It is best if the ribbons are touching one another along their length so that they are rafted together. This makes it easier to manipulate the ribbons later. Place the slide on a slide warming tray that is set at a temperature that is 5–10ºC below the melting point of the paraffin. Usually this is a temperature that feels very warm to the touch but not hot enough to burn you. As the water warms on the slide, the sections will expand, greatly increasing the length of the ribbons. When the ribbons look completely expanded (this takes about 1 minute), remove the slide from the warming tray. Tip one corner of the slide against absorbent paper, such as a Kimwipe®, to wick off the water without disturbing the ribbons. You need to remove only as much water as will pour off. This leaves a film of water behind that allows the repositioning of any ribbons that have become displaced. Using a dissecting needle, reposition the ribbons so that they are arranged as you wish on the slide. Place the slide back on the warming tray for about 10 seconds. This will help flatten any slight ripple you might have caused when repositioning the ribbons. It is extremely important that you do not melt the paraffin, since this will seriously distort your sections. Remove the slide from the tray and place in a slide holder.

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Drying slides The slides must now dry for at least 6–8 hours (or overnight) before staining. If any water remains prior to staining, the sections will not stick properly to the slide. It is best to dry the slides in a slide-drying oven (set on warm) or in a jar that contains desiccant.

Staining During the processing of the slides, the general scheme is that first the paraffin is removed from the sections using xylene or toluene, since the paraffin prevents staining of the tissues. The slides are then processed down to water, since most staining solutions are water-based. This requires a graded series of alcohols to prevent severe convection currents that would damage the tissues. Following staining, the slides are dehydrated again through a graded series of alcohol, cleared in xylene or toluene, and coverslips are applied using a plastic mounting medium that is miscible with the clearing agent. For clearing agents, it is better to use toluene than xylene since it causes less shrinkage. For alcohol, it is better to use ethanol than methanol since it is gentler on tissue and is nontoxic. Setting up the staining series There are a variety of types of staining dishes that can handle a few slides up to many slides at a time. For up to nine slides, Coplin jars (Figure 12.4) are used (place every other slide at an angle so that it is sharing a slot with the slide in front of it on one side and sharing a slot with the slide behind it on its other side). It is best, however, to process no more than five at a time; each slide is slid into a pair of slots and is separated from the next slide by a pair of glass ridges. This ensures that no slide touches another slide. A distinct advantage to Coplin jars is that most have screw tops, which will prevent evaporation of solutions between use. The Coplin jars should be set up and labeled prior to the beginning of staining. For a typical hematoxylin, eosin, and alcian blue staining process, set up the following series of Coplin jars and label both the jars and their tops. Use label tape and always label in pencil since toluene (and xylene) as well as alcohol typically dissolve the ink of other markers. On a jar that is in the “down” series, a downward pointing arrow can be used to indicate this, and on a jar that is in the “up” series, an arrow pointing upward can be used. In the down series, there are two jars of toluene (or xylene) to ensure that all paraffin is removed. On the up series, there are two jars of 100% ETOH to ensure that all water is removed from the sections. These are the two most critical steps in the series for determining success.

Figure 12.4 A Coplin jar used in staining slides. Slides are placed in slots between glass ridges to keep them from touching one another.

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Coplin jars for a hematoxylin, eosin, and alcian blue staining series • • • • • • • •

Toluene (or xylene) I — ↓ Toluene (or xylene) II — ↓ 100% ETOH — ↓ 95% ETOH — ↓ 70% ETOH — ↓ Distilled water — I Alcian blue Distilled water — II

• • • • • • • •

Hematoxylin Tap water 70% ETOH — ↑ Alcoholic eosin 95% ETOH — ↑ 100% ETOH I — ↑ 100% ETOH II — ↑ Toluene — ↑

Using the hematoxylin, eosin, and alcian blue staining series Hematoxylin and eosin (H&E) are the most commonly used general nuclear and cytoplasmic stains. Hematoxylin (a basic dye) stains acidic components, primarily nucleic acids, a dark blue and is therefore used as a nuclear stain. Eosin (an acidic dye) has an affinity for cytoplasmic elements. Its yellowish-red color makes it an ideal counterstain to hematoxylin. Alcian blue stains glycosaminoglycans (e.g., mucus, chondroitin sulfate of cartilage) and is a simple addition to the H&E series that adds useful information. Reasons for the order of things Because alcian blue is used at a very acidic pH, it must be used prior to hematoxylin in the staining series to avoid removing any bound hematoxylin. A tap-water bath is placed after hematoxylin as a “bluing agent.” Tap water is slightly basic, and a basic solution is needed to change the color of the bound hematoxylin from a rusty brownish color to a dark, almost black, blue. Moving up from eosin should be done as quickly as possible to avoid losing too much eosin from the tissues. Processing can be speeded up by gently dipping the slides in and out of the solutions to increase the speed of diffusion. This must be done gently to avoid detaching the sections from the slide. Filtering stains before use A precipitate forms in most stains, requiring that they be filtered before use. Hematoxylin will need to be filtered if it has been sitting for more than a day. Alcian blue should also be filtered but doesn’t need filtering as frequently—once a week should be sufficient. Alcoholic eosin forms little precipitate. Simply check for a precipitate and filter if necessary. (Unbleached coffee filters make inexpensive filters that are sufficient for removing the precipitate from stains.) Remember that toluene and xylene are toxic, and these should only be used where there is good ventilation. Use in a fume hood (or handle them outside!). Staining schedule Toluene (or xylene) I — ↓ Toluene (or xylene) II — ↓ 100% ETOH — ↓ 95% ETOH — ↓ 70% ETOH — ↓ Distilled water — I Alcian blue (filter before use) Distilled water — II Hematoxylin (filter before use) Tap water — several changes 70% ETOH — ↑ Alcoholic eosin

20 min. 20 min. 5 min. 5 min. 5 min. 5 min. 10 min. 5 min. 2 min. 1 min. each 5 min. 5 min.

Histological Techniques 95% ETOH — ↑ 100% ETOH I — ↑ 100% ETOH II — ↑ Toluene (or xylene)* — ↑

1 1 1 2

min. min. min. min.

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dipping dipping dipping and hold

*On transferring your slides to the final toluene (or xylene), if you notice any white clouding of the fluid, this means that there is still water on the slide and in the tissues. You must immediately go back to 100% ethanol. If you don’t, there will be water droplets in your preparation that will make your slide look like salad dressing.

Formulary for staining solutions Alcoholic eosin eosin Y, saturated solution in 95% ETOH (approx. 0.5 gm/100 ml) 95% ethanol Alcian blue alcian blue distilled water glacial acetic acid thymol (to prevent mold)

10 ml 45 ml

1 gm 100 ml 3 ml a crystal

Filter before use.

Hematoxylin There is true art to making hematoxylin solutions. You can buy them already made, or you can make them yourself. For any hematoxylin solution, it is important to know the shelf life of the solution. Some will keep for years (Delafield’s and Erhlich’s), and some last only a month or two (Harris’s). Ehrlich’s hematoxylin hematoxylin ammonium alum (NH 4Al (SO4)2·12H2O) 100% ethanol glycerin distilled water

2 gm 3 gm 100 ml 100 ml 100 ml

Ripen for 6–8 weeks. Add 10 ml glacial acetic acid. Keeps for years.

Mounting Coverslips You are now ready to put a coverslip on your slide using a mounting medium. Since you are going from toluene (or xylene) to this mounting medium, this step should be done in a fume hood. The mounting medium you use should be a neutral mounting medium such as Histoclad. Cheaper mounting media, such as Permount, are acidic. They cause the stain to fade and oxidize over time, making a cloudy ring around the edge of the slide that gradually works its way to the middle. Any slide that needs to be permanent is worth the extra cost of a neutral mounting medium.

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The hardest part of mounting is devising a method to lower the coverslip without introducing air bubbles into your preparation. Once you start the mounting procedure, work with speed. Do not let your sections dry out. They will do so very quickly. The amount of mounting medium you should put on the slide depends on the length of the coverslip you are using. Use an eyedropper to put a pencil’s width of medium down the length of the slide to match the length of the coverslip. Hold one end of the coverslip with forceps, and lower the other end onto the slide, holding it in place with a dissecting needle. Slowly lower the rest of the coverslip. If an air bubble is introduced, it will usually move to the edge as you lower the coverslip. Once the coverslip is in place, very slight pressure on the top of the coverslip using the end of a pencil or dowel (you can use the handle end of your dissecting needle) will push out any bubbles on the edges. This must be done gently to avoid damaging the sections. In most cases, bubbles should just be left where they are. A few bubbles are better than smashed sections. If you have too much mounting medium so that it is oozing out from under the coverslip, don’t touch it. After it dries for a week, excess mountant can be removed with a razor blade (carefully). Allow your slides to dry flat in a dust-free environment (such as a small cardboard box used for stationery or a flat cardboard slide holder). These should be left at room temperature for several days before observing them under the microscope. After a week, once they are thoroughly dry, they can be stored on their sides in regular slide boxes.

Accompanying Materials See Vade Mecum: “Histotechniques.” This chapter of the CD illustrates, with movies and still pictures, how to do each of the procedures described here.

Selected Bibliography Bird, M. 1995. The Town That Died: A Chronicle of the Halifax Disaster. Nimbus Publishing, Ltd., Halifax, N.S. A history of the explosion caused by picric acid loaded on a munition ship that caught fire in Halifax harbor during World War I, the worst human-made explosion before the nuclear explosions of WW II. Boon, M. E. and L. P. Kok, (eds.). 1992. Microwave Cookbook for Microscopists. Coulomb Press, Leyden, Leiden, The Netherlands. This gives many of the details that will be useful in applying microwave techniques to your histology protocol. Gleiberman, A. S., N. G. Fedtsova and M. G. Rosenfeld. 1999. Tissue interactions in the induction of anterior pituitary: role of the ventral diencephalon, mesenchyme and notochord. Dev. Biol. 213: 340–353. This research paper is a fine example of the use of paraffin histological techniques in immunohistochemistry. Gu, J. (ed.). 1997. Analytical Morphology: Theory, Applications and Protocols. Eaton Publishing Co., Natick, MA. A valuable book to the sophisticated microscopist who needs to delve into immunohistochemistry and fluorescence microscopy. It also has an excellent discussion of the use of a microwave oven to speed up the processing of tissues. Humason, G. L. 1979. Animal Tissue Techniques, 4th Ed. W.H. Freeman, San Francisco. This is a classic, and still very useful, handbook on histological techniques. It gives very complete instructions on each step in the processing of tissue, as well as recipes for fixatives and stains, and explains what each is specific for. Login, G. R. and A. M. Dvorak. 1994. Methods of microwave fixation for microscopy. A review of research and clinical applications: 1970–1992. In W. Graumann, Z. Lojda, A. G. E. Pearse and T.

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H. Schiebler (eds.), Prog. Histochem. Cytochem. 27: 1–127. This guides you through steps needed to adjust the microwave protocol for processing tissues to the particular microwave oven you are using. Presnell, J. K., G. L. Humason and M. Schreibman. 1997. Humason’s Animal Tissue Techniques, 5th Ed. Johns Hopkins Univ. Press, Baltimore. This is a welcome reissuing of Humason’s classic volume.

Suppliers A large biological supply house, such as: Fisher Scientific 585 Alpha Dr. Pittsburgh, PA 15238 1-800-766-7000 www.fishersci.com A full line of histological supplies, including reagents, stains, staining dishes, slides, coverslips, embedding media, mounting media, microtomes and microtome supplies, slide warmers, and slide boxes.

Ward’s Natural Science Establishment, Inc. P.O. Box 92912 5100 West Henrietta Road Rochester, NY 14692-9012 1-800-962-2660 www.wardsci.com Ward’s offers an economical line of student microtomes for cutting single sections at a time, not suitable for serial sectioning. They also sell a selection of histological supplies and neutral mounting media.

Glossary Alcian blue: A dye that, when in an acidic solution, stains glycosaminoglycans (e.g., mucus and chondroitin sulfate of cartilage). Bouin’s fluid: A type of fixative that contains picric acid, formalin, and glacial acetic acid. It is particularly good for fixing lipid-rich tissues, and is therefore often used to fix embryonic specimens. Carnoy’s fixative: A fixative that contains glacial acetic acid and ethanol. Though a harsh fixative on tissues, it has the advantage of not leaving any toxic residues behind once it has evaporated. Chuck: In histological procedures, a chuck is the piece of wood, synthetic fiber, or metal on which the paraffin block is mounted. The chuck fits into the chuck holder of the microtome. Clearing: The process in a histological preparation in which tissues become clear and translucent. Common clearing agents include xylene and toluene, which are immiscible with water, and require that the tissues be dehydrated prior to clearing. Glycerin is a clearing agent that is miscible with water, and can be used without dehydrating the specimen.

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Coplin jar: A type of glass jar that is used for staining histological sections mounted on slides. The jar has sets of parallel grooves that allow multiple slides to be stained without their touching one another. Coverslip: A thin transparent cover that is placed over a specimen on a microscope slide. They are usually made of glass and are of the same refractive index as the mounting medium used for making histological slides. Cytological: Cellular. Dehydration: The removal of water. In histological procedures, a specimen is usually dehydrated through a graded series of alcohol prior to embedding or mounting. Dietrich’s FAA: A fixative that contains formalin, alcohol, and acetic acid. Because of the acetic acid, it tends not to overshrink or overharden tissues. Embed: The process of infiltrating a specimen with a substance such as paraffin so that the specimen can be subsequently sectioned for histological examination. Eosin: An acidic dye that has an affinity for cytoplasmic elements. Its yellowish-red color makes it an ideal counter stain to hematoxylin. Fixation: The process of fixing a tissue. Fixation kills the tissue, as well as any bacteria present, and coagulates (cross-links) proteins. Fixative: Anything that fixes a living organism so that it can’t be degraded by bacteria or other microorganisms. Fluorescence microscopy: A type of microscopy in which molecules in the specimen are made to fluoresce—that is, emit light by being activated with short-wavelength illumination such as the ultraviolet light from mercury arc lamps or lasers. The fluorescing molecules can be fluorescent dyes that stain components of cells; some biological molecules, such as chlorophyll, fluoresce themselves. Formaldehyde: A pungent gas, HCHO, used in solution as a fixative. It is sold as a solution (about 40% formaldehyde in water) called formalin. OSHA has determined that formaldehyde is carcinogenic, so care should be taken to avoid fumes and gloves should be worn to avoid contact with the skin. Glycosaminoglycan (GAG): A long, unbranched polysaccharide (formerly called mucopolysaccharide) made up of repeating units of a disaccharide in which one of the sugars is an amino sugar. Complexes with protein to make proteoglycans. Out in the extracellular matrix, they cause extensive swelling of spaces by absorbing water. Examples of GAGs are hyaluronic acid (hyaluronate), chondroitin sulfate, and heparan sulfate. Hematoxylin: A basic dye that is used as a histological stain. It stains acidic components, primarily nucleic acids, a dark blue and is therefore used as a nuclear stain. Cytoplasmic ribosomes are also darkly stained. Histological study: The study of tissues. Immunohistochemistry: Using antibodies, specific to target molecules, complexed to histological stains in order to locate even minute quantities of the target molecules within tissues and cells. Mayer’s albumen adhesive: An adhesive that is used to affix paraffin sections to a glass slide. It is made from egg whites and glycerin. Microtome knife: Any knife that fits on a microtome and is used to cut histological sections of a specimen. Microtome: A machine used for making exceedingly thin sections of a specimen. Different types of microtomes include the sledge microtome and rotary microtome.

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Mounting medium: The medium in which a specimen is mounted for observation under the microscope. A mounting medium can be water soluble, such as glycerin jelly, or immiscible with water, such as a plastic polymer dissolved in a solvent. When a mounting medium dries, it is firm and transparent, and typically has the same refractive index as glass. OSHA: Occupational Safety and Health Administration, a division of the U.S. Department of Labor. Paraffin embedding: Infiltrating a histological specimen with paraffin. A specimen is first dehydrated and cleared, and then immersed in melted paraffin. Once infiltrated with paraffin, the specimen is placed in a mold containing melted paraffin; when the paraffin cools, the paraffin block can be sectioned on a rotary microtome. Picric acid: A yellow crystalline substance that is an excellent fixative of lipid-rich tissues. It is also highly explosive in dry form and therefore must be stored with water covering the crystals. (The destruction of the Halifax harbor during World War I was caused by an explosion of picric acid aboard a munitions ship.) Rotary microtome: A machine used for sectioning embedded specimens into very thin slices. It holds a sharp steel blade and has a wheel that is turned to advance the specimen towards the blade. Serial sections: Histological sections of a specimen that are taken sequentially, and kept in order. Usually every section is mounted in correct order for microscopic observations. Slide holder: A simple apparatus for holding slides. Usually it is made of metal or glass, with slots that allow multiple slides to be loaded into the holder. Slide warming tray: An electrically heated tray, much like a food warming tray. The rheostat allows the tray to be adjusted to an appropriate heat, warm enough to flatten paraffin sections, but not so hot that it melts them. Stained: In histological procedures, this means dyed with histological dyes to contrast various components of cells and tissues. Staining series: In histological procedures, the series of steps that first prepare sections for staining, then stain the sections, then prepare the sections for final mounting of a coverslip over the sections. Washing: In histological procedures, this is the step following fixation in which the fixative is washed out of the tissue.

Histological Techniques Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. Why do you think you need to fix a tissue before processing it for paraffin sectioning?

2. How can you counterbalance the shrinking effects of alcohol in a fixative?

3. After fixing a tissue, what must you do to it prior to embedding it in paraffin?

4. What type of embedding molds can be used for paraffin embedding, and why is it important that they be watertight?

Name: _________________________

Histotechniques - 2

5. When trimming a paraffin block for cutting, what shape should the block face be?

6. What common tool do you use to handle paraffin sections? How do you store paraffin sections before mounting them on slides?

7. Before mounting sections on a slide, what must you do to ensure that the sections won't come off the slide during processing?

8. Why is toluene or xylene the first step in a staining series when processing paraffin sections?

9. Why do you think a graded series of alcohols is used in a staining series? Why not take the slides directly from 100% alcohol to water?

10. What is a routine nuclear stain? What is a routine cytoplasmic stain? What other stains do you know about, and what components do they stain?

13

chapter

Planarian Regeneration

Regeneration, the process of molding tissues into an exact replica of missing parts, is a mystery worthy of many hours of armchair biology, punctuated by periods of clever experimentation. It is a field for adventurers, where amateurs can have the advantage, since their minds are still free of prejudices. When you venture into the field, be wary of simple solutions. Doubt all theories and devise your own. Test theories through experiments, and let your organism serve up the answers. The oddities that appear to defy explanation may be the clues that everyone else threw away in frustration. Pocket these to mull over in quiet hours. And above all, don’t let confusion cloud your admiration for the phenomenon: regeneration is truly astonishing. Remember also that the ultimate prize would be understanding regeneration in enough detail to be able to trick a normally nonregenerating system, such as the human hand, into regenerating parts tragically lost in accidents. Planarian flatworms offer an ideal system for regeneration experiments. Mutilation of an organism is never a game; however, in the planarian, cutting an animal in half is doing no more than the animal does to itself. When reproduction is in order, the animal adheres tightly to the substrate with its posterior end, pulls forward with its anterior end, and literally tears itself in two. The two halves then regenerate their lost half. This process of asexual reproduction, called fission, produces genetically identical individuals—true clones. Many times an experimenter has returned to a control animal after a few days to find the single control is now two. You may see it happening in your own dishes, so watch carefully. Tearing oneself in half may seem a peculiar mode of reproduction, but it is not the only mode available to the planarian. Its sexual form of reproduction, however, may seem no less peculiar than its asexual mode. Planarians are hermaphrodites, containing both male and female reproductive organs within a single individual. Among many planarians, sexual reproduction is rare; it may occur only seasonally, and in some races it doesn’t occur at all. When it does, fertilization is internal, and the paired organisms each inseminate the other. Sperm are introduced into the mate using a copulatory bulb inserted into a genital pore. Sperm travel up the oviducts and are held in storage near the ovaries until the eggs are ovulated. Eggs traveling down the oviducts are fertilized by the stored sperm. Yolk cells are then added around the eggs by specialized regions of the oviduct. A cocoon, or egg capsule, is secreted to surround a group of eggs, and these capsules are extruded through the genital pore and attached to solid substrata. Though the possibility exists for an animal to fertilize its own eggs, this probably rarely, if ever, occurs.

Collecting and Maintaining Planarians Planarians are free-living, freshwater flatworms, classified as triclad turbellarians. The common genera—Planaria, Dugesia, Polycelis, and Phagocata—include many species that have been used for

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regeneration studies. Two in particular, Dugesia dorotocephala and D. tigrina (brown planarian), are large enough to be satisfying for classroom studies and are commonly available from biological supply companies. If you are braving the wilds to collect your own planarians, there are several hunting methods you can try. Planarians are often found in abundance on the undersurface of stones or wood in the swiftly flowing water below dams, in spring-fed streams, along the shores of lakes, and even in marshes. If the water is not frozen, collections can be made even in midwinter. You may find the animals simply by turning over stones and logs and seeing them adhering to the undersides. You can dislodge the animals using a stream of water from a pipette directed into a bucket or using a soft paintbrush to transfer them. Sometimes you can catch a number of planarians by placing small pieces of fresh liver at the edges of flowing water, waiting 15–20 minutes, and then inspecting the pieces. Planarians, attracted to the liver by chemotaxis, should be feeding on its shaded underside. Another method, less delicate, is to bring in masses of submerged vegetation from streams or lakes. The vegetation should be kept covered with water until it starts to decay. If planarians are present, they will collect at the surface of the water and along the sides of the container, where they can be picked off with a soft paintbrush. Planarians in the laboratory should be maintained in very clean conditions. They should be kept in water in enamel pans or glass finger bowls that are kept in darkened conditions. The water cannot be tap water, since the chlorine in treated water is toxic to these sensitive animals. Spring or well water, or water from the collecting site, should be used, and the water should be changed one or two times a week. Each time the water is changed, the dishes should be either changed or rubbed down to remove the accumulated “scuzz” that builds up from the copious mucus these animals secrete. If the animals must be maintained for a long period prior to the experiments, they should be fed once or twice a week. Once a week is sufficient for maintaining the cultures; twice a week is only necessary if you want to increase the stock quickly. The animals can be fed on raw beef liver or egg yolks. Leave the food in the dishes no more than 2–3 hours, then remove it and change the water. This will prevent bacteria from fouling the water. Prior to your regeneration experiments, the animals should be starved for a week. This will empty the gut of food and will avoid bacterial contamination during the recovery period. Animals will not be fed again until regeneration is complete.

Observations of Normal Anatomy and Behavior Use a soft paintbrush to transfer animals from the large enamel pans or finger bowls to a petri dish containing well or spring water. You will be keeping one animal as a control. The others will be used for regeneration experiments. Anatomy Observe the intact animal under the dissecting microscope, and learn its anatomy using Figure 13.1. This knowledge will be invaluable to you when you start analyzing your regenerates. You will be able to discern a great deal more anatomy by making a planarian “squeeze.” Put a ring of petroleum jelly on a coverslip, place the planarian in the center with a drop of water, then press a second coverslip on top. Apply enough pressure to squeeze the animal flat without damaging it. Place this on a slide and observe it under both the dissecting microscope and the low power of your compound microscope. By turning the preparation over, you can observe both the dorsal and ventral surfaces. Make sure you

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Pigment cup Cerebral ganglion Auricle

Ovary

Lateral nerve cord

Oviduct

Sperm duct

Anterior intestine

Pharynx Testis

Mouth Posterior intestine Copulatory bulb Genital atrium Genital pore

Figure 13.1 Schematic diagram of planarian anatomy. Dorsal view.

adjust the light in a variety of ways; certain structures will be seen better with direct light, others with transmitted light. Notice that the dorsal surface is pigmented and more darkly colored than the ventral surface. When doing regeneration experiments, this will be important for keeping track of dorsal and ventral surfaces of a piece you might be grafting. Look for the gut; this should be the most obvious structure, almost filling the body. It lies dorsal to most other organs. In the midline and opening ventrally is the extrusible pharynx. Turn the preparation so that you are looking at the ventral surface. Notice that the

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pharynx is a highly muscular structure and will look quite dense. The pharynx is used to suck food—fish debris, small prey, even other planarians—into the gut. (If a planarian fissions in two and then cannibalizes its other half, does it count as self-destruction?) The food, once it enters the mouth, travels through the gut, which is tripartite (hence the name triclad). The gut, with a single anterior branch, two posterior branches, and numerous subbranches, extends to almost all parts of the body. Food is processed down to the size of small particles in the gut. These particles then are phagocytized by gut epithelial cells for their final digestion. Undigested food in the gut exits by the same door it entered, the mouth; there is no anus. The animal first fills its gut with water and then forces it out by muscular contraction to irrigate the entire gut. Look from the ventral side for the bilobed brain from which extend two lateral nerve cords that have numerous transverse connections. The nervous tissue will be whitish. Attached to the brain anteriorly are the photoreceptors, which are clustered together into two clumps, each shaded by a cup of pigmented cells. These paired eyespots (or eyecups) give the animal its “cute” cross-eyed look. The pigmented cup ensures that the photoreceptors are receiving light only from one side. When you are observing regenerates to determine whether head regeneration has taken place, the presence of these eyespots will be one of your clearest indicators of new head formation. Look at your animal from the ventral side for the presence of reproductive organs. You will be lucky if you see any, not because they are invisible, but because in many planarians these develop only during seasons of sexual reproduction—usually spring and summer—and regress for the rest of the year. If the reproductive organs are present, you should see paired ovaries anteriorly, with oviducts that lead down to the genital atrium. Testes are multiple and appear along the length of the body. They empty into sperm ducts that are connected to a copulatory bulb in the genital atrium. Behavior After you have observed the internal anatomy, release your animal from its glass sandwich, place it in your dish of water, and let it recover. Use your other planarians to observe some of the basic behavior of these animals. Record your observations, for these will be used as a standard against which you will evaluate your regenerates. Look carefully at the animal. How does it move? Does it glide, swim, or crawl? If you flip it over, can it move on its dorsal surface? Can the animal right itself? The animal has a ciliated ventral surface, and this will allow the animal to glide along surfaces, but far more frequently, it uses its longitudinal and circular musculature for movement, crawling along a surface of mucus that it secretes from its ventral surface. Test for various senses in the animal. How does it respond to touch? Try poking and stroking the animal at its anterior and posterior ends. Does the animal respond differently depending on where it is touched? The animal does have tactile receptors, which are concentrated in the head. Your animal should also exhibit chemoreception (as you already know if you caught them on a liver-baited fishing line). The chemoreceptors are concentrated in ciliated grooves located on the auricles. These auricles are the earlike extensions on the head (adding another measure of “cuteness” to the animal). If you place a small drop of liver juice in one part of the dish and create a slight current with a pipette toward the animal, the animal may respond for you, demonstrating its chemoreception. Record its responses.

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You may have already noticed that planarians tend to aggregate in any shaded region of the dish. They are demonstrating negative phototaxis, responding to the light that they sense with their photoreceptors by moving away from it. You can determine the angle of light that each eyespot is able to perceive by shading your dish and shining light from different directions. What advantage is there to the animal of being negatively phototactic? Once you have observed and recorded all the behavioral responses you can think of testing, move on to the regeneration studies.

The Regeneration Process: Patterns and Theories of Regeneration Regeneration experiments must be carefully planned so that they test specific theories. Before you start, it is important that you know something about planarian regeneration, know the models that are thought to explain it, and dabble in creating your own. Study the next section well before you make your first cut. The regeneration process The first step in regeneration is wound closure and wound healing. When you make your cuts, you should be able to observe these processes by watching the cut surface under the dissecting microscope. First the wound will close by muscular contractions of the body wall. This will take about 10 minutes. The epithelium then heals over the wound, actively spreading, which takes an additional 20 minutes. Once the wound has healed, a blastema begins to form. The blastema is an accumulation of undifferentiated cells that eventually will differentiate into the missing parts. The cells of the blastema are called neoblasts. They are embryonic-like cells that are found throughout the body, held in reserve for just this task. When a cut occurs, the neoblasts in the immediate area coalesce to form the blastema. (There is evidence that neoblasts can also migrate in from more distant regions, but they probably do so only in unusual circumstances when local neoblasts have been destroyed.) The cells at the base of the blastema are highly mitotic, while those in the rest of the blastema are not. In animals held at 22–24°C, the blastema usually forms within 1–2 days and should be clearly visible to you by 3–4 days as an unpigmented region. The timing of events is highly dependent upon temperature, and it is slower at lower temperatures. At 22–24°C, differentiation proceeds quickly, and by 4–6 days you should see differentiated structures in the regenerating region; by 2–3 weeks, regeneration should be complete, with regenerates having reestablished normal body proportions. This type of regeneration, in which the lost parts are built anew from a population of undifferentiated cells, is called epimorphosis. This is in contrast to a process called morphallaxis, in which differentiated cells are remodeled into the new parts. Planarians undoubtedly undergo epimorphosis, but this should not blind us to the possibility that morphallaxis may also be contributing. Some research indicates that morphallaxis may even be playing a major role in the process (Chandebois, 1984).

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Pattern formation and positional information It has been known since the 1700s that when a planarian is cut in half, or into even smaller portions, each piece is able to accurately regenerate an entire animal. Even early workers in the field, such as Charles Manor Child and Thomas Hunt Morgan (two names that should be known to any budding developmental biologist), realized that there must be a system of information that specifies what should be regenerated. How is this information encoded? Child and Morgan considered that gradients of substances must be involved, but substantiating this was difficult. Even now, almost a century later, we still don’t have any specific answers. What we do have is a set of terms that define what we’re looking for: pattern formation refers to the process of creating specific patterns and shapes out of tissues, and positional information refers to the information necessary to create these patterns and shapes (see Wolpert, 1978). The models proposed to explain pattern formation and positional information are too many to examine here, so we will concentrate on two models, a diffusion gradient model and the polar coordinate model. These should provide you with ammunition to start inventing other models on your own—it’s open season, and your models can be as good as anyone’s. Diffusion gradient model

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A number of authors have suggested a diffusion gradient model for positional information in regenerating systems (see Wolpert, 1978). Whether or not such a model is applicable to planarian regeneration is something you can test in your experiments. A diffusion gradient model assumes that positional information is established by a gradient of a chemical substance (morphogen). Cells would know their positions by sensing concentration levels of the morphogen, and they would know their polarity (which way is “up”) by sensing the direction of the gradient. Such a model divides the organism into a number of regions, usually along the anteriorposterior axis (represented in Figure 13.2 as A–E; the number of regions was chosen arbitrarily). The regions represent different threshold levels of the morphogen. Cells are thought to respond by forming different structures at different thresholds.

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Figure 13.2 Schematic diagram of a diffusion gradient model of positional information as it would relate to the planarian. Anterior-posterior positional information is represented as a gradient in which different threshold levels along the gradient establish boundaries, dividing the organism into regions labelled A–E. (After Wolpert, 1978 and Gilbert, 1991)

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The model predicts a number of outcomes that you can test. For example, if the animal were cut in half, the model predicts that the level of morphogen at the cut surface of each piece would tell the cells forming the blastema where they were in relation to the whole and their polarity. The anterior half would know to regenerate a tail end, and the posterior half would know to regenerate a head end (see Figure 13.4A). This is fairly straightforward. It is far more challenging to design experiments that “confound” and really test the model. For example, suppose, as shown in Figure 13.3C, only section B were removed and the remaining two pieces reannealed? What would happen? The model predicts that there would be remodeling of tissues posterior to the cut due to a change in gradient levels in these regions. Does this in fact occur? Such an experiment would be a good test of the model.

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Figure 13.3 Schematic diagram of the polar coordinate model of positional information as it would relate to the planarian. The diagram shows the model proposed by French, Bryant, and Bryant (1976) for vertebrate and insect limbs, as it would relate to the planarian. (A) Anterior–posterior positional information is represented as a series of values ranging from A to E. (B) Circumferential positional information is represented as points on a clockface. Each number on the clockface represents a different positional value. (C) To test the model for anterior posterior positional information, section B can be removed and the remaining pieces annealed. The model predicts that regeneration will occur to intercalate a new section B between sections A and C. (D) To test the model for circumferential positional information, a pie-shaped slice containing positions 3 and 4 can be removed. Wound healing brings positions 2 and 5 into juxtaposition. The model predicts that regeneration will occur to intercalate positions 3 and 4 between positions 2 and 5.

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Polar coordinate model The polar coordinate model, proposed by French, Bryant, and Bryant (1976), was first developed to explain patterns of regeneration in vertebrate and insect limbs. It attempts to explain patterns of regeneration in three dimensions. It is a model well worth understanding, and in your studies you can test whether or not it is applicable to planarian regeneration. The polar coordinate model states that cells recognize their position in the whole because of positional information that each cell receives (like a zip code) designating where the cell is in relation to the rest of the organism. Only two systems of information are needed to generate this zip code: one that gives positional information along the anterior-posterior axis, and one that indicates position within the circumference. This information is thought to be coded not in diffusible morphogens, but in the molecules found on the surfaces of cells. The model designates two sets of values that give positional information. The set of values designating position along the anterior-posterior axis is represented by the letters A–E (see Figure 13.3A; the number of letters is arbitrary); the other set of values, designating position in the circumference, is represented as numbers on a clockface (Figure 13.3B; again, the number is arbitrary, but having 12 creates a clever metaphor). The zip code of a cell is its own combination of a letter and number. The model postulates that each cell knows which values it should be sandwiched between. As long as a cell sees the appropriate values surrounding it, it remains happy. If those values are inappropriate, however, regeneration will occur until a correct sequence is reestablished. For example, if a section corresponding to B were removed (Figure 13.3C), allowing sections A and C to be juxtaposed, regeneration would take place and would intercalate (insert) a new section B between A and C. If a slice were taken out of the circumference that, in the clockface metaphor, brought numbers 2 and 5 into juxtaposition (Figure 13.3D), regeneration would occur to intercalate the missing values 3 and 4 between the 2 and 5. Examples of regeneration in vertebrate and insect limbs have shown that intercalation does occur in these systems, and various manipulations have resulted in often bizarre patterns of regeneration. To explain these patterns, two rules of intercalation have been added to the model. Rule one Intercalation will always proceed by the shortest route to restore positional continuity. If the numbers 2 and 5 are juxtaposed, for example, 3 and 4 will be intercalated rather than 1, 12, 11, 10, 9, 8, 7, 6, 5, and 4—which also would restore continuity but would be the long way around. Rule two Regeneration will proceed by restoring proximal structures first, and then progressively more distal structures—that is, it will first complete the circle of positional values at the cut surface and then regenerate more distal structures. Designing experiments to test the polar coordinate model of positional information can be great fun. For example, you can remove a piece of the planarian, rotate the piece, and then replace it (see Figure 13.5A). This provides a myriad of possibilities for confusing polarities. If you rotate it such that the ventral side is now dorsal, you will have inverted the “clockface.” The cells of the graft now have circumferential positional values that don’t match those of its graft site. The model predicts that there should be regeneration along the edges of the graft, even though technically you haven’t removed any tissue from the animal. If the model is correct, you should get duplication of structures in the region of the graft. You can convince yourself of this by diagramming the regenerate, showing clockface values for the graft and the area where the graft is placed. Then, using rule one above, insert the series of numbers that would restore positional continuity. Imagine what this would represent in terms of actual body parts.

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Patterns of regeneration A number of bizarre patterns of regeneration emerge when students are given free reign to cut as they wish. Some of these oddities can be explained very well using the models presented. For example, if a T-cut is made so that the head is removed and the stump is cut longitudinally part way down its length (see Figure 13.4C), two heads regenerate. The polar coordinate model predicts that this would occur, since, on each side of the cut, the region would sense the loss of half its “clockface,” and each half would intercalate the missing values until two complete heads had been formed.

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Figure 13.4 Patterns of regeneration in the planarian. (A) When the planarian is cut in half, the anterior half regenerates a tail end, and the posterior half regenerates a head end. (B) A thin slice taken from the anterior region can regenerate two heads, a pattern called “Janus-heads.” A thin slice taken from the posterior region can regenerate two tails, called “Janus-tails.” (C) When a T-cut is made where the head is removed and the stump is split part way down its length, two heads regenerate. Shaded areas represent the original piece; unshaded areas represent regenerated portions.

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Some of the patterns of regeneration, however, are difficult to explain. If, for example, a short slice is removed from the planarian, the slice may not regenerate into a complete worm. If it is taken from the anterior region and the slice is wider than it is long, it can regenerate a head anteriorly and another head posteriorly (Figure 13.4B). Such a two-headed regenerate has been called “Janus-heads” (an allusion to the Roman god Janus, who guarded gateways by having two faces, one looking forward and another looking back). A narrow slice taken from the posterior region can regenerate two tails, referred to as “Janus-tails” (Figure 13.4B). Can one of the models presented explain these patterns of regeneration? A typical explanation using the diffusion gradient model has been that if a slice is too narrow, the difference in concentration of the morphogen at the anterior and posterior surfaces is too small to be detectable, so the polarity of the piece cannot be established. Do you believe this? Can you come up with a different or better explanation? Mead and Krump (1986) have done some important work establishing that it is the ratio of length to width in a slice taken from a planarian that determines whether it will regenerate normally, and not the absolute width of the slice. As the ratio of length to width drops below 1.0, abnormal regenerates increase; at a ratio of 0.8, as many as 90% of the regenerates are abnormal. How might you fit this information into one of the models discussed? Can you use it to suggest a different model? Further experiments only thicken the plot. If a narrow slice is removed, but there is a delay between the time that the first and second cuts are made (see Figure 13.5C), different patterns of regeneration result. If the second cut is made after a delay of two or more hours, regeneration can be completely normal (Child, reviewed in Brøndsted, 1969). If the delay is short, only 5–12 minutes, regeneration is abnormal, and the percentage of abnormal regenerates is actually higher than when the second cut is made almost immediately (in less than 1.5 minutes after the first) (Mead, 1991). Time obviously is an important parameter in establishing positional information. Can you now modify one of our positional information models to explain this, or devise your own explanation? Your ideas will be as good as anyone’s. It is frustrating that we still do not know what the actual substances are that bestow positional information. Several suggestions have been made for anterior-posterior gradients that might establish positional information (metabolic gradients; inhibitors and inducers secreted by the nervous system; electric fields; and vitamin A, which acts as a morphogen in some other systems). You could test for these in your experiments. One exciting development is the discovery of homeobox genes in planarians (Garcia-Fernández, Baguñá, and Saló, 1991). This family of genes is known to be involved in pattern formation and appears to be doing the same in the planarian. Two of the planarian homeobox genes (Plox-4 and Plox-5) have been shown to be expressed in an increasing gradient along the anteriorposterior axis of both the intact animal and the regenerate blastema (Orii et al., 1999). The presence of these homeobox genes in planarians gives us products to be looking for, and suggests genetic manipulations that might crack open some answers.

Experimental Procedures Planning your experiments A major task will be to think of clever experiments that test the diffusion gradient or polar coordinate models, or models of your own. A few suggestions are diagrammed in Figure 13.5; they represent only a small part of the smorgasbord of possibilities. You can come up with many, more inventive ones on your own. All of your work will be important to understanding positional information.

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Figure 13.5 Suggestions of experiments that will test models of regeneration. (A) A piece removed from the planarian and replaced in different orientations will juxtapose positional values that don’t match. A number of positions can be tried, including flipping the graft so that its ventral side is aligned with the dorsal side of the animal. (B) Different halves of animals can be aligned so that anterior–posterior and circumferential axes are manipulated either simultaneously or independently of one another. (C) The parameter of time can be investigated by altering the time at which a second cut is made when removing a slice from the animal. Shaded areas represent dorsal surfaces of the organism, unshaded areas are ventral surfaces.

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You should design your experiments in advance. These should be written in your laboratory notebook, with each experimental plan on a separate sheet, since you will be adding your observations for each experiment to these sheets. You should have a standard format for each experiment in which you: 1. On an outline diagram of a planarian, show exactly what manipulations you plan to do. 2. State what model you are testing and in what ways your experiment tests this model. 3. Describe, with diagrams, what results the model predicts. Procedures You have already observed that your animals are energetically motile, and this will be a disadvantage when you start to make your cuts. The animal can be slowed significantly by cooling it. One method is to transfer the animal to a glass slide that is sitting on ice. Since you will have to do all your cuts under a dissecting microscope, realize that the light will be heating up your specimen, counteracting the effects of the ice, so keep the light at a distance if possible. Cuts can be made with microknives or two microneedles pulled across one another in a scissor action. If you use a single microknife to cut, you will find it convenient to have a needle or probe in the other hand to steady the animal during the cutting. A thin slice of a specified width can be made by using two razor blades. Tape the two blades together with a spacer such as a small piece of cardboard positioned between the two blades to give the desired width for the cut. This device also makes the two cuts simultaneously rather than sequentially, which is an important element in the experiment. A hole of a specified diameter can be punched out of the organism by using a Pasteur pipette. Break off the bill of the pipette at the level that will give the bore size you want. The unpolished end of the pipette is an excellent cutting device and can be used as you would use a cork borer. When the animal is stretched out, quickly make your cut. If you need to manipulate or transfer pieces, use microneedles or a soft paintbrush. You may find that the cuts and manipulations you make are not the ones you had initially planned. Don’t despair. Simply change your experimental design accordingly. If you are doing a grafting experiment, the trick is to align the pieces accurately and then to keep the regenerate relatively still during wound healing so that the graft anneals in place. You undoubtedly will invent your own solutions, but here are some tips. Pieces can be aligned on a piece of wet filter paper kept moist from below by cotton soaked in ice-cold spring water. In general, a graft that is placed in a hole constructed for it will stay in place more readily than two pieces placed side-by-side. Also, keeping the regenerate cold during wound healing is important for keeping the pieces aligned: a cold room set at 10ºC is ideal for this, or you can put the regenerate in the refrigerator for up to 24 hours after the operation. Whatever cuts you make, record their exact positions using diagrams, and save all your pieces. Each piece can give you information about regeneration. You will be keeping your regenerates over a period of time. Since regenerates move and climb over partitions in a dish, each regenerate should be put into a separate small petri dish. Be sure to label the dishes with your name and their experiment number. The water in your cultures must be changed frequently, every other day or so. This is a nuisance, since it’s difficult not to throw out the regenerate with the bathwater. Using a pipette to remove the old water is the safest method. Also, remember that chlorinated tap water is toxic. Use only well water or spring water for your cultures.

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Record keeping and analysis Each time you change the water you should record the progress of your regenerates. As you observe them, recall that if the regenerates are kept at room temperature, you can expect to see a blastema within a day or so, differentiated tissues in 4–6 days, and a complete, proportional regenerate in 2–3 weeks. Sometimes it is difficult to determine what you are seeing. Adjust your light for transmitted and then reflected light. Different structures may reveal themselves under different lighting. Also, remember that the blastema will be unpigmented, and that the presence of pigmented eye cups is a sure sign of head regeneration. Keep careful records of the progress of your regenerates, using diagrams to illustrate what you observe. These should be drawn to scale, since size is data. Record approximately how many times larger you have drawn each regenerate. Make a checklist of behavioral responses that you are monitoring, such as responses to touch and light; speed and type of movement; direction of movement; and the ability of a regenerate to right itself. Test for these each time you make a set of observations. Compare these responses to that of your control animal. Do not feed cultures during regeneration. You are not being cruel; they will not be capable of feeding until regeneration is complete, and food in the water will simply rot and contaminate your cultures. If any of your tissues die during the experiment, remove them so that they don’t become a source of contamination. When you start analyzing your data, present it in chart and graph form wherever possible. This will simplify making comparisons among different experiments and will allow you to see trends and the timing of events more easily. Realize that in the best of all possible worlds you would have repeated experiments a number of times and would have been able to report the frequency with which a certain pattern of regeneration occurred. (Frequency is important data.) Determine whether your results are similar to those predicted by the model you are testing. If they are different, try to reformulate the model so that it conforms to your results or create your own model. This is no simple chore, but it is true science. In your excitement and frustration, keep in mind that it is this method of investigation—designing models and then testing them—that fuels advancing science at explosive speeds. Your fuel may be the high-octane we’ve been looking for.

Accompanying Materials See Vade Mecum: “Flatworm.” This chapter of the CD illustrates the anatomy and behavior of the flatworm as well as various regeneration patterns in labeled photographs and movies. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. Contains concise explanations of diffusion gradient models and the polar coordinate model as they relate to regeneration.

Selected Bibliography Baguñá, J. 1998. Planarians. In Cellular and Molecular Basis of Regeneration: From Invertebrates to Humans, P. Ferretti and J. Géraudie (eds.). John Wiley and Sons, Ltd., New York, pp. 135–165. A superb and comprehensive review of what is presently known about planarian regeneration. Baguñá, J., E. Saló, J. Collett, C. Auladell and M. Ribas. 1988. Cellular, molecular, and genetic approaches to regeneration and pattern formation in planarians. Fortschr. Zool. 36: 65–78. A

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concise, well-written review of planarian regeneration studies, dealing less with theoretical models than with descriptive analysis of the process. Brøndsted, H. V. 1969. Planarian Regeneration. Pergamon Press, Oxford. This is a classic. Its almost encyclopedic record of previous experiments is invaluable. If in riffling through its pages you find the same experiments you are doing, do not be discouraged. All good experiments are worth repeating. Chandebois, R. 1984. Intercalary regeneration and level interactions in the freshwater planarian, Dugesia lugubris. II. Evidence for independent antero-posterior and medio-lateral self-regulating systems. Roux Arch. Dev. Biol. 194: 390–396. This is the second in a series of three papers in the same journal. The inventiveness of the experiments to test models of regeneration make the papers well worth examining. French, V., P. J. Bryant and S. V. Bryant. 1976. Pattern regulation in epimorphic fields. Science 193: 969–981. The authors of the polar coordinate model excited the world of regeneration studies with this presentation of their model. Though very readable, it can be found in an even more palatable style in a later article, “Biological regeneration and pattern formation,” in the July 1977 issue of Scientific American. Galtsoff, P. S., F. E. Lutz, P. S. Welch and J. G. Needham (eds.). 1959. Culture Methods for Invertebrate Animals. Dover Publishing, New York. This is a republication of a work originally published in 1937 and is still worthy of another printing. It is an excellent source of the nitty-gritty details that are needed for culturing animals in the laboratory. Garcia-Fernández, J., J. Baguñá and E. Saló. 1991. Planarian homeobox genes: Cloning, sequence analysis, and expression. Proc. Natl. Acad. Sci. USA 88: 7338–7342. A truly exciting paper, this marks the first genetic study to find homeobox genes in the planaria. Gilbert, S. F. 1991. Developmental Biology, 3rd Ed. Sinauer Associates, Sunderland, MA. An earlier edition of the text currently in use. It includes a diagram on page 615 that is useful in studying the gradient diffusion model of positional information. Goss, R. J. 1969. Principles of Regeneration. Academic Press, New York. This is a delightfully written book. It is ageless and well worth reading with an excellent chapter on planarians. Gremigni, V. 1988. Planarian regeneration: An overview of some cellular mechanisms. Zool. Sci. 5: 1153–1163. A concise review, this concentrates primarily on blastema formation and the origin of blastema cells. Kato, K., H. Orii, K. Watanabe and K. Agata. 1999. The role of dorso-ventral interaction in the onset of planarian regneration. Development 126: 1031–1040. An intriguing paper that suggests that the trigger to regeneration in planarians is the juxtaposition of dorsal and ventral epithelia during wound healing. Malacinski, G. M. and S. V. Bryant (eds.). 1984. Pattern Formation: A Primer in Developmental Biology. Macmillan, New York. An important collection of papers on regeneration. Though none is specifically on planarian regeneration, a number of models are discussed, including the polar coordinate model. Mead, R. W. 1991. Effect of timing of cutting on patterning and proportion regulation during regeneration of the planarian Dugesia tigrina. In Turbellarian Biology, S. Tyler (ed.). Kluwer Academic Publishers, Dordrecht, Netherlands, pp. 25–30. This is one of the landmark papers in planarian regeneration that will help to fine-tune investigations. By showing that small increments of time are important factors in pattern formation, it will allow geneticists and biochemists to focus their studies within specific time frames.

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Mead, R. W. and J. Christman. 1998. Proportional regulation in the planarian Dugesia tigrina following regeneration of structures. Hydrobiologia 383: 105–109. A study that looks at the “normalizing” of proportion in regenerates after structures have reformed. Mead, R. W. and M. A. Krump. 1986. Abnormal regeneration in the planarian Dugesia tigrina as a function of the length:width ratio of the regenerating fragment. J. Exp. Zool. 239: 355–364. Another paper from Robert Mead’s laboratory; this one focuses attention on the fine parameters of regeneration, showing that the ratio of dimensions of a piece of planarian rather than absolute dimensions determines patterns of regeneration. This is an extremely important clue for figuring out pattern formation and must be fit into any model of positional information. Morgan, T. H. 1898. Experimental studies of the regeneration in Planaria maculata. Arch. Entwicklungsmech. Org. (Wilhelm Roux) 7: 364–397. For historical reasons alone this should be fascinating to the student of regeneration. The first paper by Thomas Hunt Morgan on his planarian research, it catalogues a number of his experiments and shows the style in which science was reported in these early days. Orii, H., K. Kato, Y. Umesono, T. Sakurai, K. Agata and K. Watanabe. 1999. The planarian HOM/HOX homeoboxgenes (Plox) expressed along the anteroposterior axis. Dev. Biol. 210: 456–468. This excellent paper gives evidence that homeobox genes help establish anterior-posterior patterning in planarians. It includes a useful chart comparing homeobox genes of a number of organisms. Rieger, R. M., S. Tyler, J. P. S. Smith III and G. R. Rieger. 1991. Platyhelminthes: Turbellaria. In Microscopic Anatomy of Invertebrates, Vol. 3: Platyhelminthes and Nemertinea, F. W. Harrison and B. J. Bogitsch (eds.). Wiley-Liss, New York, pp. 7–140. This is an impressive, well-written chapter on turbellarian biology. You will not find anywhere else such a complete, up-to-date review of all that is known at the light-microscopic and ultrastructural levels about the anatomy of these animals. Winston, J. E. (ed.). 1999. Libbie Henrietta Hyman: Life and Contributions. American Museum Novitates, Number 3277. American Museum of Natural History, New York. This entire issue of Novitates is devoted to articles on Libbie Hyman, one of the giants in the field of invertebrate zoology who devoted much of her life to the study of flatworms. Wolff, E. 1961. Recent researches on the regeneration of planaria. In Regeneration, D. Rudnick (ed.). Ronald Press, New York, pp. 53–84. This is an important chapter summarizing the model proposed by Etienne Wolff that gradients of inducer and inhibitor substances are the basis of positional information in planarians. Wolpert, L. 1978. Pattern formation in biological development. Sci. Am. 239(4): 154–164. This is a beautifully illustrated article, written by the author of the term, “positional information.” It provides an excellent, clear presentation of various models of regeneration.

Suppliers Dugesia dorotocephala and D. tigrina can be obtained from: Connecticut Valley Biological Supply Co., Inc. P.O. Box 326 82 Valley Road Southampton, MA 01073 1-800-628-7748

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Carolina Biological Supply Co. 2700 York Road Burlington, NC 27215 1-800-334-5551 www.carolina.com

Glossary Auricle: In the planarian, ear-like extensions on the head in which are found ciliated grooves where chemoreceptors are concentrated. Blastema: In regenerating organisms, the accumulated group of undifferentiated cells that form at the point of an amputation. The blastema has regenerative capabilities and will eventually differentiate into the missing parts. Chemoreceptor: A receptor that can sense the presence of a chemical. Chemotaxis: Attraction by a chemical that causes migration up a concentration gradient of the chemical. Copulatory bulb: In a planarian, a muscular organ used to introduce sperm into the female reproductive tract of another planarian. Diffusion gradient model: A model for positional information in regenerating systems which assumes that positional information is established by a gradient of a chemical substance (a morphogen). Cells are thought to gain different positional information depending upon where they were in the concentration gradient. Epimorphosis: A type of regeneration in which the lost parts are built anew from a population of undifferentiated cells. Planarians undergo epimorphosis. Epithelium: A major type of tissue in an organism, found lining body cavities and covering the outside of the body. The epidermis is an example of a specific type of epithelium. Eye cups: In the planarian, the photosensitive structures in the head region. These exist as a cluster of photoreceptors shaped by a cup of pigmented cells. The pigmented cup ensures that the photoreceptors are receiving light only from one side. Fertilization: Fusion of egg and sperm. Fission: A form of asexual reproduction in which a cell or organism splits in two. Planarians reproduce preferentially by fission. Genital atrium: In the planarian, the region of the reproductive tract that receives products from both the oviducts and sperm ducts. The copulatory bulb protrudes from the genital atrium at the time of copulation. Genital pore: In a planarian, the pore through which gametes are released, both sperm and fertilized eggs. Hermaphrodite: Having both male and female reproductive structures in a single individual. Simultaneous hermaphroditism, as found in the planarian, involves both male and female structures being present at the same time. Sequential hermaphroditism, as seen in certain fishes, involves the organism switching sexes during its life cycle. Homeobox gene: A family of genes known to be involved in pattern formation. Each contains a homeobox region, a conserved 180 base-pair motif, and each codes for a transcription factor. A mutation in a homeobox gene results in the disruption of pattern formation. Homeobox genes are homeotic genes.

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Homeotic gene: A gene whose expression determines the specification of a body region. In Drosophila, the expression of the homeotic gene Antennapedia, for example, specifies the identity of the second thoracic segment. Intercalate: To insert. Janus-heads: A pattern of regeneration found in the planarian in which two heads are produced joined back-to-back. This pattern is observed when a narrow slice is removed from the anterior region of the worm. The thin slice regenerates a head anteriorly and a second head posteriorly. The term “Janus-heads” is an allusion to the Roman god Janus who guarded gateways by having two faces, one looking forward and one looking back. Janus-tails: A pattern of regeneration found in the planarian in which two tails are produced joined together with opposite polarity. This pattern is observed when a narrow slice is removed from the posterior region of the worm. The thin slice regenerates a tail posteriorly and a second tail anteriorly. The term “Janus-tails,” though an inappropriate allusion to the Roman god Janus, is used because of its similarity to the Janus-heads pattern of regeneration. Lateral nerve cords: In a planarian, the paired nerve cords that run from the anterior to the posterior end of the animal and lie along the ventrolateral sides of the organism. There are numerous cross branches connecting the two cords. Microknife: A knife with a small blade, suitable for microdissection under a dissecting microscope. It can be made inexpensively from chips of razor blade, a wooden dowel, and superglue. Microneedle: A needle that is very thin, suitable for microdissection under a dissecting microscope. It can be made inexpensively from inset pins, a wooden dowel, and superglue. Mitotic: Dividing by mitosis. Morphallaxis: A type of regeneration in which differentiated cells are remodeled into the new parts. Some research indicates that morphallaxis may play a role in planarian regeneration. Morphogen: A diffusible molecule, secreted at a distance from its target cells, that specifies the fate of the target cells. It may specify more than one cell type through a concentration gradient of the morphogen, in which case different levels of concentration specify different fates. Mucus: A slimy secretion containing mucins, glycoproteins which upon hydration form a viscous, jelly-like substance. Negative phototaxis: Directed movement away from light. Neoblasts: Found in certain organisms that are capable of regeneration, a population of undifferentiated cells that are kept in reserve until the animal is cut. They are then used to form the regenerative blastema. Planaria have neoblasts. Ovary: The female gonad in which germ cells form ova. Oviduct: The duct in a female that carries eggs from the ovary. Ovulation: The release of an egg (ovum) from the ovary. Pasteur pipette: A pipette with a long narrow bill with a relatively small bore. It can easily be modified into a pipette with smaller bore by heating the bill, pulling it out when the glass is soft, and breaking it off where the bore is the desired size. Pattern formation: The process of creating specific patterns and shapes out of tissues during development and regeneration. Phagocytize: A form of cellular eating in which particles of food or cellular debris are taken into a cell by way of endocytotic vesicles. Pharynx: In a planarian, the region of the gut just behind the mouth, which lies near the middle of the ventral surface of the animal. The pharynx is muscular and extrusible, and can be protruded through the mouth to suck in food.

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Photoreceptors: Light-sensitive receptors. Phototaxis: Directed movement toward light. Planarians: A group of free-living triclad turbellarians found in freshwater lakes, ponds, and streams. Common genera include Planaria, Dugesia, Polycelis and Phagocata. Polar coordinate model: A model for positional information in regenerating systems that attempts to explain patterns of regeneration in three dimensions. The model designates two sets of values that give positional information, one along the long axis, and the other around the circumference. Each cell is thought to know which values it should be sandwiched between. If the values it sees are inappropriate, then regeneration will take place to intercalate the missing values. Positional information: The information necessary to create specific patterns and shapes in tissues during development and regeneration. Regeneration: The process of molding tissues into an exact replica of missing parts. Certain organisms are superb at regeneration, for example planarian flatworms, hydra, sea stars, and salamanders, to name a few examples. Sperm: The male gamete. Tactile receptors: Receptors that respond to touch. In the planarian, these are concentrated in the head region. Testis: The male gonad in which germ cells form sperm. Triclad turbellarians: A group of free-living flatworms in which the gut has three branches, a single anterior and two posterior branches. A planarian is a triclad turbellarian. Tripartite: Having three parts. The gut of a triclad turbellarian, such as the planarian, is tripartite, having three branches, a single anterior and two posterior branches. Turbellarians: The class of primarily free-living flatworms within the phylum Platyhelminthes, as distinct from the parasitic groups in this phylum, the tapeworms and flukes.

Flatworm Regeneration Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. What anatomical structures can you observe in the living planarian?

2. What sensory structures does a planarian have at its anterior end?

3. What reproductive structures are found in a single individual planarian? Your answer defines the term "hermaphroditic."

4. What does a planarian use to move?

Name: _______________________

Flatworm - 2

5. What behaviors did you observe in the planarian? What adaptive value do you think each of these might have?

6. How does a planarian feed? What organ is used for ingesting food, and where is this organ?

7. What mechanisms of reproduction do planaria use?

8. If a planarian is cut near the pharynx, what often happens to the pharynx?

9. What is a blastema, and how long does it take to form in the planarian?

10. What type of cuts can result in the regeneration of two heads on a single planarian body?

14

chapter

Amphibian Development

Breeding In areas where snow blankets the ground for much of the winter, one of the surest signs of spring is a chorus of frogs cracking the stillness of evening with their din or, more subtly, the appearance of spermatophores left on the leafy bottom of a temporary pool by an amorous salamander. It is the male amphibians that respond first to the new pulse of spring. Male frogs and toads (anurans, meaning “no tail”) make all the racket at the breeding pools belting out their “song.” Male salamanders (urodeles, meaning “tailed”), lacking vocal cords, quietly lure their mates with a complex and intimate dance. The females arrive at the breeding pools more sedately. Attracted by the male’s song or dance, they choose their mate. Mating differs according to the mode of fertilization. Fertilization in most anurans is external, and to ensure a successful mating, the sperm must be deposited on the eggs as they are laid. Male frogs and toads, therefore, usually grasp the female around the abdomen in an embrace called amplexus. This stimulates the female to extrude her eggs, and as she does, the male releases his sperm over the eggs. This usually is the last the parents concern themselves with their offspring. After mating, most frogs and toads abandon their progeny to the elements. Most salamanders have internal fertilization. For mating to take place, a female must pick up a packet of sperm, the spermatophore, which the male lays on the bottom of a pool. The female chooses her partner as he woos her with his courting dance, a series of weaves and caresses; the dance brings the female over the spermatophore, which she clasps with the lips of her cloaca. Once lodged in the cloaca, the tip of the spermatophore releases its sperm, and the sperm find their way to a specialized region of the cloaca called the spermatheca. Here they are stored until the female starts laying her eggs. As the eggs pass by the spermatheca, they are fertilized by the sperm. A female may lay her eggs soon after mating, or she may wait months, depending on the species. Usually the site for laying is a shallow pool or a well protected, moist nest. In those species that have nests, the female often guards her eggs until hatching. Some amphibians return to the same breeding site year after year, and certain species of salamanders are known to return exclusively to the site where they were raised as larvae. This homing behavior makes these amphibians especially sensitive to the destruction of their habitat—a problem that today threatens so many species of plants and animals.

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Environmental Hazards Affecting Amphibian Development For decades, a local pond or woods has provided the developmental biologist with a richly stocked field laboratory for amphibian studies. In recent years, however, this laboratory has become poorly stocked in many regions. Declines in amphibian populations have been reported over wide geographic areas: in the Americas, Britain, Australia, and the Amazon basin. In the United States, extensive declines have been documented for the Northwest and Sierra Mountains; they also are reported for the Northeast and Southeast. The decline appears to be due primarily to the loss of wetlands, increased ultraviolet radiation, and the pollution of the breeding sites that remain. Amphibians are especially vulnerable to these habitat changes during their developmental stages, because in most species the embryos and larvae develop in water, and the eggs and embryos, being surrounded only by permeable jelly coats and a vitelline membrane, are unprotected from pollutants. The individual environmental factors that can adversely affect amphibian development are numerous and together create a complicated interplay of effects. One of the culprits is acidic precipitation. Acid rain and snow have lowered the pH of breeding sites, affecting the pH levels of temporary pools more significantly than those of permanent ponds. It is estimated that roughly 10–27% of temporary ponds in the Northeast have pH levels lower than 5.0. Since temporary pools provide the breeding sites of roughly 30% of all salamander species and 50% of all frog species in North America, their acidity could have a significant effect on populations. Sensitivity to acidity varies widely among different species. Though the embryonic stages of some species show complete mortality at pH levels of 4.0–5.0—a typical range for an acidified pool—others survive well in this range. Acidity is not the only culprit in this story, however; it acts in concert with other factors. Lowered pH levels cause aluminum to be leached out of the substrate, and high levels of aluminum have been found to be very toxic to the early larval stages of amphibians. But the formula for toxicity is not simple. The toxicity of the aluminum is very dependent on the pH of the water, and the combination of aluminum and pH that is lethal to one group can be quite different from the combination that affects another group. Another severe threat to amphibian development is long stretches of dry weather that dry up temporary pools. This quickly kills any embryos and larvae that are left in the pools. Acid rain, once again, can be a contributor to the problem. Acidity that may not be low enough to kill the embryos can still slow larval growth. The extended period required for these larvae to come to metamorphosis becomes critical if it leaves the larvae at the bottom of a dry pool. Sudden temperature changes can also be a threat to the survival of amphibian embryos. Though the embryos usually have a wide range of temperature tolerance (the range for Rana sylvatica embryos, for example, is 5–21°C), a sudden freeze can kill off any embryos that are caught near the water’s surface. More problematic can be the synergistic effects of temperature and acid stress. An embryo developing at the top of its temperature range will succumb if exposed to an acid stress as well. Another hazard to amphibian development, one that has received attention lately with the thinning of the protective upper atmospheric ozone layer, is the increased exposure to ultraviolet radiation. This can be particularly damaging to amphibians because of the long exposure of their embryos to sunlight. In addition, the synergistic effects of ultraviolet radiation and low pH are particularly damaging and can greatly increase mortality. You will be measuring only a few of the environmental factors that could be affecting amphibian development in your local areas, since other factors, such as aluminum levels and synergistic effects,

Amphibian Development

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are probably beyond your monitoring abilities. However, by monitoring pH, temperature, and the drying out of pools, and correlating this with survival rates of the embryos you observe, you should get a general picture of amphibian reproductive success in your area.

Preparing for the Field Trip Learn about your local amphibians first Most amphibians in the Northeast breed in the spring or early summer. Those in the Southeast may breed even in the winter months, coming out in a warm rain. In the more arid West, breeding is often rhymed closely with periods of rain. The eggs—whether hidden under rocks or logs, attached snugly to submerged vegetation, or broadcast haphazardly to float in rafts at the water’s surface—are usually easily found if you know when and where to look. So schedule your trip for a time when several species of amphibians are breeding (this will differ radically, depending on your geographic region. Check a regional guide to amphibians that gives you specifics about your area; some are listed in the bibliography.) Table 14.1 is a short listing that will help. Determine which species you can expect to find in your area and where you might find them. It is always advisable to consult a herpetology enthusiast about their field notes on “who, where, and when.” (These invaluable people quietly exist everywhere—just start asking around. Often they will be willing to come along on your field trip.) Determine a route for your trip that will pass through both woods and fields and will encounter a pond, temporary pools such as ditches, and slow-moving water. Swamps and marshes are wonderful if you can find them. Try to schedule both a daytime and a nighttime field trip. Eggs are most easily found by day, but the night brings forth both mating behavior in salamanders and frogs and the varied choruses of the frogs.

Table 14.1 Breeding information on some commons North American amphibians

Distribution

Season eggs laid

Laying habitat

Acris gryllus Cricket frog

Southern and central U.S.

February– October

Hyla regilla Pacific tree frog

Western U.S. and Canada

Hyla versicolor Gray tree frog

Species

Where eggs are found

Egg clutch size

Egg

Shallow meadow pools and swamps

Attached to stems of grass in shallow water or strewn on bottom

Eggs laid separately, few up to 250, jelly firm

0.9–1.0 mm diam. Brownish above, white below

January– May

Quiet water, pools, ponds, swamps

Attached to vegetation at or below water surface

Eggs laid in small, loose, mass, 10-70 eggs, loose jelly

1.3 mm diam. Brown above, yellowish below

Eastern U.S. and southeastern Canada

May– June

Temporary pools or quiet permanent water

Loosely attached to vegetation at water surface

10–40 eggs, loose jelly

1.1–1.2 mm diam. Very light brown to gray above, white to yellow below

Pseudacris (Hyla) crucifer Spring peeper

Eastern U.S. and southeastern Canada

April– May

Temporary or permanent water

Eggs attached singly to submerged vegetation

Up to 900 eggs, jelly firm

1.1 mm diam. Dark brownblack above, white below

Rana catesbeiana Bullfrog

East of Rockies and West Coast

May– August

Permanent water with vegetation

Thin, floating mass at surface

12,000–20,000 eggs in mass up to 0.6 m across, jelly loose

1.23–1.7 mm diam. Black above, creamy white below

Frogs

(continued)

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Table 14.1 Breeding information on some commons North American amphibians (continued) Species

Distribution

Season eggs laid

Laying habitat

Rana clamitans Green frog

Eastern North America

June– August

Rana palustris Pickerel Frog

Eastern North America

Rana pipiens (species complex) Leopard frog

U.S., Canada, and Mexico, except for West Coast

Where eggs are found

Egg clutch size

Egg

Permanent quiet water with vegetation

In shallow water, floating mass, attached to vegetation

1500–5000 eggs in mass 15–30 cm diam., jelly loose

1.5 mm diam. Black above, white below

April– May

Permanent water with vegetation

In center of water, attached to submerged sticks

2000–3000 eggs in globular mass, 5–10 cm diam. Firm jelly

1.6 diam. Brown above, white to yellow below

March– May (after wood frog) in North

Permanent water with vegetation such as lake inlets, slow streams, ponds, or overflows

Laid in the shallows, often in communal masses

2000–4000 eggs in flattened oval masses, 7.5–15 cm × 5–7.5 cm, jelly loose

1.6 mm diam. Black above, white below

February– December in South Rana sylvatica Wood frog

Northern U.S. and Canada

March– May

Temporary pools

Often in large communal masses, attached to submerged stems or branches

Up to 3000 eggs in globular mass, 6–10 cm diam. Jelly firm

2.0 mm diam. Black or chocolate above, ivory white below

Bufo americanus American toad

Eastern and Midwest U.S. north of Georgia, southeastern Canada and west through Ontario

April– May

Shallow, open pools

Threaded among submerged vegetation or stretched out on bottom

6000 eggs in pairs of long (4 m or longer), spiral strings, eggs in single row down string, jelly firm

1.0–1.4 mm diam. Black above, white below

Bufo boreas Western toad

Northwestern U.S. and Canada

March– September

Ponds and shallow pools

Long strings strewn among grasses usually at margins of pool or in shallow water

Up to 17,000 eggs in long strings, eggs in one or two rows down string, jelly firm

1.5–1.75 mm diam. Black above, white below

Bufo cognatus Great plains toad

Dakotas south through Texas and Southwest to Calif.

April– September

Pools, irrigation ditches, overflow from streams

Long strings in shallow water

Up to 20,000 eggs in long strings, eggs in one or two rows down string, jelly firm

1.18 mm diam. Black above, white below

Bufo woodhousii fowleri Fowler’s toad

Central and eastern U.S.

April– June

Shallow water of pools, lake margins, or ditches

Laid in strings tangled in aquatic vegetation below surface

8000 eggs, in long tangled strings, eggs in one or two rows down string, jelly firm

1.0–1.2 mm diam. Black above, white below

Bufo woodhousii woodhousii Woodhouse’s

Great Plains and western U.S. to Calif.

March– July

Shallow water of pools, lake margins, or ditches

Laid in strings tangled in aquatic vegetation below surface

Up to 25,000 eggs, in long tangled strings, eggs in one or two rows down sting, jelly firm

1.0–1.2 mm diam. Black above, white below

Ambystoma jeffersonianum Jefferson salamander

Eastern U.S. and southern Canada

Early Spring

Temporary or permanent water

Laid singly, attached to one another and to a twig.

100–300 eggs, jelly capsules often green from unicellular algae, jelly thick and soft

2–2.5 mm diam. Dark brownblack above, lighter below

A. laterale Bluespotted salamander

Northeastern U.S. and Canada

April

Shallow (30-40 cm deep or less) temporary pools with vegetation or leafy bottoms

Laid in loose groups that attach lightly to vegetation or fall to bottom

Single or loose groups of 2–6 eggs, large clear jelly capsules

2–2.5 mm diam. Dark brownblack above, lighter below

Toads

Salamanders

(continued)

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Table 14.1 Breeding information on some commons North American amphibians (continued) Season eggs laid

Laying habitat

Where eggs are found

Egg clutch size

Egg

Eastern U.S.

January– May

Temporary pools

Attached to sticks about 15 cm below surface

100–200 eggs in oval mass about 10 cm across, jelly firm, often milky opaque

2.5–3 mm diam. Dark brown or gray above, whitish to yellow

A. opacum Marble salamander

East and Midwest, not north of Boston– Chicago line

September– October

Under moss or log at edge of water or in dry beds of temporary ponds or streams

Laid singly in a depression likely to be flooded by fall rains

50–232 eggs, guarded by female, two separate jelly envelopes easily distinguished

2.7 mm diam. Dull yellow

A. tigrinum Tiger salamander

U.S., Canada, Mexico

December– April

Temporary pools

Attached to twigs about 1 inch below surface

23–110 eggs, globular or oblong mass, 55–70 mm × 75–100 mm, jelly loose

2.5 mm diam. Light to dark brown above, pale cream to buff below

Eurycea bislineata Two-lined salamander

Central and eastern U.S.

April– June

Flowing water of small brook

Cemented individually by stalk to underside of rocks

12–36 eggs, two distinct jelly envelopes, outer one extended into slender jelly stalk

2.5–3 mm diam. White to pale yellow

Hemidactylium scutatum Four-toed salamander

Eastern U.S.

MidApril– June

Cavities, 7–10 cm deep, in moss or grass overhanging water

Laid singly but adhere in cluster

19–50 eggs, not aggregated into common jelly envelope, but cling to one another, not stalked, guarded by female

2.5–3 mm diam. Very light pigment above, yellowish below

Necturus maculosus Mudpuppy

East of Rockies, north of Georgia

Spring and summer

Lakes or streams

Excavations scooped out from underneath large rocks or submerged logs

60–100 eggs, attached singly by stalks to roof of nest, three jelly envelopes, guarded by female

5–6 mm diam. Light yellow

Notopthalmus viridescens Eastern newt or Red-spotted newt

Eastern U.S., southeastern Canada

April– June

Sluggish water

Attached singly to submerged stems, leaves or wrapped individually in leaves

20–30 eggs, separate, not in common mass, elliptical jelly envelopes, outer jelly covered with thin filaments

1.5 mm diam. Light to dark brown above, yellowish green below

Plethodon cinereus Redbacked salamander

Northeastern U.S.

June– July

Cavities in well-decayed logs or stumps

Suspended from roof of nest in small clusters

4–17 eggs, outer jelly 3.0–5.0 mm diam. envelope covered with White above, white mucous stands, very pale yellow converge to form attachto pale yellowment stalk, held together orange below like bunch of grapes and suspended by common pedicel, guarded by females

Taricha granulosus (Triturus granulosus) Rough skin (Oregon) Newt

West Coast, S.E. Alaskacentral Calif.

Late December– July

Streams, lakes, and ponds

Laid singly or small clusters, attached to vegetation and other objects in water

Single or 2–3 eggs, outer jelly envelope 3.63 mm, jelly firm

1.8–2 mm diam. Light brown above, white below

Taricha torosus (Triturus torosus) California newt

California coastal ranges

December– May

Streams, ponds, and reservoirs

Laid in small clumps, attached to sticks, vegetation, and other objects under water

7–29 eggs, globular mass, 15 mm across, jelly firm

1.88–2.5 mm diam. Pale brown above, whitishyellow below

Species

Distribution

Ambystoma maculatum Spotted salamander

Source: Primarily after Hunter, Albright, and Arbuckle, 1992; Rugh, 1948; and Wright and Wright, 1949.

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Equipment needed and recording of data For the field trip, you will need long-handled fish nets for egg masses beyond arm’s reach; clean plastic (because glass breaks) containers with tops; index cards; a pencil (because pen runs); a measuring tape; a sturdy thermometer (a metal pocket thermometer is especially nice); some distilled water (for washing electrodes if necessary and wetting soil samples); and a pH-measuring device. (There are all levels of sophistication of pH meters, and they vary in reliability; see the section on pH measurements. Choose a method that fits your goals and purse.) Whenever you find a clutch of eggs, record information about the eggs on an index card, and place the card along with a container of three or four eggs (more is unwarranted) and water or soil from the site. Your collection of eggs will be brought back to the lab for measurements and observations of development. If there are only very few eggs at the site, you will not be collecting any, but record the information about them. The information you take down is important for making reliable identifications of the species of eggs, for updating records on “who, what, where, and when” amphibians breed in your area, and for evaluating whether and which amphibians in your area are under stress from environmental factors. The information recorded on the card should be as listed below. It is useful to prepare cards in advance that state the information that is needed and blanks for filling in. Working in teams of three or four lightens the load. Personnel and date Who is collecting the data, and the date. Macrohabitat Where were the eggs found (e.g., bog, marsh, woods, field)? If a woods, type of trees. Record air temperature. Mesohabitat Were the eggs found in a lake, stream, pond, temporary pool, under a log, etc.? Microhabitat If eggs are found in water, what is its temperature and pH? How deep is it? Is it still or running? How much vegetation? Is the bottom grassy or leafy? What type of leaves? If eggs were found under a log, what is the temperature and pH of the substrate? Is the log rotten? What type of tree did it come from? Are the eggs laid in moss or leaf detritus? What type of leaves? Where in specific habitat eggs found If eggs are found in water, are they at the water’s edge or in the middle of the pond? Are the eggs floating or submerged? If submerged, how far under the water’s surface? Are they attached? If so, to what? If in a nest, what are its dimensions? Is it near any water? Description of egg mass Are the eggs laid singly or in a mass? What are the dimensions of the egg mass? How many eggs in the mass? (If there are too many eggs to count, you can estimate by counting the number of eggs in a small sample, calculating from their measurements the volumes of the small sample and the entire mass, and figuring from these numbers the total number of eggs.) Is the jelly firm or loose? Is it milky or clear? Is it green (contains green algae)? Description of egg Color, both above and below; diameter; stage of development (see staging series in Table 14.3); percentage of healthy eggs; description of unhealthy eggs. Species of the egg From your information above and that given in Table 14.1, what species do the eggs appear to be? You will be verifying your identification back in the laboratory. Equipment for pH measurements in the field It is always best to make your pH measurements in the field rather than back in the lab. This will give the most accurate readings, since the pH of water changes as it is trundled around and loses CO 2. The best method for accurate pH measurement is to use a portable electric pH meter. The Orion™ meter

Amphibian Development

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with an Orion Ross™ electrode is particularly accurate, but extremely expensive. Far less expensive meters are available (see the Suppliers list at the end of the chapter). Those that both read and adjust for temperature are the best. If you use an electric pH meter, you should test your meter for its sensitivity to ionic effects. The low ionic concentrations of the soft water found in ponds and pools make it difficult for electrode pH meters to give accurate readings. After calibrating your pH meter with a known acidic pH buffer, measure a low ionic acidic solution, also of known pH, to determine if there is any fault in the reading. For a low ionic acidic solution, you can use a 0.0001 N solution of hydrochloric acid (HCl). This will have a pH of 4.0. If the meter can’t read this dilute solution, try another electrode. Be sure the pH buffer and low ionic acidic solutions are at approximately the same temperature. Always rinse the electrode several times with distilled water between buffers and before reading each sample, and make two or three readings of any sample to ensure accuracy. Colorimetric pH measurement kits also exist, and these can be relatively inexpensive and convenient for field work. But a word of caution about their use. They have been found to give inaccurate readings in acidic soft water. Discrepancies are usually in the range of ± 0.2 units of pH, but can be as high as 0.8 pH units. These kits use dyes of different pH that change color as the pH changes. The color change is measured against a color chart to determine the pH of the solution. You will get the most accurate readings by using dyes that have a pH that is at or very near to the pH of the solution being measured. Paper pH test strips are the least expensive method of measuring pH, and those with the range most appropriate for this study are sold as acid rain test kits and soil pH strips. They are the least reliable method of measuring pH, but are usually within a lab’s budget. Come dressed for the field On the day of the field trip, come dressed for the weather, and wear boots (or waders if you have them) and polarized sunglasses. The polarized glass will allow you to see through any glare on the water surface, and you’ll spot eggs that would otherwise be invisible. If the field trip is at night, or in the evening extending into dark, bring a flashlight and red cellophane to cover the light. Amphibians are insensitive to red light, so you’ll be able to observe their behavior without disturbing it. If it is raining, rejoice. Amphibians love wet weather and will be out in great numbers. Know government regulations States and provinces often have regulations regarding amphibians, so check with the appropriate Nongame and/or Endangered Species Program or Department of Wildlife, Natural Resources, or Conservation before embarking on your trip.

The Field Trip As you trudge along your planned route, examine all bodies of water, regardless of size. Often, it will be the tire rut filled with rainwater that some amphibian has chosen as a nursery for her brood. As a rule of thumb (though there are many exceptions), you can expect to find the eggs of frogs and toads in ponds or temporary pools, and those of salamanders in temporary pools or under and inside logs. At first you may have trouble spotting an egg mass. Wear your polarized sunglasses to reduce glare on the water, get down on your knees, and paw around. Egg masses that are submerged, and especially those

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sitting on the bottom of a pool or pond, become covered with sediment. Often all that you see to alert you to their presence is a globular shape. In early spring in the Northeast, you can expect to find the eggs of the wood frog (Rana sylvatica); spring peeper (Pseudacris crucifer previously Hyla crucifer); leopard frog (Rana pipiens); and spotted salamander (Ambystoma maculatum); all of these are found in water. These eggs are easily distinguishable (some are shown in Figure 14.1). The eggs of the spring peeper will appear singly, at the edges of pools, and are very difficult to see. The eggs of spotted salamanders are in globular masses with firm jelly. The jelly is sometimes milky and opaque. The eggs of the wood frog are slightly smaller than those of the spotted salamander and are usually found in larger masses, with jelly that is less firm. The egg masses of the leopard frog usually are huge and black with loose jelly and are the most easily spotted, but they may be rafted out in the middle of the pond where you can only get to if you brought waders. The communal rafting of these eggs is an adaptation that increases survival, since the center eggs retain more heat and are protected from sudden drops in temperature.

(A)

(D)

5 mm 10 mm

(B)

(E) Sperm cap Stalk

3 mm

10 mm (F)

(C)

10 mm 5 mm

Figure 14.1 Egg masses and a spermatophore of some common North American amphibians. (A) Eggs of the mudpuppy, Necturus maculosus, hanging by jelly stalks from the underside of a rock in a lake. (B) A spermatophore of the spotted salamander, Ambystoma maculatum. The gelatinous stalk supports a sperm cap. (C) Egg mass of the Jefferson salamander, Ambystoma jeffersonianum, laid on a twig in a vernal pool. (D) Eggs of the spring peeper, Pseudacris crucifer, laid on grass blades submerged at the edge of a small pond. (E) Eggs of the leopard frog, Rana pipiens, laid in a floating mass on the surface of a pond. (F) String of eggs of the American toad, Bufo americanus, which would be laid as paired strings on submerged vegetation or on the bottom of a shallow pool.

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The normal bicoloration of amphibian eggs is an adaptation as well. The dark upper (animal) hemisphere is difficult to see against the dark water when viewed from above and therefore protects against aerial predators. The light lower (vegetal) hemisphere is difficult to see for any hungry swimmer looking up at the eggs against a light sky. When such bicoloration is not needed for camouflage, such as in eggs that are laid under rocks or logs, the eggs often are unpigmented. For any egg mass that is found, record the necessary information on an index card. Remember, working in teams will reduce the amount of work for each of you. Make your pH and temperature measurements on water from the exact location and depth at which the eggs were found. If there are more than 50 eggs in the mass, or there are a number of similar masses in the vicinity, remove three or four eggs and put these in a plastic container along with the index card and enough water to fill half the container. These will be brought back to the laboratory for observation. If you find eggs under a log, these are likely to be those of a salamander, and the female may be around guarding them, so look carefully for her. In general, these eggs should not be disturbed, since usually there are very few. If there are plenty (over 50), two or three may be collected along with a small soil sample. Measure the temperature of the soil and take a pH reading, moistening the soil first with some distilled water. Remember that we are concerned with possible environmental effects on developmental success, so look carefully at the eggs. It is important to determine how many look healthy and how many appear moribund (abnormally whitish or deteriorating). If there are deformities, what are they, and how many eggs are affected? Don’t forget to look for the spermatophores left by male salamanders (see Figure 14.1). These little cone-shaped, gelatinous structures can be spotted from a moving bicycle, if you know what you’re looking for. When you are going at a speed of 15 miles per hour, they often look like the white triangles of hardened sap seen on pine cones (and I’ve come to a screeching halt many times mistaking the two). They are usually found in clusters of ten or more, often on leaves in the bottom of water-filled ditches bordering woods. If you find some, take one or two back to the lab. If you are lucky, the sperm cap will still be intact, and you can put it on a slide under the microscope to see the living sperm. During your trip, cover as many habitats as possible, and always observe good ecological manners.

Rules for the Road Walk softly and carry a long-poled fishing net. Be inquisitive—turn over rocks and logs, search through wet pond grasses. Always replace the rock or log and leave the habitat as you found it. Never tromp as a large group through a pond or other small body of water. Send out a single scout, so as to leave the habitat as undisturbed as possible. Be patient—pause for long minutes to listen and look. Never overcollect. Observe and put back. If you catch an adult amphibian, wet your hands before holding it, and hold it only for a short period. Do not pass it around. Its skin is fragile and dries out very quickly. Once you have finished your observations on the eggs brought back to the laboratory, always return them to their specific habitat as soon as possible.

Night Sounds If you are lucky enough to schedule a night frog walk, you may be serenaded by a cacophony of songs from males of various species of frogs and toads. Marshes and swamps are among the best places to

Amphibian Development Table 14.2 Songs of some common North American male anurans

Species

Time calling is Air/water heard in various temperature when regions males first start calling Male mating call

Acris gryllus Cricket frog

Late January– October in North Carolina

/Water can have a surface film of ice

Sounds like a rattle or metallic click in rapid succession

Rana sylvatica Wood frog

Late March or early April-late April, early May in central Maine

50 •F/54 •F

Short abrupt “wrunk,” emitted singly or several in rapid succession, sounds like quacking of ducks

Pseudacris crucifer Spring peeper

Early April–May in central Maine, greatest on warm, rainy nights

52 •F/above 50 •F

Simple loud peep ending with an upward slur, repeated 15–25 times/minute, occasionally a short trill

Rana pipiens Leopard frog

Late April, early May in central Maine for 1–4 weeks

55 •F/59 •F

Long drawn out guttural snore up to 3 sec., release call is a short series of trills, and warning is an untrilled scream

Bufo americanus American toad

Mid–April–early July in central Maine

/40 •F

Long musical trill, 6–30 sec.

Rana palustris Pickerel frog

Late April or early May–May in central Maine

50–65 •F /50–65 •F

Low-pitched croak, somewhat like a rusty door opened slowly, or snoring grunt, similar to leopard frog but shorter and softer

Bufo woodhousii fowleri Fowler’s toad Bufo woodhousii woodhousii Woodhouse’s toad

After B. americanus 65 •F /65 •F B.w.f., April in N. Carolina, B.w.w., February in Arizona

Nasal “waaaaah” 1–5 sec. long, repeated 4–8 times per minute

Hyla versicolor Gray tree frog

May–August in central Maine

Above 68 •F/

Short, very loud trill, lasting 0.5–3 seconds, repeated about 10 times in 30 sec.

Rana clamitans Green frog

May–August in central Maine

/70 •F

Single explosive note like the plucking of a loose banjo string, may repeat 3–4 times, each note softer than the last

Rana catesbeiana Bullfrog

Last to breed in Maine, mid- to late June and July

68–72º •F/66º •F

Deep-throated “jug-o-rum”

Source: From information in Hunter, Albright, and Arbuckle, 1992; J. Markowsky, pers. comm.; Rugh, 1948, B. K. Sullivan, pers. comm.; and Wright and Wright, 1949.

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hear such a concert. If possible, find an anuran enthusiast, ask them which anurans call when in your area, and bring them along on your trip. A very useful aid to prepare you for your walk is a narrated CD or cassette tape of night sounds, which you should listen to several times before your walk. (Two such recordings are A Guide to Night Sounds and The Calls of Frogs and Toads, listed in the bibliography. They are inexpensive and easily obtainable.) Also, Table 14.2 lists some common North American frogs in the order that they are heard in the spring. The time that they are heard will vary among different regions. You should be able to extrapolate from the table approximate times when you might hear these frogs in your area. Though air and water temperatures are listed for when the males first start calling, this is very approximate, since precipitation is an equally important factor. In general, calling will start after a period of rain. The chart’s descriptions of the songs, of course, are subjective, and if you compose better ones, feel free to amend the chart.

Back at the Laboratory Species identification Your first job back in the laboratory is to verify your field identification of the species of your eggs. Recheck your identifications against Table 14.1, then use your dissecting microscope to look at each group of eggs you collected. If your eggs are in the early stages of development, you can measure the diameters of the eggs and their surrounding jelly capsules (there should be one or more of these), and use these in your identification. Compare the measurements with those given in Table 14.1 and Figure 14.2. Correct your data card if necessary, and make your collection available for others to see. Normal development Now examine your eggs to observe as many stages of development as possible. Notice that the egg itself is not transparent. This is due to the large amount of yolk stored within, concentrated primarily in the vegetal hemisphere. The egg is classified as a mesolecithal egg, meaning that it contains a moderate amount of yolk. Most eggs are pigmented with melanin, a black pigment, distributed primarily in the animal hemisphere, giving the egg a bicoloration of dark above and light below. The concentration of melanin varies among the species. The less melanin, the lighter brown the egg. For years embryologists have sought out light-colored amphibian eggs to use for fate-map experiments. The light pigmentation allows for experiments using vital dyes in which a small number of cells on the surface are stained and then followed throughout gastrulation. Determine what stages of development your embryos are in, using Table 14.3 and Figures 14.3 and 14.4. If you were doing this for publication, you would use a staging series that is more specific to the group being studied. For our purposes, this simplified series can be used for both anuran and urodele embryos, though it was constructed by Roberts Rugh for the anurans Rana pipiens and Rana sylvaticus. You will be lucky if you have stages as young as cleavage, for this will mean that, unless the water from which you collected the eggs was exceptionally cold (which slows down development considerably), the eggs probably were laid that day or the night before. If you really have hit the jackpot, you will have eggs that have just been fertilized. In these you will see a lightly pigmented gray crescent-shaped region, marking an area directly opposite the point of sperm entry. This gray crescent material is the precursor to the notochordal mesoderm. The first cleavage usually, but not always, bisects this gray crescent. The first, second, and fourth cleavage furrows start at the animal pole and work their way toward the vegetal pole. The third and fifth cleavage furrows are perpendicular to these.

Amphibian Development

Hyla regilla Bufo americanus

Bufo cognatus Bufo woodhousii

Bufo boreas

Hyla versicolor

Rana clamitans Rana catesbeiana

Ambystoma jeffersonianum

Rana palustris

Ambystoma maculatum

Hemidactylium scutatum

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Acris gryllus

Pseudacris crucifer

Rana pipiens Rana sylvatica

Ambystoma opacum

Ambystoma tigrinum

Notopthalmus viridescens

Eurycea bislineata

Plethodon cinereus Taricha granulosus

5 mm Necturus maculosus

Taricha torosus

Figure 14.2 Eggs from some common North American amphibians, showing relative size and the diameter and number of jelly capsules. The central dark structure in each diagram is the egg, surrounded by its jelly capsules. The vitelline membrane, which lies snugly around the egg, is not shown. The relative darkness of the egg represents the amount of pigmentation that is found in the animal hemisphere of the egg. The most lightly shaded eggs are unpigmented. Those more darkly shaded are light-to-dark brown, and the darkest are black. The first two rows of eggs are those of common anurans. The lower rows are eggs of common urodeles. (After Bishop, 1941; Stebbins, 1951; Wright and Wright, 1949.)

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If you have mid-cleavage stages, notice that cells in the vegetal half are larger and less numerous than in the animal half. This is a consequence of yolk being concentrated in the vegetal half. As in any embryo, a large amount of yolk is an impediment to cleavage. In the amphibian egg, this yolk slows down cleavage in the vegetal region, and by mid-cleavage the animal half has pulled ahead in its cleavage rate and ends up with more but smaller cells. If your embryos have passed the cleavage stages, they may be in gastrulation. Look for the presence of a blastopore. In the amphibian, surface cells move toward the blastopore and move inward first at the dorsal lip, then the lateral lips, and finally at the ventral lip of the blastopore, a process called involution. Among amphibians, there is some variation in which cells move over the lips of the blastopore. In the South African clawed frog, Xenopus laevis, used in so many developmental studies, it is the endoderm; but in most North American amphibians, it is the mesoderm. In these, the dorsal lip is where the gray crescent material, or notochordal mesoderm, involutes. Once the entire blastopore is formed, a small plug of yolky endodermal cells is caught between the lips of the blastopore. This yolk plug is eventually internalized. Following the major gastrulation movements, the neural tube starts forming from the neural plate. Look on the dorsal side of your eggs for evidence of an invaginating neural plate. The sides of the plate gradually meet in the midline and fuse to form the neural tube.

Table 14.3 Stages in amphibian development

Stage

Description

Ambystoma Rana sylvaticus Rana pipiens punctatum in hours at in hours at in hours at 18˚C 18˚C 18˚C

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

Unfertilized Gray crescent 2-Cell 4-Cell 8-Cell 16-Cell 32-Cell Mid-cleavage Late cleavage Early gastrula, dorsal lip forming Mid-gastrula Late gastrula Neural plate Neural folds Rotation Neural tube Tailbud Muscular response Heartbeat Gill circulation, hatching

0 1 2.5 3+ 4.5 5+ 6 12 16 19 24 28 36 40 45 50 58 65 75 90

Source: After Rugh, 1948.

0 1 3.5 4.5 5.7 6.5 7.5 16 21 26 34 42 50 62 67 72 84 96 118 140

0 15.5 hrs 21 hrs 25 hrs 27 hrs 31 hrs 2 days 4 days 4.5 days 5 days 6 days 7 days

Amphibian Development (B)

(A)

(C)

(D)

(E)

(I)

(J)

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Gray crescent (G)

(F)

(H)

Dorsal lip (K)

(M)

(L)

Yolk plug

(N)

Yolk plug

(O)

Neural plate

Figure 14.3 Developmental stages of amphibians. Though there are a number of different staging series available for different species of anurans and urodeles, this simplified series (based on Rugh, 1948) can be used, with some exceptions, for most species of anurans and urodeles up to hatching. The diagrams illustrate the stages listed in Table 14.3. (A) Stage 1: Unfertilized egg. (B) Stage 2: Fertilized egg. There is a shifting of cytoplasm that uncovers a lighter area of pigmentation called the gray crescent. This is the precursor to the notochordal mesoderm. (Does not appear in the African clawed toad, Xenopus laevis. (C) Stage 3: 2 Cells. The first cleavage plane usually bisects the gray crescent. (D) Stage 4: 4 Cells. The second cleavage plane is parallel to the first. (E) Stage 5: 8 Cells. The third cleavage plane is perpendicular to the first two. (F) Stage 6: 16 Cells. The fourth cleavage plane is parallel to the first two. (G) Stage 7: 32 Cells. The fifth cleavage planes are parallel to the third. (H) Stage 8: Mid-cleavage. Cleavage is faster in the animal hemisphere than in the vegetal hemisphere, so the cells in the animal half are smaller and more numerous than those in the vegetal half. (I) Stage 10: Dorsal lip forms. Gastrulation is just beginning as cells involute forming the dorsal lip of the blastopore. (J) Stage 11: Mid-gastrula. The embryo is shown from the lower side. Cells are involuting from all sides of the blastopore. Large, yolky endoderm cells caught in the middle form the yolk plug. (K) Stage 12: Late gastrula. The embryo is shown from the lower side. As gastrulation proceeds, the yolk plug gets smaller. (L) Stage 13: Neural plate forms. The embryo is shown from the dorsal side. Neural ectoderm begins the process of forming a neural tube. (M) Stage 14: Neural folds form. The embryo is shown from the dorsal side. The edges of the neural plate are folding upwards. (N) Stage 15: Rotation begins. The embryo shown from the dorsal side. The neural folds are closing to form a neural tube. The epidermis of the embryo is ciliated, and the beating of the cilia cause the embryo to rotate within its vitelline envelope. (O) Stage 16: Neural tube. The embryo is shown from the dorsal side. The neural tube is now closed. The bulge of the forming eyes can be seen at the anterior end.

Amphibian Development Figure 14.4 Stages in amphibian development continued. (A) Stage 17: Tail bud forms. The embryo is shown from the dorsal side. The tail begins forming. A short stubby tail bud is seen at the posterior end. (B) Stage 18: Muscular response begins. The embryo is shown in lateral view. The embryo and tail bud are elongating, and the gill plate, which will form external gills, is visible. Muscular movements have started, and the embryo can be seen to twitch within its vitelline envelope. (C) Stage 19: Heartbeat begins. The embryo continues to elongate, the gill plate is subdivided, and the heart has begun to beat. (D) Stage 20, gill circulation and hatching. The embryo hatches from its vitelline envelope, the gill plate is further subdivided, and gill circulation is evident. (E) A generalized diagram of a young tadpole, the larva of an anuran. They possess oral suckers which allow them to attach. The gills are no longer visible, being enclosed in an opercular chamber with usually only one opening to the outside. (F) A generalized diagram of a young larva of a urodele. Called simply a salamander larva, it has a pair of balancers and visible external gills, which become long and featherlike as the larva grows. (After Rugh, 1948.)

(A)

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(B)

(C)

Gill plate

Tail bud (D)

External gills developing

(E)

(F)

Oral sucker

Balancer

Opercular opening

External gills

If your embryos are rotating within their vitelline envelope, then you know that they are already ciliated. This will occur just after the two sides of the neural plate have met in the midline. At this point, the brain should begin to look subdivided, and the bulges of the optic cups are obvious. Following this, the tail, starting as a short tail bud, begins to elongate as the entire embryo elongates. A gill plate will form, which eventually will be molded into external gills. Both anuran and urodele larvae form external gills, but the anurans secondarily enclose them in an opercular chamber, closing them off except for a single (though sometimes two) opercular opening(s). If your embryos are ready to hatch, they will be secreting a hatching enzyme that degrades the vitelline envelope and allows them to escape. You will know that hatching is near if the embryos are contracting their musculature, which will look like sudden twitches, and if their heart has started beating. If you have embryos this advanced, be sure to watch them carefully. Hatching is an exciting event to be witness to. Urodele and anuran larvae initially look quite similar, but this similarity soon wanes as they continue their development. Urodele larvae, called simply salamander larvae, continue to elongate, their gills become long and featherlike, and a pair of balancers develops. These balancers aid in keeping the larva from sinking into the mud. Salamander larvae are carnivores and only eat food that is

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alive and wiggling. Hand-feeding salamander larvae becomes a game of making raw liver look like a wriggling insect larva by waving it on the end of a stick. Anuran larvae are called tadpoles. The body becomes ovoid followed by a long narrow tail. The intestines are especially noticeable, being long and coiled within the short body, making it look (I’ve always thought) like an hors d’oeuvre ready for a cracker. Tadpoles are vegetarians and need this large intestinal surface area to absorb nutrients. Tadpoles lack balancers but have an oral sucker that allows them to attach to sticks and other objects. Environmental effects on amphibian development Construct a chart showing the species of eggs that you found, percent survival, their developmental stage, any developmental defects observed, air temperature, and the temperature, pH, and condition of drying of their habitat. Look for specific effects due to acidity (described below), and add these to the chart. One of the major effects of acidity on amphibian development is the loss of perivitelline space and the failure of the vitelline envelope to expand as the embryo elongates. The result is that the embryo becomes tightly coiled within the rigid, spherical envelope. In many cases, these coiled embryos never hatch. The acidity appears to inhibit the hatching enzyme. If they do manage to hatch, their vertebral columns are severely deformed and their tails are curled and bent. Other effects that have been noted are abnormal gill development and abnormal swelling near the heart. Physiologically, acidity disrupts the sodium and chloride balance within the embryo by inhibiting active uptake of these ions. Passive loss of the ions results in a net loss that eventually causes death. Acidic conditions may also arrest development during early cleavage. In this case, the egg capsules become opaque and shrink. It has been reported that low pH can even prevent the first step in development—fertilization—by inactivating sperm, though the pH at which this occurs is debated. A pH lower than 4.0, however, is undoubtedly harmful to amphibian sperm. Determine whether any of the abnormalities you found in your samples appear to be related to acid stress, and compare your results with those listed in Table 14.4. The table shows clearly that the range of pH tolerance varies among different species. What the table does not show is that this tolerance can vary even among different populations of a single species. For example, though the critical pH range for Ambystoma maculatum is given as 5–7 based on studies of several populations, another population was reported to have a critical range that started as low as pH 4.2. It appears that different populations are adapted to the natural pH of their habitat. For example, bogs typically are very acidic, with a pH as low as 4.2, so it can be expected that populations that lay their eggs in bogs will be more acid-tolerant than populations found in ponds or vernal pools. The data for a number of the species in the table below came from populations laying in bogs. The same species that you find laying in non-bog habitats may be far less acid-tolerant than these. Can you determine probable lethal, critical, or optimal pH levels for your embryos from your data? If so, add these to Table 14.4, recognizing that this is only preliminary data. Draw conclusions from your studies, and share these with the class. Data compiled from the entire class can be entered into your laboratory notebook. These data are valuable. Compiled over several years they may show trends that will be useful to environmental study groups. I invite you to send your carefully (of course) compiled results to an amphibian protection group in your area. The names of these vary among states and provinces, but you most likely will be able to find the name of one by calling your state or provincial Nongame and/or Endangered Species Program. If none is listed, look in the phone book for government listings to find numbers for a Department of Wildlife, Natural Resources, or Conservation that you can call instead.

Amphibian Development

Table 14.4 pH tolerance of some common North American amphibians during development

Species

Lethal pH

Critical pH (<50% survival)

Optimal pH (maximal survival)

Ambystoma jeffersonianum embryo

4.0–4.6

4.0–5.0

5–6

Ambystoma maculatum embryo

4.0–5.0a

5.0–7.0 b

7–9

Acris gryllus embryo

4.1

4.2–4.6

?

Bufo americanus* embryo

3.5–3.9

4.0–4.5

?

Hyla versicolor* embryo

3.8

3.9–4.3

?

Pseudacris crucifer* embryo

3.8

4.0–4.2

?

Rana catesbeiana* embryo

3.9

4.1–4.3

?

Rana clamitans* embryo

3.8

3.8–4.1

?

Rana palustris* embryo

4.0

4.2–4.4

?

Rana pipiens Sperm Embryo

4.5c 5.0

5.5c 6.0

6.5–7.0 6.3–7.5

Rana sylvatica Embryo Larva Metamorphosis

3.0–3.5d 3.25 3.25

3.75 3.5 (60%)e 3.5 (90%)

4.0–7.6 4–7.6 3.75–7.6

*Data from species living in a bog a

Failure to retract yolk plug, posterior trunk deformities, perivitelline space shrinks forcing embryo into tight coils. b c

d e

Swelling of chest near heart and abnormal gills.

Measured in buffered solutions. Arrested in early cleavage, egg capsules become opaque and shrink.

Abnormal, curled and bent tails. Source: Based on data in Pierce, 1985; Pierce, et al., 1984; Pough, 1976; and Schlichter, 1981.

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Adopt an egg mass Your estimates of developmental success will be greatly improved if you are able to return to a collecting site over a period of time and repeat your observations of percentage of survival within an egg mass, repeating temperature and pH readings and recording amount of drying of the habitat. To do this, it would be best to adopt one egg mass for continued monitoring. If time allows, you could test effects of ultraviolet radiation in the field by protecting some of the eggs using UV-blocking filters as discussed in Chapter 7. As a control, you can keep some of the eggs from the mass in the laboratory. Keep these eggs at a favorable temperature (18°C is usually adequate) and grow the embryos in Holtfreter’s solution, a balanced salt solution designed especially for amphibian embryos. Compare the success rate of development between the embryos grown in the laboratory and those left in the field. Add these data to your chart. And remember at all times not to contribute to the problem of declining amphibian populations—never take large numbers of eggs from the field, do not expose embryos in the field to additional hazards, and return your collected embryos to their original habitats at the end of your studies. Holtfreter’s solution NaCl KCl CaCl2 NaHCO3 Distilled water

0.35 gm 0.005 gm 0.01 gm 0.02 gm 100 ml

Stay in touch For those who have now developed or have always had a love for these distant relatives of ours, you can keep abreast of the issues through a number of websites. One site maintained by the IUCN/SSC Declining Amphibian Populations Task Force, http://www.open.ac.uk/daptf/index.htm, publishes an excellent, up-to-date “Froglog Newsletter.” Several others include Herp-Net, www.herpetology.com/herpnet.html, which is set up for international communication on amphibians and reptiles, www.frogweb.gov, a site for linking the public, press, and researchers, and www.in.gov/dnr/fishwild/endangered/naamp/nindex.htm, a site maintained by the North American Amphibian Monitoring Program, which is looking for volunteers for frog calling surveys, terrestrial salamander monitoring, and creating amphibian atlases to document habitat.

Accompanying Materials See Vade Mecum: “Amphibian.” This chapter of the CD shows in movies and labeled photographs the gametes, development, and metamorphosis of amphibians. Time-lapse photography of Xenopus laevis development is also included, as well as a gallery of adult frogs along with the calls of the males during mating season. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. Three different chapters give diagrams and discussion of the amphibian life cycle, cleavage, gastrulation, and metamorphosis.

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Fink, R. (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderalnd, MA. Sequence 8. This shows gastrulation in the South African clawed frog, Xenopus laevis.

Selected Bibliography Bantle, J. A., J. Dumont, N. R. Finch and G. Linder. 1991. Atlas of Abnormalities: A Guide for the Conduct of FETAX. Oklahoma State Publications, Stillwater, OK. The FETAX system was developed to standardize testing of toxicity levels of environmental hazards on amphibians. By using the laboratory raised Xenopus frogs rather than native species, it avoids putting even more pressure on these endangered species. Bishop, S. C. 1941. The Salamanders of New York. New York State Museum Bulletin No. 324. University of the State of New York, Albany, NY. This is a very detailed account of salamanders that occur in many regions besides New York. Specifics on breeding, egg masses, and individual eggs make it invaluable for species identification of eggs. Bishop, S. C. 1943. Handbook of Salamanders: The Salamanders of the United States, of Canada, and of Lower California. Comstock Publishing, Ithaca, NY. This is one of the classics that all herpetologists know well. More conversational than his earlier work, it is both entertaining and an excellent reference. Blaustein, A. R., J. M. Kiesecker, D. P. Chivers, D. G. Hokit, A. Marco, L. K. Belden and A. Hatch. 1998. Effects of ultraviolet light on amphibians: Field experiments. Amer. Zool. 38: 799–812. This excellent review of the effects of UV radiation on amphibians also discusses the synergistic effects of ultraviolet radiation and low pH, which can greatly increase mortality. Blaustein, A. R. and D. B. Wake. 1995. The puzzle of declining amphibian populations. Sci. Amer. 272: 52–57. A beautifully illustrated article on a number of the threats to amphibian species, including UV radiation, destruction of habitat, pollution, disease, and human uses. Doyle, R. 1998. Amphibians at risk. Sci. Amer. 279: 27. This short article shows maps of areas in the United States where amphibians are at risk. Duellman, W. E. and L. Trueb. 1994. Biology of Amphibians. Johns Hopkins University Press, Baltimore. A general textbook about amphibians, covering anatomy, physiology, reproduction, ecology, and evolution. Its discussion of mating behaviors and breeding patterns is especially useful for this laboratory. Elliott, L. 1998. A Guide to Night Sounds NatureSound Studio, P.O. Box 84, Ithaca, NY 14851-0084. This extremely useful collection is available as an audiocassette or CD and comes with a guide. It is a wonderful introduction for the novice. And it’s cheap. It includes calls of ten of the most common anurans as well as night-calling birds, a few insects, and some mammals. It can be ordered from the NatureSound website, www.naturesound.com, or from North Sound by calling 1-800-336-6398. Elliott, L. 1998. The Calls of Frogs and Toads. NatureSound Studio, P.O. Box 84, Ithaca, NY 14851-0084. More comprehensive than the selection above, this collection is available in both audiocassette and CD form. It includes the sounds of 42 species of anurans along with a booklet on identification. It can be ordered from the NatureSound website, www.naturesound.com, or from North Sound by calling 1-800-336-6398. Freda, J. 1991. Effects of acidification on amphibians. In Acidic Deposition: State of Science and Technology, Vol. II. Aquatic Processes and Effects. U.S. National Acid Precipitation Assessment Program, Washington, D.C., pp. 13-135–13-151. A good summary of the effects of acid rain on amphibian development, this both reviews and critiques early and more recent studies.

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“Froglog.” Newsletter from the World Conservation Union (IUCN), Species Survival Commision (SSC), Declining Amphibian Populations Task Force (DAPTF), and Center for Analysis of Environmental Change (CAEC). This is an extremely informative and concise publication available on the web at www.open.ac.uk/daptf/froglog/. Heyer, W. R., M. A. Donnelly, R. W. McDiarmid, L. C. Hayek and M. S. Foster (eds.). 1993. Measuring and Monitoring Biological Diversity: Standard Methods for Amphibians. Smithsonian Institution Press, Dept. 900, Blue Ridge Summit, PA 17294-0900, 1- 800-782-4612. Very recent, and inexpensive in its paperback version, this is an important book for anyone who would like to help in conducting amphibian counts for their geographic area and protecting local populations. It gives standardized methods for sampling and information on analyzing and using data. Houck, L. D. 1998. Integrative studies of amphibians: from molecules to mating. Amer. Zool. 38: 108–117. This is a nice discussion of courtship pheromones used by male salamanders in courtship. Howells, G., 1990. Acid Rain and Acid Waters. Ellis Horwood Limited, West Sussex, England. This has only a short section on the effects of acid rain on amphibians, but helps to give a general understanding of the global effects of acid rain. Hunter, M. L., Jr., J. Albright and J. Arbuckle (eds.). 1992. The Amphibians and Reptiles of Maine. Maine Agricultural Experiment Station, University of Maine Press, Orono, ME. Hunter, M. L., A. J. K. Calhoun and M. McCollough. 1999. Maine Amphibians and Reptiles. University of Maine Press, Orono, ME. All herp-enthusiasts, whether seasoned pros or the newly initiated, should have this book. It is an extremely complete compilation of information about amphibians and reptiles in Maine, but most of the information is applicable to many regions. It is delightfully written and beautifully illustrated with color photographs, pen-and-ink drawings, and includes a CD on calls of male frogs. La Clair, J. J., J. A. Bantle and J. Dumont. 1998. Photoproducts and metabolites of a common insect growth regulator produce developmental deformities in Xenopus. Environ. Sci. Technol. 32: 1453–1461. A paper that uses the frog embryo teratogenesis assay-Xenopus (FETAX) method to determine the level of toxicity of a commonly used insecticide. Pierce, B. A. 1985. Acid tolerance in amphibians. BioScience 35: 239–243. This is a very useful, easily understood review article, including a table and a long reference list. Pierce, B. A., J. B. Hoskins and E. Epstein. 1984. Acid tolerance in Connecticut wood frogs (Rana sylvatica). J. Herpetology 18: 159–167. An important study on the effects of acid rain, it includes information on different stages of development as well as larvae. Phillips, K. 1995. Tracking the Vanishing Frogs: An Ecological Mystery. Viking Penguin, New York. Geared toward a lay audience, this well-written account reveals the devastation to amphibian populations caused by human activities. Pough, F. H. 1976. Acid precipitation and embryonic mortality of spotted salamanders, Ambystoma maculatum. Science 192: 68–72. One of the early articles on the effects of acid rain on amphibians, it is cited in all review articles on the subject. Rugh, R. 1948. Experimental Embryology: A Manual of Techniques and Procedures. Burgess Publishing, Minneapolis, MN. An extremely valuable, out-of-print manual that was written by one of the venerable experts in amphibian development. The careful descriptions of techniques and developmental stages, illustrated with drawings and photographs, are invaluable to any embryologist. Look for it on out-of-print book lists, and snatch it up when you see it. Schlichter, L. C. 1981. Low pH affects the fertilization and development of Rana pipiens eggs. Can. J. Zool. 59: 1693–1699. This study, though somewhat confusing, is the first to attempt to determine

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effects of acidity on amphibian sperm. The conclusions are now in some dispute, since buffered solutions were used and the buffers themselves might have been toxic to the sperm. Stebbins, R. C. 1951. Amphibians of Western North America. University of California Press, Berkeley. An extremely thorough guide that includes information on breeding, egg masses, and larvae for each of the species covered. Stebbins, R. C. 1985. A Field Guide to Western Reptiles and Amphibians, 2nd Ed. Peterson Guides, Princeton, NJ. This is an excellent guide, not as detailed as the volume above, but very handy in the field. Tome, M. A. and F. H. Pough. 1982. Responses of amphibians to acid precipitation. In Acid Rain/Fisheries. Proceedings of an International Symposium on Acid Precipitation and Fisheries Impacts in Northeastern North America, Ithaca, New York, August 2–5, 1981. American Fisheries Society, Bethesda, MD, pp. 245–254. An older review article than that by Freda, this is still useful, including a number of charts and tables. Twitty, V. C. 1966. Of Scientists and Salamanders. W. H. Freeman, San Francisco. A truly delightful account of one embryologist and his work, this stands as a valuable historical work and entertaining bedtime reading. Tyning, T. F. 1990. A Guide to Amphibians and Reptiles. Little, Brown, Boston. This well-written, informative guide concentrates on relatively few species, but gives extremely useful information about their behavior and mating. Wright, A. H. and A. A. Wright. 1949. Handbook of Frogs and Toads of the United States and Canada, 3rd Ed. Comstock Publishing, Ithaca, NY. What the Bishop handbook is for salamanders, this classic is for anurans. Illustrated with photographs of adults, larvae, and egg masses, it an important reference to add to any herpetologist’s or naturalist’s library. Wyman, R. L. 1990. What’s happening to the amphibians? Cons. Biol. 4: 350–352. This is an excellent summary of the problem of declining amphibian populations.

Suppliers Cole-Parmer Instrument Company 625 East Bunker Ct. Vernon Hills, IL 60061 1-800-323-4340 www.coleparmer.com Cole-Parmer has a large array of inexpensive to top-of-the-line pH meters. They also have inexpensive glass pocket thermometers.

Fisher Scientific 711 Forbes Ave. Pittsburgh, PA 15219-4785 1-800-766-7000 www.fishersci.com Fisher offers a large array of pH measuring devices. Among these are the most expensive and reliable Orion meters with Orion Ross electrodes, inexpensive and expensive pocket pH meters, colorimetric pH measuring kits, pH paper, and pH paper for soil testing. They also have a wide range of metal and glass pocket and

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field thermometers. You also can get 0.01 N HCl from Fisher and dilute it to 0.001 N (pH 4.0) for testing electrodes on low ionic acidic solutions.

Ward’s Natural Science Establishment Inc. P.O. Box 92912 5100 West Henrietta Road Rochester, NY 14692-2660 1-800-962-2660 www.wardsci.com Ward’s offers a colorimetric pH measuring kit (for acid rain), an inexpensive portable water testing instrument that tests pH and temperature called Water Test Laboratory, and an acid rain test strip kit. They also have metal and glass pocket thermometers, from inexpensive to expensive.

Glossary Acid rain: The general term for precipitation with a pH below 5.6. Pollutants, created through human activity, have contributed significantly to acid rain, resulting in the acidification of lakes as well as soils. This acidification is threatening both the life of organisms in the lakes as well as forests, leading to forest declines. Amplexus: Among the anurans, the embrace by the male of the female during mating. The male grasps the female around the abdomen. This stimulates the female to extrude her eggs. The male releases his sperm over the eggs as they are extruded. Fertilization, therefore, is external. Anamniote: Vertebrates that do not have an amnion during development, the fishes and amphibians. Animal pole: The point on the surface of the egg where polar bodies are given off. Eggs usually orient to gravity with their animal pole oriented uppermost. Anurans: Literally, no tail. The group of amphibians lacking a tail; frogs and toads. Archenteron: Literally, ancient gut. The primitive gut formed during gastrulation. Balancer: A structure on the side of an organism that helps the organism to stabilize while flying or swimming. In Drosophila, the balancers sit posterior to the wings and represent a modified pair of wings. In the salamander larva, the balancers aid in keeping the larva from sinking into the mud. Blastocoel: The cavity within a blastula. Blastopore: In a gastrulating embryo, the point at which cells are moving inward to form the archenteron. In deuterostomes, the blastopore becomes the anus; in protostomes, the mouth forms at or near the blastopore opening. Blastula stage: The stage in embryonic development following cleavage and preceding gastrulation. The embryo may form a hollow blastula (coeloblastula) or a solid blastula (stereoblastula). The cavity formed in a coeloblastula is called the blastocoel. Cleavage: The stage in embryonic development, normally following fertilization, in which the egg, or portions of the egg, divide by mitosis into a number of cells. Cleavage results in the formation of the blastula. Cloaca: Literally means sewer. In a chordate, the terminal region of the endodermal hindgut. In general, it receives the ducts and products from the gonads, digestive, and excretory organs. Coelom: The body cavity in which the gut is suspended. It is lined by mesoderm.

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Dorsal lip of blastopore: In an amphibian embryo, this is the region where gastrulation movements first appear as surface cells move inward by involution. Ectoderm: The germ layer in an embryo that gives rise to the epidermis and nervous system. In many cases, as in vertebrates, an anterior and posterior invagination of ectoderm gives rise to the stomodeum and proctodeum, respectively. Endoderm: The germ layer in an embryo that gives rise to the epithelium (lining) of the gut and gut derivatives. Epidermal ectoderm: The region of ectoderm that forms the epidermis (the epithelial layer of the skin). Epidermis: The epithelium of the skin. It is typically a stratified squamous epithelium, with outer keratinized layers of cells that protect against dehydration and physical abrasion. Epithelium: A major type of tissue in an organism, found lining body cavities and covering the outside of the body. The epidermis is an example of a specific type of epithelium. Exogastrula: A deformity in developing embryos in which the archenteron, instead of invaginating, evaginates, forming an external archenteron. External gills: Gills which protrude into the external environment. Fertilization: Fusion of egg and sperm. Gastrulation: The stage in embryonic development that follows cleavage. During gastrulation, cells rearrange themselves, the endoderm and mesoderm cells moving inward and the ectoderm cells spreading around the outside, with some ectoderm invaginating to form such structures as the stomodeum and proctodeum. Germ layers: The ectoderm, mesoderm, and endoderm of a developing embryo. Gill plate: In an amphibian embryo, the precursor to the external gills. Gray crescent: In an amphibian egg, a crescent-shaped lightly pigmented region formed just after fertilization. It is formed by a shifting of the highly pigmented cortical cytoplasm relative to the less pigmented cytoplasm beneath. It forms directly opposite the point of sperm entry and represents an area that will later form the notochordal mesoderm. Hatching enzyme: An enzyme produced by an embryo that creates a hole in the vitelline envelope allowing the embryo to escape. Herpetology: The study of amphibians and reptiles. Holoblastic cleavage: Complete cleavage. The pattern of cleavage in which the entire egg is divided into smaller cells. Holtfreter’s solution: A balanced salt solution designed especially for amphibian embryos. Intermediate mesoderm: Nephrogenic mesoderm. Invagination: The folding inward of a sheet of cells. This is a common type of movement during gastrulation. In the sea urchin, for example, the archenteron first forms by an invagination of the vegetal plate. Involution: A type of gastrulation movement in which a sheet of cells folds inward and spreads over an inner surface. Larva: Early stage or stages in the life cycle, differing significantly in morphology and ecology from the adult. Lateral plate mesoderm: Mesoderm lying lateral to the intermediate mesoderm. It delaminates into two layers: the splanchnic and somatic layers. The splitting creates a cavity, the body coelom. Melanin: A black pigment common in many animals. Mesoderm: The germ layer in an embryo that gives rise to the muscular and circulatory systems and most of the skeletal and urogenital systems.

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Mesolecithal egg: An egg with a moderate amount of yolk, such as the amphibian egg. Metamorphosis: The transformation from the larval to the adult stage. Neural crest cells: Cells found only in vertebrate embryos that start out residing on the crest of the neural tube and subsequently migrate to become a host of structures. Pigment cells (except for the pigmented retina), membrane bones in the face and skull, dentine-secreting cells of the teeth, the adrenal medulla, and sympathetic ganglia of the autonomic nervous system are just some of the structures formed. Neural ectoderm: The region of ectoderm that forms the neural tube. Neural plate: The neural ectoderm prior to its invagination to form the neural tube. Neural tube: The tube formed by the neural ectoderm. The precursor to the brain and spinal cord. Neurula: The stage in embryonic development when neurulation is occurring. Neurulation: The morphogenetic movements of the neural ectoderm that form a neural tube out of the neural plate. Notochord: The axial supporting structure in a chordate embryo formed from notochordal mesoderm. Notochordal mesoderm has been called the “primary organizer” because of its inducing functions that initiate an axial organization of the embryo. The notochord is a diagnostic characteristic of the chordates. Notochordal mesoderm: The region of mesoderm that forms the notochord. Opercular chamber: In the larva of an anuran, the chamber that surrounds the gills. In frog and toad larvae, gills start out as external gills, but are secondarily enclosed by a flap of skin that grows over the gills to create the opercular chamber. Opercular opening: In the larva of an anuran, the opening of the opercular chamber to the outside. Optic cup: In a vertebrate embryo, the precursor to the neural and pigmented retinas of the eye. During eye development, optic vesicles that form as outpocketings of the diencephalon invaginate to form the optic cups. Oral sucker: A structure in the oral region of an organism that allows the organism to attach to a substrate or another organism. In the anuran larva, the oral sucker allows the larva to attach to sticks and other objects. Ozone: O3. An allotropic (alternative) form of oxygen. In our upper atmosphere, it protects the Earth’s surface from UV-B radiation. In the lower atmosphere, it is a pollutant that is harmful to the health of organisms. PAR: Stands for photosynthetically active radiation. This is the range of wavelengths (400–700 nm) that can activate photosynthetically active pigments. It is not considered harmful. Perivitelline space: The space between the egg cell membrane (or embryo) and the vitelline envelope that surrounds the egg (or embryo). Radial cleavage: The cleavage pattern in which each cleavage plane is either parallel or perpendicular to the animal–vegetal axis. Somatic lateral plate mesoderm: The layer of lateral plate mesoderm lining the coelom and that is associated with the epidermal ectoderm to form the somatopleure. This layer of mesoderm is not able to form blood vessels. Somites: Blocks of mesoderm that become subdivided into a sclerotome, myotome, and dermatome. Sclerotomes form vertebrae and ribs, myotomes form skeletal muscles for the back and appendages, and dermatomes give rise to dermis. Somitic: Of the somite.

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Spermatheca: A specialized region of a female reproductive tract for storing sperm. Female salamanders have a spermatheca that represents a specialized region of the cloaca. As eggs pass by the spermatheca, they are fertilized by the sperm. Spermatophores: A small packet of sperm produced by some species with internal fertilization. Male salamanders produce spermatophores. The male deposits the spermatophore on the bottom of a pool and leads the female over it. The female picks up the packet with the lips of her cloaca. Splanchnic lateral plate mesoderm: The layer of lateral plate mesoderm lining the coelom and that is associated with the endoderm to form the splanchnopleure. This layer of mesoderm is able to form blood vessels. Tailbud: In an embryo, the posterior bud that will form the tail. Ultraviolet radiation: Radiation of wavelengths that fall between 200 and 400 nanometers (nm). Urodeles: Literally, tailed. The group of amphibians that have tails; salamanders. UV-A: The longest wavelengths (320–400 nm) within the ultraviolet range. This range is harmful to biological organisms, but is less harmful than UV-B or UV-C. Most of the UV radiation reaching the Earth’s surface is in the UV-A range. UV-B: The range of wavelengths within the ultraviolet that fall between 280–320 nm. (Some sources use 290–320 nm.) UV-B radiation is significantly more harmful to biological organisms than UVA. Most of the UV-B radiation coming from the sun is blocked by the ozone in the Earth’s upper atmosphere. Holes developing in this ozone layer are allowing greater amounts of UV-B radiation to penetrate to the Earth’s surface. UV-B filters: Filters that block UV-B radiation, but allow UV-A radiation and PAR to pass through. Examples include glass, Xerox transparencies, plastic milk bottles, and Mylar film. UV-C: Ultraviolet radiation in the range of 200–280 nm. This is the most highly dangerous range of UV radiation for biological organisms. It is absorbed by oxygen in the atmosphere, such that it never reaches the Earth’s surface. UV-opaque filters: Filters that block UV-A and UV-B radiation, but allow PAR to pass through. Examples include standard plexiglass, sunglasses rated as 100% UV-blocking, light-block plastic milk bottles, and many cosmetic sunscreens. UV-transparent filters: Filters that allow UV-A, UV-B radiation and PAR to pass through. Examples include plastic petri dishes, plastic sandwich bags, plastic food wrap, UV-transparent plexiglass, and cellulose acetate. Vegetal pole: The pole that is opposite the animal pole. Yolk is more concentrated at the vegetal pole. Eggs usually orient to gravity such that their vegetal pole is oriented downward. Vertebrata: The class of chordates that include the fish, amphibians, reptiles, birds and mammals. Diagnostic features of the vertebrates are a vertebral column (hence its name), neural crest cells during development, and the presence of bone. Vitelline envelope: An extracellular coat surrounding an ovum. Xenopus laevis: The South African clawed frog or African clawed frog (previously commonly called the South African or African clawed toad). It is a species of frog that has become a model organism for amphibian development because of the ease with which it can be raised and maintained in the laboratory. Yolk plug: In an amphibian embryo, the plug of yolky endoderm cells caught between the lips of the blastopore during gastrulation. It is eventually internalized. Zygote: A fertilized egg following fusion of the egg and sperm.

Amphibian Development Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. How does salamander mating differ from that of frogs?

2. What is a spermatophore? What are the various parts of a spermatophore?

3. What are some environmental factors that threaten amphibian survival?

4. Why do you think amphibian embryos are particularly vulnerable to environmental hazards?

Name: _______________________

Amphibian - 2

5. What are some of the amphibian species that live in your area?

6. What type of egg does an amphibian have? What is its pattern of cleavage?

7. What protection does the vitelline envelope provide the amphibian embryo?

8. What is the gray crescent, and when do you see it in amphibian development?

9. What is a yolk plug in the amphibian gastrula?

10. What differences can you list that distinguish frog larvae from salamander larvae?

15

chapter

Zebrafish Development

Teleost: Danio rerio

A New Model Organism The zebrafish, Danio rerio, is a relatively new and important addition to the family of model organisms. Appreciated for its transparent eggs, rapid embryonic development (in only 24 hours all the major organ systems have been laid down), quick generation time (it takes only 2–3 months for the fish to reach reproductive age), and ease of maintenance, it became a favored laboratory organism beginning in the 1970s, due primarily to the efforts of George Streisinger, who recognized the advantage of zebrafish for genetic studies. Since then, two large-scale mutagenic screens of the zebrafish genome have created a wealth of information for genetic and developmental studies (see, e.g., Development, 123 published in 1996). The entire issue is devoted to the description of approximately 2000 mutations that disrupt zebrafish development.) The zebrafish is a small (4–5 cm), tropical, freshwater teleost fish, closely related to the goldfish, and originally identified in the Ganges River in northeastern India. It is now commonly available in pet stores and can be purchased from a number of biological supply companies as well. It can be easily maintained in aquaria kept within a 25–28.5ºC temperature range, and spawning can be induced at any time of year through an appropriate feeding regime and light–dark cycle. Each mature female is able to lay as many as 100–200 eggs, and these, since they are demersal, sink to the bottom and can be collected simply by siphoning them up from among the marbles that are placed at the bottom of a spawning chamber. Because the embryos develop quickly in a beautifully transparent egg, the zebrafish allows an exquisite look at the entire course of vertebrate development, unencumbered by either the opaqueness of the amphibian egg or the inaccessibility of the amniote egg (reptiles and birds developing within an opaque shell and placental mammals within the uterus of the mother). Not only does the zebrafish lend itself to the study of normal vertebrate development, it also can serve as a convenient indicator of possible teratogens in our diet and environment. It is these that our study will focus on. A number of substances can be tested. For example, the teratogenic effects of excess vitamin A have recently become a severe threat because of the use of isotretinoin (Accutane®), a form of vitamin A that is used to help treat severe acne. Can the zebrafish embryo be used as a model to determine the detrimental effects of excess vitamin A? Lithium is another common medication (used to treat bipolar disorder) that has been known for over a hundred years to produce embryonic malformations. Are there safe

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levels of lithium, and can these be determined using zebrafish embryos? Most people are aware that our environment is riddled with pollutants, but few realize the detrimental effects that these can have on embryos. For example, if the power plant down the road from you is spewing out heated water that raises the temperature of the water ways near the effluent pipes, will this have a detrimental effect on the development of organisms in the area? You can get some answers by testing the temperature range that zebrafish can tolerate. Are they immune to temperature fluctuations? Many of the pollutants we produce in industrialized countries have estrogenic effects (pesticides including DDT, plastics, even the containers that you drink your milk and orange juice from). These are called endocrine disruptors. Can we use zebrafish to detect the ill-effects of these substances? The first thing we must do in pursuing this is to learn the details of normal zebrafish development. As you review normal development, think of possible experiments you might design that would test possible teratogens. There are a number of suggestions given later in this chapter, far more than can be covered in a single laboratory period. Because of the nature of these experiments, you may be choosing to spend several weeks studying zebrafish.

Normal Development Development in the zebrafish is exceedingly fast as compared to that in the chick. What takes days to occur in the chick embryo happens within hours in the zebrafish embryo. As you study the development of the zebrafish, you should be referring back to the chapters on chick development to review the general features of vertebrate embryogenesis. Also, consult Table 15.1 and Figures 15.1–15.3 for descriptions of each of the stages of zebrafish development. Fertilization The zebrafish egg is small (approx. 0.5–0.6 mm in diameter prior to fertilization). It is a macrolecithal egg, meaning that it contains a relatively large quantity of yolk (macro=large; lecithal=yolk). Surrounding the egg is an extracellular layer called the vitelline envelope, and outside of this is a tough extracellular coat called the chorion. Fertilization in the zebrafish is external. After brief courtship maneuvers in which the male swims after the female, the female sheds her eggs, and the male deposits sperm over the eggs. These sperm, released into the water, must now make their way through the tough chorion to the surface of the egg in order to fuse with the egg. They accomplish this by finding the micropyle, a small hole through the chorion at the animal pole. The first sperm to arrive at the egg cell surface attaches to microvilli on the egg surface. These microvilli elongate to create a fertilization cone while the sperm fuses with the egg cell membrane, and the micropyle subsequently becomes plugged, preventing further sperm from entering. This is one of the mechanisms used to prevent polyspermy. A further block to polyspermy is initiated by a calcium-dependent cortical granule reaction, which is activated by the fusion of the egg and sperm. In the cortical granule reaction, the cortical granules of the egg fuse with the egg cell membrane, dumping their contents into the extracellular space. The components of the granules in the extracellular compartment absorb water, and as a result, the chorion swells and lifts away from the surface of the egg. When the chorion is fully lifted (12 minutes post-fertilization), the egg diameter has increased to approximately 1 mm. Upon fertilization, cytoplasmic streaming within the egg reorganizes the previously homogeneous contents, segregating the yolk-free cytoplasm to the animal pole until it sits as a cap of clear cytoplasm on the yolky sphere below. This clear cytoplasm is the blastodisc (Figure 15.1A).

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Table 15.1 Staging series of embryonic zebrafish development, at 28.5ºC. (After Kimmel et al., 1995.) S t a g e a n d length o f incubation

Description of embryo

Fertilization (0 hours)

Newly fertilized egg undergoes cytoplasmic streaming, producing blastodisc at animal pole.

Cleavage (45 minutes – 2 hours)

Cleavage begins after 45 minutes, and continues synchronously at 15minute intervals.

2-cell (1st cleavage) (45 minutes)

Cleavage is meroblastic (incomplete). The two cells are continuous with the yolk.

4-cell (2nd cleavage) (1 hour)

Cleavage is meroblastic, cells are arranged in 2 rows of 2 cells per row (2 × 2).

8-cell (3rd cleavage) (1 1/4 hour)

Cleavage is meroblastic, cells are arranged in 2 rows of 4 cells per row (2 × 4).

16-cell (4th cleavage) (1 1/2 hour)

Cleavage of outer cells is meroblastic, central 4 blastomeres are complete. Cells are arranged in 4 rows of 4 cells per row (4 × 4).

32-cell (5th cleavage) (1 3/4 hour)

Cleavage of non-marginal blastomeres is complete. Cells are not always arranged regularly, though can be in 4 rows of 8 cells per row (4 × 8). In side view appears to be two tiers of cells.

64-cell (6th cleavage) (2 hours)

Horizontal cleavage planes produce two tiers of cells, though in side view it appears to be three tiers.

Blastula (2 1/4 – 3 1/2 hours)

Embryo continues to divide, becoming metasynchronous, adding tiers of blastomeres.

128-cell (7th cleavage) (2 1/4 hour)

Cells sit as a high mound in five irregular tiers.

256-cell (8th cleavage) (2 1/2 hours)

Cells are seen as seven tiers.

512-cell (9th cleavage) (2 3/4 hours)

Midblastula transition begins. Cell cycle begins to lengthen. Nine irregular tiers of cells are seen.

~1000-cell (10th cleavage) (3 hours)

Yolk syncytial layer (YSL) forms. About 11 tiers of cells can be seen.

High (3 1/3 hours)

Blastoderm is a high mound of cells. More than 11 tiers of cells can be seen.

Oblong (3 2/3 hours)

Blastoderm begins to flatten so that the egg looks oblong.

Sphere (4 hours)

The blastoderm continues flattening until the egg becomes spherical.

Dome (4 1/3 hours)

A short period during which the internal YSL domes upward toward the animal pole. This marks the beginning of epiboly, the movement of the blastoderm down around the yolk.

30% epiboly (4 2/3 hours)

Epiboly continues, reaching 30% of the distance between the animal and vegetal poles.

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Table 15.1 (Continued) Stage and l e n g t h o f incubation

Description of embryo

Gastrula (5 1/4–10 hours) 50%-epiboly (5 1/4 hours)

Germ-ring (5 2/3 hours) Shield (6 hours) 90%-epiboly (9 hours) Bud (10 hours)

Epiboly continues as gastrulation movements within the deep layer of cells separate the germ layers into an upper epiblast and lower hypoblast. The germ ring becomes visible at 50% epiboly. Embryonic shield forms. Brain rudiment is thickened by 90% epiboly. Tail bud forms and epiboly is complete.

Segmentation (10 1/3–22 hours) 1-somite (10 1/3 hours)

First somite forms.

5-somite (11 2/3 hours)

By the 5-somite stage optic vesicle is prominent.

14-somite (16 hours)

By the 14-somite stage, the otic placode is visible as are the brain neuromeres, and the trunk somites have a V-shape.

20-somite (19 hours)

By the 20-somite stage, muscular twitches are evident, the otic placode is now an otic vesicle, the eye lens is evident, hindbrain neuromeres are prominent, and the tail is well extended.

26-somite (22 hours)

By the 26-somite stage, the embryo flexes side to side. Otoliths can be seen in the inner ear (otic vesicle).

Pharyngula (24–42 hours) Prim-5 (24 hours)

By the Prim-5 stage, pigmentation in the pigmented retina of the eye as well as in the skin becomes apparent, the heart begins beating, and red blood cells can be seen in the yolk sac.

Prim-15 (30 hours)

By the Prim-15 stage, the embryo responds to touch, and spontaneous movements are reduced. The retina is well pigmented. The caudal artery extends halfway to the tail.

Prim-25 (36 hours)

By the Prim-25 stage, there is tail pigmentation, blood circulation is strong, a single aortic arch can be seen, and the caudal artery extends three-quarters of the way to the end of the tail.

High-pec (42 hours)

By the High-pec stage, a dechorionated embryo rests on its side after swimming. There is a complete dorsal stripe, xanthophores can be seen in the head region and iridophores can be seen in the retina. The mandibular and hyoid arches are visible.

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Table 15.1 (Continued) S t a g e a n d l e n g t h o f incubation

Description of embryo

Hatching (48 – 72 hours) Long-pec (48 hours)

Hatching occurs asynchronously. By the Long-pec stage, dorsal and ventral stripes meet at the tail. Iridophores are numerous in retina, head has a distinct yellow cast to it. 2–4 aortic arches can be seen with circulation in them.

Pec-fin (60 hours)

By the Pec-fin stage, movements are very rapid, the pectoral fin is flattened into a fin shape. Iridophores are seen in the dorsal stripe, early jaw cartilages are visible, and circulation can be seen in the 5–6 aortic arches.

Protruding-mouth (72 hours)

By the Protruding-mouth stage, the mouth is wide open and protrudes anterior to the eye. The dorsal body is as yellow as the head. Gill slits can be seen, and cartilage is present in the branchial arches.

Cleavage Cleavage begins approximately 45 minutes after fertilization (see Table 15.1 and Figure 15.1). Cleavage is initially incomplete (meroblastic). The cytoplasm of the blastodisc divides, but the cleavage furrows cannot penetrate the dense yolk below (Figures 15.1A–D). The first cells therefore are contiguous with the yolk, and up through the 8-cell stage, any substance injected into the yolk will also enter the cells of the blastoderm. The first six cleavages are synchronous, occurring at 15-minute intervals. The cleaved blastodisc is now called the blastoderm (Figure 15.1G). At the sixth cleavage, the first horizontal orientation of a cleavage plane occurs, resulting in two tiers of cells. Cleavage from here on becomes metasynchronous, in that waves of cell division pass through the blastoderm, and a multi-tiered mound of cells is produced. The body axes of the embryo are not specified in the cells of these early cleavages, which is unusual among vertebrate embryos. Blastula During the blastula stage, the cells within the blastoderm form three distinct layers. The first of these (forming between the ninth and tenth cleavages) is a layer called the yolk syncytial layer (YSL) (Figure 15.1I). This is created when cells around the edge of the blastoderm fuse with the yolk below. Initially, there is just a ring of nuclei, but these nuclei move and additional nuclei are added from the edge of the blastoderm, so that eventually there is an entire layer of nuclei lying within the yolky cytoplasm beneath the blastoderm. Next, the outer-most cells of the blastoderm establish themselves as a one-celllayer thick enveloping layer (EVL), and the cells left between the EVL and the YSL become the deep cells (Figure 15.2.A). The deep cells are the cells that will later form the body of the embryo. The EVL protects the developing embryo by forming a protective layer called the periderm, an outer layer of cells that is eventually sloughed. The YSL will lead the rest of the cells down around the yolk sphere during gastrulation. This spreading movement of the blastoderm, down around the yolk, is called epiboly, and the first stages occur while the embryo is still classified as a blastula. During this time, the blastoderm begins to

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flatten. Starting out with a high dome of blastoderm cells sitting atop the yolk sphere (Figure 15.1I), the blastoderm thins as it moves down around the yolk until the egg looks spherical (Figures 15.2A–D). The movements within the YSL that bring the rest of the overlying cells around the yolk during epiboly are dependent upon the action of microtubules and their associated motor proteins. Any environmental factor that disrupts microtubules, such as cold temperatures, therefore, would seriously interfere with epiboly.

Blastodisc (A)

(B)

1-Cell 15 min. (D)

2-Cell 45 min. (E)

8-Cell 2 rows of 4 1.25 hours (G)

(C)

Blastoderm

4-Cell 1 hour (F)

16-Cell 4 rows of 4 1.5 hours (H)

32-Cell 4 rows of 8 1.75 hours (I) YSL

64-Cell 2 hours

256-Cell 2.5 hours

High 3.3 hours

Figure 15.1 Diagrams of early development in the zebrafish embryo from fertilization to mid-blastula. Shown with chorion removed. (A) After fertilization, the nonyolky cytoplasm separates to form a clear blastodisc. (B) The first cleavage begins after 45 minutes. (C) Second cleavage produces four cells arranged in 2 rows of 2 cells each. (D) Third cleavage gives 2 rows of 4 cells each. (E) The fourth cleavage gives 4 rows of 4 cells each; the central blastomeres are complete, cut off from the yolk. (F) The fifth cleavage produces 32 cells, with central cells complete. (G) The sixth cleavage produces 64 cells in two tiers, with all the cells complete, with no connections to the yolk. (H) The eighth cleavage produces 256 cells, which appear as seven tiers of cells. (I) By 3.3 hours of development, the blastoderm is a high mound of cells. The yolk syncytial layer has been formed by cells fusing with the yolk below. Compare diagrams with descriptions of stages in Table 15.1.

Zebrafish Development

(A) EVL

(B) Deep cells

(C)

Germ ring

YSL Embryonic shield 30%-Epiboly 4.7 hours

(D)

Head region

Tail bud Bud 10 hours

50%-Epiboly 5.3 hours

Eye (E)

Otic placode

Somites 10-Somite 14 hours

Shield - Dorsal View 6 hours

(F)

Lens placode

Rhombomeres

Olfactory Somites placode 18-Somite 18 hours

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Figure 15.2 Zebrafish development continued. (A) 30% epiboly. The blastoderm now consists of three layers of cells: the outer enveloping layer (EVL), the yolk syncytial layer (YSL), and the deep cells. (B) 50% epiboly. The blastoderm continues to move down around the yolk. Involution begins around the margin of the blastoderm. (C) Shield stage, shown in dorsal view. Involution of deep cells at the margin of the blastoderm creates a thickened germ ring. Deep cells converging towards the dorsal side create a more pronounced thickening, the embryonic shield. (D) Bud stage, 100% epiboly. The body of the embryo, cupping the yolk along the dorsal side, now has a distinct head and tail region. (E) 10-somite stage. Somites are forming. The eye primordium and otic placodes are visible. (F) 18-somite stage. Olfactory placodes are present and the lens placode can be seen.

The blastula stage is the stage at which the genes of the embryo are turned on—prior to this, all development is controlled through messages that were stored in the egg during oogenesis (maternal mRNA). This turning on of the embryonic genome is called the mid-blastula transition and begins at the tenth cell division. Prior to this time, a teratogen that interfered with gene transcription would not harm the embryo. Gastrulation Gastrulation movements, those movements that define the three germ layers, all occur within the deep cells of the blastoderm. These movements begin once the blastoderm has spread about half way around the yolk (a stage called 50% epiboly; Figure 15.2B). Deep cells at the margin of the blastoderm begin to turn inward and spread on top of the YSL, a process called involution. This creates two layers within the deep cells, the upper epiblast and the lower hypoblast. The epiblast will form the ectoderm (the germ layer that forms the epidermis, or outer layer of skin, and the nervous system) and the hypoblast will form the mesoderm (the germ layer that forms muscle, the skeletal and circulatory

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systems, the kidneys, reproductive ducts and gonads, etc.), and endoderm (the germ layer that forms the inner lining of the gut and gut derivatives). The initial stages of involution create a thickened ring of deep cells, called the germ ring. As involution proceeds, deep cells converge towards the dorsal side creating a more pronounced thickening within the germ ring called the embryonic shield (Figure 15.2C). This embryonic shield is analogous to the dorsal lip of the blastopore of an amphibian embryo. It is here that the body of the embryo will be organized. By the time that the blastoderm has moved down around the yolk completely (100% epiboly or bud stage; Figure 15.2D), the body of the embryo will be cupping the dorsal side of the yolk as a long narrow thickening, with the brain region beginning to form anteriorly and the tail bud visible at the posterior end. Only ten hours have passed since the time the egg was fertilized and development began. Organogenesis Organogenesis is rapid, and most organ primordia have formed by 24 hours of development. (To review the structures and stages of vertebrate organogenesis, refer back to the chapters on chick development. Similar structures will be seen here, but they are forming much more rapidly.) Recall that a 24-hour chick embryo (having developed at a warmer temperature—38ºC) only has the beginnings of the neural region defined and approximately 4 pairs of somites. The 24-hour zebrafish embryo, on the other hand, has its full set of 30 pairs of somites, its heart is beating, and pigmentation is forming in the eye and skin (Figure 15.3A). Somites The first pair of somites form just after 10 hours of development, and from then on, a new pair of somites forms about every half-hour. This means that within the space of a few hours, you should be able to observe several sets of somites forming (Figures 15.2D–E). Neural tube As the somites are forming, neurulation is also occurring. The neural plate is apparent by 10 hours of development, just as the first pair of somites is forming (bud stage). The neural plate then folds inward forming first a neural keel (13 hours), and then a neural rod (16 hours). This rod secondarily cavitates to form the hollow neural tube (beginning at 18 hours when the embryo has 18 somites). Regions of the brain can now begin to be identified as distinct swellings called neuromeres. By 18 hours, there should be about 10 neuromeres; in anterior-to-posterior order, the first three neuromeres correspond to the telencephalon, diencephalon, mesencephalon (midbrain), and the last seven neuromeres (rhombomeres) correspond to the rhombencephalon, or hindbrain (Figure 15.2F). Sensory placodes of the head Various sensory structures of the head can also be identified by 18 hours (18-somite stage; Figure 15.2F). Look for the olfactory placodes anterior to the telencephalon, prominent optic (eye) cups forming as outpocketings of the diencephalon and their associated lens placodes, and the otic (ear) placodes forming alongside rhombomere 5 of the hindbrain. Pharyngeal arches Following 24 hours, the embryo begins to respond to touch, and pigmentation becomes pronounced. Within the pharyngeal region, the jaws and operculum are forming from the first two pharyngeal arches, and the more posterior pharyngeal arches (also called branchial arches that will form the gills) are becoming apparent. Fin fold and fin buds Also at 24 hours, the developing tail fin primordium (median fin fold) surrounding the tail bud can be identified (Figure 15.3A). The pectoral fin buds begin their development later, being identifiable as small mounds at 30 hours of development (prim-15 stage) and becoming more prominent by 48 hours (long-pec stage; Figure 15.3B).

Zebrafish Development (A)

(B) Lens

Otic placode

Olfactory placode

Optic cup Lens Otic placode

Yolk

Optic cup

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Heart

Pectoral fin

Olfactory placode Heart

Somites

Yolk Somites

Opening of pronephric duct

Opening of pronephric duct

Median fin fold Tail fin Prim-6 25 hours

Long pec 48 hours

Figure 15.3 Zebrafish development continued. (A) Prim-6 stage. By this stage, the median fin fold has formed, the heart is beating, and the embryo is sensitive to touch. (B) Long pec stage. The embryo hatches between 48 and 72 hours of development. By 48 hours, the pectoral fin buds are visible.

Lateral line Among the structures that will form in the zebrafish that you did not observe in the chick is the lateral line system. This system, which consists of a series of canals containing sensory structures called neuromasts, is designed to detect low-frequency motions within the water, and thereby inform the fish about such things as prey and predators within close range. The lateral line begins to form as a placode in the epidermis just posterior to the otic placode. By the 18-somite stage, it begins a posterior migration within the epidermis, migrating the distance of two somites every hour, until it reaches the tail at about 40 hours of development. Hatching Between 48 and 72 hours, the embryo hatches. The cartilaginous skeleton of the pharyngeal region is being laid down, and the larva is beautifully colored with the yellow sheen from its iridophores.

Preparing For Your Laboratory Studies Zebrafish are easily maintained in the laboratory and require no more than the facilities that you might have at home if you keep fish in a freshwater aquarium. Unlike at home, however, in the laboratory strict regulations have been established for the handling and care of vertebrate animals. Check with your institution’s animal care committee (IACUC) for the necessary training and forms before obtaining your zebrafish.

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Culture procedures Tanks and water You do not need a large tank to maintain these fish. A 10-gallon aquarium is sufficient for approximately 25 to 50 fish, and smaller tanks can be used for fewer fish. The water used can be tap water, as long as it has been aged for at least a day to release any chlorine in it. Alternatively, distilled water can be used if a small amount of salts are added (60 mg instant ocean/liter). The water should be kept between 26 and 28.5ºC (78–83ºF). The tanks should be cleaned by siphoning out debris and replacing a third of the water each day. Water quality can be tested using aquarium testing kits that measure pH, and ammonia and nitrite levels. The water should be filtered using a standard tank-filter. Also, adding plants to the tank, either artificial or living, will provide shelter for the fish (a living plant such as the Java fern, Microsorium pteropus, will also improve the water quality, though it can introduce unwanted parasites as well; artificial plastic plants will avoid this possibility.) Because zebrafish are photoperiodic in their breeding, they will only breed when kept on a specific light/dark cycle; 14-hour light/10-hour dark cycle. The fluorescent fish-tank lights available in pet stores plugged into an automatic timer switch work well for this. If you want to obtain a large number of fertilized eggs on a specific day, maintain males and females separately until the night before you will need embryos. If you want a steady, daily supply of eggs, or a system requiring less maintenance, you can keep males and females together in the same tank. This allows the adults to establish a more normal social hierarchy and is less disruptive to the fish prior to mating. Placing opaque paper on the outside of all but one wall of the tank will limit distractions and help to keep the fish calm. Sexing Adult Fish Sexing the adult fish is not difficult, though you do have to look closely to see differences between the sexes. Males are longer, slimmer, and more yellow, especially on the belly. Females are plumper and more silvery. Feeding For general maintenance, adults should be fed once or twice a day. Food can be ground up dry or moist trout pellets (e.g., Ranger® quarter inch brood food) or dry flake food (e.g., Tetra®). Provide enough food so that all fish are fed and the food is eaten within 5 minutes. Prior to breeding, adults (aged 7 months to 1.5 years is an optimal) should be fed a rich, highprotein diet, three times a day. Among the best food sources is brine shrimp (Artemia; live or frozen, though live is best). Brine shrimp can be easily raised from eggs; put approximately 10 ml of shrimp eggs into 2000 ml of sea water or instant ocean 2000 ml. Keep this well aerated at 28.5ºC. After 48 hours, the baby shrimp can be used as food: filter the shrimp through a fine-meshed cloth or a commercially available net made for this purpose, washing the shrimp with fresh water. Dilute the shrimp in fresh water (1:3; shrimp to water). A Pasteur pipette’s worth of this mixture feeds approximately eight adult zebrafish. Other food sources include Daphnia, Drosophila (adults or larvae), or frozen “white worm” (Chaoborus; the larval stage of this dipteran insect which are also called glass worms). Live food should be alternated with flake food. Do not overfeed, since this will reduce egg Obtaining embryos laying. Provide the amount of food that will be eaten within five minutes. day before the eggs are males needed; four males in and eight females usually ensurefertilized good results. you have If you are maintaining and females separate tanks, then to obtain eggs on(Ifa specific been males and together females together in the same this tank is also breeding tank, day, maintaining put males and females in a “breeding tank”tank, in a ratio of one malethe to two females onand the a

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system for collecting eggs, as described below, should be placed in the tank the night before you will be collecting eggs.) A breeding tank can be created in a number of different ways. It can be small (as small as 4”× 6” for 3–6 fish; 18” × 18” for 12–20 fish), and made of plastic or glass. The key element is any device that will keep the adults away from the spawned eggs, so that the eggs won’t be eaten after they are laid. A simple method for doing this is to place a glass baking-dish, containing a single layer of marbles, into the tank on the night before eggs are to be collected. Place plastic plants in the bakingdish as well; the fish will swim in and out of these plants while spawning. Alternatively, a nylon mesh can be used secured within the tank to separate the adults from the bottom; the holes in the mesh should be 3–4 mm in diameter. Mating usually occurs within an hour of “dawn,” the beginning of their light period. Once the eggs have been laid, they can be removed from the tank. If you used a baking-dish containing marbles and plants to collect the eggs, remove the plants and carefully lift the dish out of the tank. Pour the water from the dish into a large beaker; then remove the marbles from the dish, and wash any further eggs left in the dish into the beaker. The eggs will settle to the bottom of the beaker and can be collected with a pipette (opening at tip should be 2 mm or larger) or a flexible hose with a piece of glass tubing at its end (6 inches long with a 1 cm internal diameter works well). Alternatively, you can pour the water through a tea strainer, and the eggs can then be washed from the strainer into a Petri dish. If a nylon mesh was used in the tank instead of marbles, you can siphon the eggs off the bottom of the tank using a pipette or glass tubing as described above. (If the fish are reluctant to spawn at the beginning of the light cycle, changing the water can help to stimulate them to do so.) If you are maintaining the males and females separately, return the males and females to their separate tanks following mating. If a breeding pair does not produce eggs on the first day, you can leave them together for a second day. After this, the pair should be allowed to rest for a week before being used for breeding again. Maintaining embryos For normal development, embryos and young larvae should be maintained at 28.5ºC in a balanced salt solution (egg water). Egg water can be made easily using Instant Ocean. Alternatively, Hank’s solution diluted to 10% of its original strength can be used. Egg Water* Instant Ocean Sterile distilled H2O

60 mg 1 liter

Hank’s stock solution* (Alternative to Egg Water) Solution 1 NaCl KCl CaCl2 Na2HPO 4 KH 2PO4 MgSO Solution 2 2O 4·7H Sterile distilled H 2O NaHCO 3

0.8 gm 0.04 gm 0.014 gm 0.0036 gm 0.006 gm 0.025 gm 90 0.035 ml gm

To make stock solution, mix each solution separately. Make solution 2 just prior to use and add to solution 1. For use, dilute the stock solution to 10% of its original concentration by adding 10 ml of Hank’s stock solution to 90 ml sterile distilled water. Store solutions in the refrigerator.

Zebrafish Development Solution 2 NaHCO3 Sterile distilled H2O

15 - 12

0.035 gm 10 ml

*N.B.: These solutions cannot be autoclaved since autoclaving causes the precipitation of calcium carbonate. Use sterile distilled water and sterile containers, or, alternatively, filter the solution through a sterilizing filter apparatus prior to use. Raising larvae The zebrafish embryos will hatch in approximately three days, but they do not have to be fed until the fifth day. Larvae should be maintained separately from adults, since the adult fish will eat the fry. Initially, you can keep the embryos and larvae in Petri dishes or small beakers using egg water (approximately 15 embryos/100 ml water). Change the water daily and clean out any dead or dying embryos. After hatching, the larvae can be switched to the water used for adults. During the first day after hatching, the larvae will lie on the bottom of the dish—don’t worry, their swim bladders haven’t yet filled with air. Feed larvae baby fish food (available at pet stores) or paramecia until they are nine days old. On the tenth day, move the larvae to larger containers and begin feeding them brine shrimp. By three months, they are sexually mature and can be kept with the adult fish and used for breeding.

Experiments Using Zebrafish Embryos You will be designing your own experiments; what follows are only suggestions. Remember, the major focus of these experiments is to discover possible teratogens in our environment and the effects of these on development. When you start an experiment, state clearly in the “Experimental Set-Up” section of your laboratory notebook the question or questions you are asking. Observe normal development with as much attention to detail as possible so that you can identify any abnormal development in your experimental cultures. Observing normal development Obtain embryos, either as instructed above or as provided by your instructor. Sterile procedure should be used. Use a sterile Petri dish containing egg water as a culture chamber. Devise a scheme for maintaining your culture at 28.5ºC. If your laboratory has a humidified culture chamber that can be set to specific temperatures, this is ideal, but other, less expensive devices can be utilized. A simple method is to place an incandescent lamp at a distance from the dish that is sufficient to keep the culture at the proper temperature—use a thermometer to measure the temperature. Alternatively, you can use a slide warming tray set at very low heat (again, you will need a thermometer to measure the temperature). When observing the embryos under the microscope, make sure that you provide enough heat to keep the embryos at the proper temperature. You may find that the light source for the microscope is sufficient. If not, use an additional incandescent lamp set at a short distance from the stage of the microscope. Use a thermometer to measure the temperature. Observe the embryos first under your dissecting microscope. Use its highest power (these embryos are small). To observe an embryo under the compound microscope, place it in the well of a depression slide and cover with a coverslip. If the well of the slide is not deep enough, or if you don’t have a depression slide, use a footed coverslip, making sure the spacers are thick enough so that the embryo isn’t crushed. (In this case, using small globs of vacuum silicone grease at the corners of the coverslip

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works well. After slipping the embryo underneath the coverslip, you can gently push down on the coverslip until the embryo is held in place.) Another method is to create a spacer using coverslips. Four 22–mm2 coverslips, glued together, gives an adequate amount of height for an embryo with its chorion intact. Take two of these multi-coverslip preparations, and glue them onto a glass slide spaced about 20 mm apart from one another. After placing the embryo between the two stacks, a longer coverslip (22 mm × 60 mm works well) can be placed on top. Observe normal development over a three-day period, making as many observations as time allows. Change the medium daily and clean out any dead or dying embryos. Following 24 hours, you may want to anesthetize the embryos for observation. Use a stock solution of tricaine (4 mg/ml dH20, stored at 4ºC) diluted 1:10 in embryo water. Using the description of development given above, identify as many of the structures and embryonic processes shown in boldface type as you are able. For each observation, record the time of observation, the temperature of the culture and the stage of the embryo, using the staging series in Table 15.1. And most of all, appreciate what the embryo is showing you—it is a phenomenal amount of development in a short space of time. Having observed normal development, construct a graph or chart of the normal developmental rate. You will be using these data to compare the development of your experimental cultures with normal development. Removing chorion For certain observations, it is convenient to have removed the chorion from around the embryo. Removing the chorion can also increase the exposure of the embryo to certain teratogens. Dechorionating an embryo can be done relatively easily using fine forceps. Using a dissecting microscope to see what you are doing, grasp the chorion with two pairs of fine forceps and make a tear large enough for the embryo to fall out. Pre-treating the embryo with dilute pronase (2 mg/ml embryo water) makes the chorion brittle and easier to remove. Treat for one minute (longer for older embryos); rinse 3–4 times in embryo water. Often the swishing alone during washing will cause the chorion to fall off. (Alternatively, a weaker solution of 0.5 mg pronase/ml embryo water for 3.5 minutes can be used.) Once dechorionated, the embryo is very fragile, especially in the early stages (prior to 3 hours of development). Normally, you should not dechorionate an egg prior to the mid-blastula stage. Following dechorionation, an embryo younger than 24 hours of development should be kept in an agar-coated Petri dish to avoid its sticking to the dish. Temperature experiments A number of experiments can be done using temperature as a variable. Embryos often have a narrow range of temperatures that they will tolerate. You can be asking the question, what is the temperature range for the zebrafish embryo, what defects occur outside of this range, and is a temperature that is too cold less or more harmful than one that is too warm. In your experiments, you can reflect on the effects of environmental temperature changes on the development of organisms. From your results, what would you conclude about such environmental conditions as a nuclear power plant dumping super heated water directly into a waterway? The threat of global warming? The effects of a long hot bath on the fetus of pregnant mother? (Long hot baths are a threat to the development of a human embryo—do your studies support this finding?) In your experiments, remember to watch for early effects of temperature changes on the embryo. Cold temperatures (18ºC) inhibit microtubules. If you use this temperature, does it inhibit the initial ooplasmic segregation that brings the yolk-free cytoplasm to the top of the egg? Does it inhibit epiboly? Does it inhibit axis formation? A temperature this low is undoubtedly very harmful to the

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embryo and it may not survive unless the low temperature is used for only short periods of time. With short exposures at different time points, you can determine which stages are especially sensitive. Variable lengths of treatment will give you information about levels of tolerance. A heat shock can be delivered by exposing the embryos to a temperature above 40ºC (41.4ºC is suggested). A two-minute exposure delivered 13 minutes after fertilization is sufficient to prevent the first mitotic division, and diploids become tetraploids. Remember, in these and all experiments, it is imperative to compare your experimental cultures to control cultures, since even under control conditions embryos can display defects. For temperature experiments, the control cultures would be embryos raised at a constant 28.5ºC. Chemical teratogens Only recently has it been adequately recognized that the substances ingested by a pregnant woman can cause defects in her unborn child. Up until the 1950s, and even later in some cases, potent teratogens such as mercury, thalidomide, diethylstilbestrol, alcohol, and vitamin A, to name a few, were not considered to be dangerous to human development. Now, with environmental pollutants accumulating at an unprecedented rate, and pharmaceuticals dominating western medicine, it is particularly important that we document teratogenicity of the substances that we are exposing ourselves to. To determine the risk to human fetuses, the testing of substances on non-human embryos is an important step, though it is not fool-proof. Different species show different susceptibilities to teratogens (the story of thalidomide is a tragic example—of all the organisms tested, humans have turned out to be the most sensitive to this teratogen, but it was tests on other species that led us to believe that thalidomide, taken as a sleep-aid and antinausient, would be harmless to human fetuses). You can use the zebrafish embryo to test a number of substances suspected of teratogenicity. To maintain safety in the laboratory, we will not be using substances that are considered hazardous materials (such as mercury), but there are a number of other substances that people ingest on a regular basis that can be tested. Lithium Lithium, in the form of lithium carbonate, is regularly prescribed to people with manic depressive (bipolar) disorder. It takes the peaks and valleys out of the emotional swings that a person with this disease suffers1, and is thought to do so through its control of receptors for the neurotransmitter glutamate. Even though embryologists have known since the late 1800s from studies on non-human embryos that lithium causes severe developmental defects, it is still disputed as to whether prescribed doses of lithium pose a threat to human embryos. Those who attribute teratogenic effects to these doses say that embryos are at increased risk of heart defects, especially Ebstein anomaly which affects the tricuspid valve and seriously compromises the heart’s ability to function effectively. Other defects are also reported, such as spina bifida and “floppy infant” syndrome, in which the baby is weak and unresponsive. Other reports contend that the risks have been overstated. In studies on non-human embryos, the developmental defects caused by lithium are clear. As early as 1892 (Herbst, 1892), it had been shown that echinoderm embryos exposed to lithium salts undergo exo-gastrulation. More recently, it has been shown that both Xenopus and zebrafish embryos exposed to lithium during cleavage stages are dorsalized, showing severe defects in the posterior region. (It is βthought that these defects are caused by lithium’s inhibition of glycogen synthase kinase in the Wnt/β 1

Known, actually, as early as 200 AD, when Galen prescribed alkaline spring baths for manic depressive illness.

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catenin pathway or by depletion of inositol triphosphate in the PIP cycle as the dorsal-ventral axis is being established; Klein and Melton, 1996; Stachel et al., 1993; Baraban, 1994.) In zebrafish, later exposure (at the eleventh cleavage through gastrulation) causes anterior defects, especially of the eyes, in some cases causing a lack of eyes all together. The doses that have been used to create these developmental defects in experimentally exposed zebrafish embryos are higher than the levels of lithium in the blood of a patient prescribed lithium. Typically, an embryo is exposed to 0.3M LiCl (more than 100 × the level in a patient’s blood serum). Also, the experimental exposure times used in past experiments have been brief (e.g., 10 minutes), and are not at all analogous to the constant levels a human embryo would be exposed to. It would be very interesting, therefore, to expose zebrafish embryos to levels of lithium akin to the levels a human embryo might experience. A therapeutic dose of lithium carbonate is 300–1500 mg/day, to attain a blood serum concentration of 0.7–2.0 mEq/L. For us to achieve this level in the laboratory, we would use a 0.7–2.0 mM LiCl solution (N.B.: 42 mg LiCl/liter egg water = 1 mM solution). To simulate the exposure a human embryo might experience, expose the zebrafish embryos to a constant low dose of lithium salts within the range given above. To determine which developmental periods are most sensitive, expose the embryos at varying time points. To determine the level at which a constant exposure to LiCl becomes teratogenic, use various concentrations that differ by 10-fold (1 mM, 10 mM, and 100 mM, for example). Vitamin A Vitamin A is another teratogen that is commonly available. Vitamin A is a necessary component in anyone’s diet, supplied through fish oil, liver, egg yolk, dairy products and as beta-carotene in yellow, orange, red and leafy green vegetables. As long as it is supplied in its precursor form, beta-carotene, it poses no threat to the health of either the person ingesting the beta-carotene or a developing embryo—quite the opposite, it provides great health benefits to both. Megadoses, taken as vitamin A, however can be harmful, even lethal to a person (no more than 5000 IU/daily is recommended), and have known teratogenic effects on a human embryo (vitamin A supplements should be avoided when pregnant—just eat your veggies). The recent use of isotretinoin (Accutane®), a form of vitamin A used for the treatment of severe acne, poses a particular threat to human embryos. Taken orally, it is so highly teratogenic that any woman of reproductive age prescribed isotretinoin (13-cis-retinoic acid, Accutane®) is asked to use two reliable forms of birth control. It is no surprise that vitamin A is teratogenic. One of the derivatives of vitamin A, retinoic acid (RA), is a posteriorizing signal in many developing systems and is known to influence Hox gene expression. Treatment with RA can cause a respecification of the anterior-posterior identity of regions, shifting them in a posterior direction. Birth defects attributable to RA include defects in the central nervous system such as microcephaly (small brain) and microphthalmia (small eye), craniofacial defects primarily due to underdevelopment of structures, defects in the cardiovascular system such as the tetralogy of Fallot, defects in the thymus, and reduction of the limbs. The zebrafish embryo can be used as a sensitive monitor of retinoic acid contaminations. It is already known that retinoic acid interferes with zebrafish development at very low concentrations; as low as 10 -7 M can cause microcephalic and micro-ophthalmic development, even to the point of the embryo lacking a head and eyes. The fact that certain pesticides can mimic retinoic acid adds to the significance of your investigations. (Methoprene, for example, which is commonly used as an insecticide because it mimics juvenile hormone in insects, breaks down when exposed to sunlight to form a product that acts like retinoic acid. Sold under a number of trade names such as Altosid®,

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Precor®, Kabat®, Pharorid®, Dianex®, Apex®, Fleatrol®, Ovitrol®, Extinguish® and Diacon®, it is often used by pet owners to control fleas; it is also put in cattlefeed to control horn flies, and sprayed on wetlands to control mosquitoes.)

Table 15.2 A short primer on Vitamin A, its precursors and derivatives

Name

Alternate name

Source

Action

Toxicity/ Teratogenicity

Molecular Weight

Beta-carotene

Provitamin A

Green leafy vegetables, yellow-to-red vegetables

Precursor to vitamin A

None known

MW

Vitamin A

Retinol

Liver, fish oil, dairy products

Precursor to two active forms of vitamin A, retinal and retinoic acid

Considered teratogenic in ingested doses greater than 15,000IU/day (no higher than 2700 IU/day is recommended for pregnant women)

286.46

Toxic to adults in doses above 100,000 IU/day Retinal

vitamin A aldehyde

Derived from vitamin A

Pigments in eye for vision.

Not ingested.

Retinoic acid

Vitamin A acid

Derived from vitamin A

Signaling molecule in development etc.

Highly teratogenic. See entries for all-trans and 13cis-retinoic acid below.

All-trans retinoic acid

Tretinoin (Retin-A)

Synthetic form of retinoic acid

Used topically for superficial pustular acne

Highly teratogenic to embryos in laboratory studies.

300.4

When used topically on skin, not considered teratogenic 13-cis-retinoic acid

Isotretinoin (Accutane®)

Synthetic form of retinoic acid

Used orally for severe cystic acne

Highly teratogenic (not to be taken during or prior to pregnancy). Numerous possible toxic side-effects in adults.

300.4

You can test the effects of vitamin A and retinoic acid on zebrafish development by exposing the embryos to varying levels. If you are concerned about teratogenesis in humans, then it would be appropriate to test the levels that are attained within the blood of the mother. For example, when people are treated with isotretinoin (Accutane®), their blood concentrations of isotretinoin range from 0.16–0.46 µg/ml; this is roughly equivalent to 0.5–1.5 µM (0.5–1.5 10 -6 M). It would therefore be interesting to test concentrations similar to this, using retinoic acid (either all trans, which is equivalent

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to tretinoin, or 13-cis which is equivalent to isotretinoin). By testing concentrations that vary by 10fold, for example 10-6, 10-7, 10-8 M, etc., you may be able to determine a tolerance level. Relevant levels of vitamin A to test would range from 0.2–20 µg/ml. A person’s normal blood serum levels of vitamin A are 0.2–0.5 µg/ml. When a person ingests a toxic dose of vitamin A causing chronic toxicity (100,000 IU/day), their blood levels rise to 1–20 µg/ml. Vitamin A is more difficult to test since it is oil soluble, though there is a water-soluble form available (retinol acetate-water soluble from Sigma, for example). Alcohol Alcohol, in the form of ethanol (ETOH; the form found in alcoholic drinks) is a significant threat to human embryos, capable of causing abnormalities even at low levels. Fetal alcohol syndrome (a wellrecognized collection of defects including facial abnormalities, heart defects, and mental retardation) can be caused by 1–2 alcoholic drinks/day consumed by the mother during pregnancy; fetal alcohol effect (a milder condition exhibiting primarily a slowness in learning) can be caused by much lower levels of drinking. It is now considered that no level of alcohol is safe for the fetus, and that pregnant women simply should not drink—at all. Again, the zebrafish embryo can serve as a useful model in examining the teratogenic effects of alcohol. It has already been shown that zebrafish embryos treated with ethanol (the form of alcohol that is consumed) during various stages can develop abnormalities such as notochord and spinal cord duplications and cyclopia (a condition where only one eye develops). To test levels of ethanol that are relevant to human pregnancies, it is important to know the blood levels of alcohol that occur within a mother. (A person with blood-alcohol levels of 0.1% is considered legally drunk, and levels of 0.05–0.09% constitute being “under the influence.”) You could test various levels (0.1%, 1% and 3% ethanol in egg water, for example) for varying periods of time. You could also compare constant exposure with episodic exposure. (Recognize that a human fetus is unlikely to be exposed to a constant level of alcohol. The mother has liver enzymes [alcohol dehydrogenase] that break down alcohol in a matter of hours; the embryo, lacking this enzyme, is exposed to alcohol that enters the amniotic fluid for far longer periods of time.) Endocrine disruptors It has recently become apparent that a number of the pollutants we dump into our environment are endocrine disruptors, many of them acting as estrogens or anti-estrogens. Dioxin, a by-product of making pesticides and paper, and DDT, used so widely in insect control, are particularly hazardous examples. Some of the chemicals used to harden plastics, such as p-nonylphenol, are estrogenic and enter our systems through plumbing and even through the polystyrene plastics that hold our milk and orange juice. The phthalates used to make plastics flexible and that are added to cosmetics are endocrine disruptors that have been linked to premature sexual maturation in young girls and disruption of maturation in boys. You could use zebrafish embryos to test the teratogenic effects of some of the plastics you use by first storing egg water in the containers for a period of time and then growing embryos in this solution. Since an acidic solution might draw more compounds out of the plastic than a neutral solution, you could acidify distilled water to something comparable to orange juice (pH 3.3–4.2) or soda (pH 2.5–3), and then neutralize the water prior to using it to make egg water for your cultures. You can use HCl to make solutions of various acidities. HCl is sold in concentrated form as a 12.1 M solution. To make a 0.1 M HCl, add 8.26 ml of 12.1 M HCl to enough distilled H 2O to make up one liter. Then dilute this solution to make the pH you want to test.

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0.01 M HCl = pH 1 0.01 M HCl = pH 2 0.001 M HCl = pH 3 0.0001 M HCl = pH 4 After storing your acidic solution in the plastic container you are testing, titrate the solution back to pH 7 with a 1 M solution of NaOH prior to using it for making egg water. Testing the water below the pipe You can also use zebrafish embryos to test the quality of water systems in your area. By growing the embryos in water collected from various sources in your area (e.g., a pond that collects agricultural runoff, a stream or river below an effluent pipe from a paper company), you can make conclusions about the possible teratogenic effects of this water on other aquatic organisms. Behavioral studies Some of the effects of a teratogen might be subtle and apparent only through behavioral studies. To determine these effects, you would first want to establish what the normal behaviors are and then look for variations in these. Simply discovering what behaviors are normal and the developmental time at which these are apparant is a project in itself. Some of the behaviors you might be looking for are reactions to a touch stimulus, phototactic or negative phototactic responses, mating behaviors, and the setting up of dominance hierarchies. For example, embryos start showing spontaneous contractions at about the 20-somite stage. Is the embryo sensitive to touch at this point? If not, when does the touch response develop and do spontaneous contractions still occur? Once hatched, do larvae show any phototactic or negative-phototactic behavior? Adults show negative phototaxis, exhibiting a preference for a dark, shaded tank over a bright open one. By using a small aquarium that has one side shielded with dark paper, you can test the adult response as well as the response in larvae to determine the age at which the behavior becomes apparent. In a population of zebrafish, the fish quickly establish dominance hierarchies through intense stereotypic fighting activity that includes lateral displays, frontal displays, nipping, and chasing. This lasts only a few minutes and ends with the subordinate fish retreating. By using pairs of fish in a small tank, you could ask whether these behaviors are demonstrated by both sexes equally. Do males and females fight? Is the dominant fish always a male? Always a female? Do females fight other females? After mating, it is advised that you remove the eggs from the tank since the adult fish eat the eggs. This seems an odd behavior, one that would jeopardize survival, and you could investigate it further. Do males and females both devour the newly spawned eggs? How many do they eat? Will they deplete the entire lot of eggs that they have spawned if given the chance?

Completing your experiments Watch your cultures carefully through out the rest of the laboratory period—a lot can happen in only a few hours—and check your cultures throughout the next three days as time allows. Change the culture fluid daily, and remove and make a record of any dead or dying embryos. Make comparisons between the experimental cultures and control cultures of normal development, including survival rates. Keep

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careful records of the timing of events and use the staging series to identify stages of development at each time point that you make an observation. Always record the time of your observation and the water temperature. Make graphs of developmental rates of experimental cultures as compared to normal cultures. Use diagrams to document what you see. A great deal of data will be tied up in these diagrams, so make them accurate and proportional. Watch for what the embryo is telling you. And remember that your results may be significant enough for eventual publication, since many of the suggestions given and the ideas that you’ll be coming up with on your own are a venture into virgin territory. You may want to fix your embryos or larvae to examine them as whole mounts. The procedure for this takes a number of hours to complete, but each step can be done relatively quickly.

Preparation of whole mounts Typically, there are two types of methods used in making whole mounts, one using a water-soluble and the other using a non-water soluble mountant. Described below is a procedure for a water-soluble mountant, which is faster, but creates a less permanent preparation than a non-water soluble mountant procedure (see Chapter 11 The Living Embryo and Making of Whole Mounts for a non-water mountant procedure). There are a number of staining procedures that can be used. The alcian blue/alizarin red method below is specifically for staining cartilage and bone, though it also gives a general pink background staining to all tissues; alcian blue at a low pH stains the glycosaminoglycans of cartilage, and alizarin red at a high pH stains the calcium of bone. Often you will be looking for skeletal defects in the larvae that you have treated, and this staining method can help you to detect these defects. Remember that cartilage and bone don’t begin to form until the hatching period (the first cartilage is apparent in the long-pec stage, around 48 hours of development, and the first bone in the protrudingmouth stage, around 72 hours of development). If you are fixing larvae, the larvae should be anesthetized before fixing. You can use 0.1 mg Tricaine dissolved in egg water or tank water. Fixation Fixation causes the cross-linking of proteins and prevents degradation of the tissues. One of the most common fixatives is formalin, and a good fixative for zebrafish embryos and larvae is neutral buffered formalin. The rule of thumb is to fix the specimen in an amount of fixative equivalent to ten times the volume of the specimen (realize, this isn’t a lot of fixative, usually enough to cover the specimen). If you are processing embryos that still have intact chorions, the chorions should be removed prior to fixation. Use fine forceps to carefully tear the chorion away from the embryo. Fix your specimens for 2 hours at room temperature or 8 hours (overnight) in the cold (4ºC; refrigerator temperature). Any fixative containing formalin (an aqueous solution of formaldehyde) is highly toxic to living embryos, and any glassware or tool that is used for fixation can not be used for living material. Neutral buffered formalin Formalin Distilled water NaH2PO4·H2O Na2HPO 4

100 ml 900 ml 4 gm 6.5 gm

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Washing After fixation, it is important to remove the fixative from the tissues by washing. This can be done by washing the specimens in 70% ethanol for 1 hour. Specimens can be stored in 70% ethanol if needed. Staining procedure for cartilage and bone The embryos and larvae can easily be damaged during the many transfers that must be made during the staining procedure. An effective way to minimize this damage is to transfer the embryos and larvae to depressions slides after washing. When a change of solution is required, use a Pasteur pipette to remove the solution and to replace it with the next. The staining schedule is as follows: • Stain the specimen in alcian blue for 2 hours. • Wash the specimen for 10 minutes in 70% ETOH to wash out the excess stain. • Stain in alizarin red for 2 hours. • Wash the specimen in water for 2 hours. Alcian blue solution Alcian blue 15 mg 95% ethanol 80 ml Glacial acetic acid 20 ml

Combine ingredients, adding the glacial acetic acid to the ethanol, rather than the ethanol to the acid. Filter before use.

Alizarin red solution Alizarin red S 0.5% KOH

10 mg 100 ml

Combine ingredients. Filter if necessary.

Clearing and mounting Following staining, the specimen is to be cleared in a solution that makes the tissues translucent. Glycerin is a clearing agent that is miscible in water. There are two methods you can use: the timesaving method, or the effort-saving method. Again, it is best to keep the specimen in the depression of a depression slide and to remove and add solutions using a Pasteur pipette. Put the specimen into a 20% glycerin solution diluted with 70% ETOH, and allow the alcohol to slowly evaporate. If the specimen is in the depression of a depression slide, put the slide in a glass Petri dish with its cover on to keep dirt out. Make sure that you start out with enough volume of glycerin/ETOH solution so that as the alcohol evaporates there is enough glycerin to cover the embryo. Add additional 100% glycerin as needed. After three days, replace the solution with 100% glycerin. Your specimens can be stored in 100% glycerin; add crumb of thymol to prevent mold. It is often best to store the specimens in this manner rather than making a permanent slide since this allows you to manipulate the embryo while studying it under the microscope and therefore to see it from all angles. If you choose instead to mount the specimen on a slide, place the specimen on a slide and surround it with glycerin jelly that has been melted in a warm-water bath. The glycerin jelly is the permanent mounting medium. Now place a footed coverslip over the specimen. Make sure the spacers you use are large enough to lift the coverslip at least as high as the specimen is thick. If there is not enough mounting medium to fill the space under the coverslip, use a pipette to add more glycerin jelly from the side. If there is too much glycerin jelly, leave it until the next lab when it will be thoroughly solid. Then you can remove the excess by cutting it away with a razor blade.

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Kaiser’s glycerin jelly Distilled water Gelatin Glycerin Thymol

52 ml 8 gm 50 ml 1 crystal (to prevent mold)

Heat to 75°C and stir until dissolved. Do not heat above this temperature.

Put your slide face-up in a covered box until next week. At that time, use fingernail polish to ring the edge of the coverslip, making an airtight seal. This will prevent the glycerin jelly from drying out.

Accompanying Materials See Vade Mecum: “Zebrafish.” This chapter of the CD shows, in movies, time-lapse photography, and still pictures, zebrafish development and techniques used to set up cultures of the embryos. Gilbert, S. F. 2003. Developmental Biology, 7th Ed. Sinauer Associates, Sunderland, MA. Contains an excellent section on early zebrafish development. Fink, R. (ed.). 1991. A Dozen Eggs: Time-Lapse Microscopy of Normal Development. Sinauer Associates, Sunderland, MA. Sequences 9 and 10 show zebrafish development and high-magnification timelapse of deep cell movement in the killifish.

Selected Bibliography Akimenko, M.-A. and M. Ekker. 1995. Anterior duplication of the Sonic hedgehog expression pattern in the pectoral fin buds of zebrafish treated with retinoic acid. Dev. Bio. 179: 243–247. This paper demonstrates that short, 2 hour exposures to retinoic acid (all-trans RA; 10-6 M) can induce ectopic expression of the Sonic hedgehog gene in the paired fin buds of the zebrafish, which results in abnormal fin development. Baraban, J. M. 1994. Toward a crystal-clear view of lithium’s site of action. Proc. Natl. Acad. Sci. USA 91: 5738–5739. This is a short, but excellent review of the dangers of therapeutic levels of lithium, a treatment for manic depression. It also discusses what is known about the site of lithium activity, proposed to be inositol monophosphatase, an enzyme in the PIP cycle. Blader, P. and U. Strähle. 1998. Ethanol impairs migration of prechordal plate in the zebrafish embryo. Dev. Bio. 201: 185–201. This paper discusses the effects of alcohol on development, showing that treatment during the early gastrula stage of zebrafish development can cause cyclopia. It also discusses possible mechanisms for this action of alcohol on development. Conlon, A. R. 1995. Retinoic acid and pattern formation in vertebrates. Trends Genet. 11: 314–319. This paper reviews the evidence for suggesting that retinoic acid plays an important role in the patterning of the anterior-posterior axis in the vertebrate embryo. Currie, P. D. 1996. Zebrafish genetics: mutant cornucopia. Curr. Biol. 6: 1548–1552. This article reviews the accomplishments of the large mutagenic screen that identified numerous mutations affecting development in the zebrafish. Detrich, H. W. III, M. Westerfield and L. I. Zon (eds.). 1999. Methods in Cell Biology, Volume 59. The Zebrafish: Biology. Academic Press, San Diego, CA. This valuable book contains excellent background information on zebrafish development and serves as a resource on laboratory methods, including embryo culture, cell culture, and histological techniques.

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Detrich, H. W. III, M. Westerfield and L. I. Zon (eds.). 1999. Methods in Cell Biology, Volume 60. The Zebrafish: Genetics and Genomics. Academic Press, San Diego, CA. This is the second volume in the Methods in Cell Biology series that covers methods in zebrafish research. This volume reviews genetic methods, and includes a number of useful appendices on genetic stocks, collection of sperm, in vitro fertilization, and relevant websites. Development. Volume 123. December, 1996. This entire special issue (37 articles) of the journal Development is devoted to the description of approximately 2000 mutations that disrupt development of zebrafish; it is the result of two independent large-scale mutagenic screens of zebrafish genome and phenotypic analysis. The two investigators leading the screens were Christiane Nüsslein-Volhard and Wolfgang Driever. Christiane Nüsslein-Volhard received the Nobel Prize in Physiology or Medicine (1995) along with Eric Wieschaus and Edward Lewis for their work on Drosophila. Eichele, G. 1993. Retinoids in embryologic development. Ann. New York Acad. Sci. 678: 22–36. This paper provides a concise review of the effects of retinoic acid on the development of the vertebrate limb, the anterior-posterior body axis, and the central nervous system. Eisen, J. S. 1996. Zebrafish make a big splash. Cell 87: 969–977. This review paper provides an excellent summary of zebrafish development and the mutations that affect this development. Grunwald, D. J. 1996. A fin-de-siècle achievement: charting new waters in vertebrate biology. Science 274: 1634–1635. This is a useful review of the mutagenic screens that were done on zebrafish and reported in the 1996 issue of Development, volume 123. The article explains the significance of this accomplishment and the impact it will have on developmental studies. Guille, M. (ed.). 1999. Molecular Methods in Developmental Biology, Xenopus and Zebrafish. Humana Press, Totowa, NJ. This volume outlines challenging techniques that can be used on zebrafish embryos, including cell and tissue transplantation, whole mount in situ hybridization, immunohistochemistry, and microinjection. Hammerschmidt, M., et al. 1996. dino and mercedes, two genes regulating dorsal development in the zebrafish embryo. Development 123: 95–102. In this article, the authors describe mutations that affect the dorsal-ventral patterning of the zebrafish embryo. Color photographs show the identifying characteristics of the mutants. Hennig, W. (ed.). 1992. Early Embryonic Development of Animals. Spinger-Verlag, Berlin, Germany. This excellent resource contains a chapter on zebrafish. The chapter summarizes normal development and genetic manipulations. Herbst, C., 1892. Expt. Untersuchungen uber den Einfluss des veranderten chemischen Zusammensetzung des umgebenden Medium auf die Entwicklung der Tiere. Zeitschr. F. wiss. Zool. 55: 446–518. A classic paper by a classical embryologist from over a century ago, this discusses the effects of a number of chemicals, including lithium, on embryonic development. Hill, J., J. D. W. Clarke, N. Vargesson, T. Jowett, and N. Holder, 1995. Exogenous retinoic acid causes specific alterations in the development of the midbrain and hindbrain of the zebrafish embryo including positional respecification of the Mauthner neuron. Mech. Dev. 50: 3–16. In this study, zebrafish embryos in the early gastrula stages were treated with retinoic acid, and the effect of this treatment on brain development was then analyzed. Hisaola, K. K., and H. I. Battle. 1958. The normal developmental stages of the zebrafish, Brachydanio rerio (Hamilton-Buchanan). J. Morphol. 102: 311–328. This is the first complete published description of the normal stages of zebrafish development through hatching. It provides a simplified alternative to the more extensive Kimmel et al. staging series (1995).

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Holder, N. and A. McMahon. 1996. Genes from zebrafish screens. Nature 384: 515–516. This is a short review of the huge mutagenic screens that were done to isolate mutants in zebrafish that affect embryonic development. Holder, N., and J. Hill. 1991. Retinoic acid modifies development of the midbrain-hindbrain border and affects cranial ganglion formation in the zebrafish embryo. Development 113: 1159–1170. This is an important paper showing that even a short exposure to retinoic acid causes damage to the mid- and hindbrain as well as to cranial ganglia. The damage to ganglia suggests that retinoic acid affects cranial neural crest cells. Jesuthasan, S., and U. Strähle. 1997. Dynamic microtubules and specification of the zebrafish embryonic axis. Curr. Biol. 7: 31–42. This paper discusses the role of microtubules in the reorganization of components along the animal-vegetal axis during zebrafish development. Among other things, the authors discuss how treatment with cold temperatures (18ºC or lower) during early cleavage stages leads to widespread developmental defects due to the disassembly of microtubules. Karlstrom, R. O., and D. A. Kane, 1996. A flipbook of zebrafish embryogenesis. Development 123: 461. Each frame of a time-lapse movie of zebrafish development is illustrated on the pages of this volume of the journal. Flipping through the pages allows you to see the development of the embryo as you would see it in a movie. Kimmel, C. B., W. W. Ballard, S. R. Kimmel, B. Ullmann, and T. F. Schilling. 1995. Stages of embryonic development of the zebrafish. Dev. Dyn. 203: 253–310. This is the authoritative, very detailed account of the developmental stages of zebrafish. Anyone doing serious work with zebrafish needs a copy in order to learn how to accurately stage the embryos. It can also be found reprinted in The Zebrafish Book by Westerfield. Kimmel, C. B., and R. Warga. 1986. Tissue specific cell lineages originate in the gastrula of the zebrafish. Science 231: 365–368. This paper describes fate map studies in the zebrafish embryo using a fluorescent dye as a marker that is injected into single blastomeres. Klein, P. S., and D. A. Melton, 1996. A molecular mechanism for the effect of lithium on development. Proc. Natl. Acad. Sci., U.S.A. 93: 8455-8459. This paper offers the hypothesis that lithium’s teratogenic effect is through the inhibition of glycogen synthase kinase in the Wnt/β-catenin pathway, a conclusion they make from their studies on Xenopus embryos. Klymkowsky, M. W. and J. Hanken. 1991. Whole-mount staining of Xenopus and other vertebrates. In Methods in Cell Biology, 36. Xenopus laevis: Practical Uses in Cell and Molecular Biology. (B. K. Kay and H. B. Peng, (eds.). Academic Press, San Diego, CA. pp. 419–441. This chapter gives full instruction on how to stain bone and cartilage in whole mount preparations. Laale, H. 1971. Ethanol induced notochord and spinal cord duplication in the embryo of the zebrafish, Brachydanio rerio. J. Exp. Zool. 177: 51–64. This paper describes the effects of treating zebrafish embryos with 1–3% ethanol. Embryos in the high blastula stage were exposed to ethanol for 24 hours. Developmental abnormalities included fusion of the optic cups mid-ventrally and duplications of the notochord and spinal cord. Metcalfe, W. K. 1994. The zebrafish. In Embryos: Color Atlas of Development. J. Bard, (ed.). Wolfe Publishing, London, England. pp. 135–147. This chapter is from a beautifully illustrated book on embryonic development in various model organisms. The full-color pictures are exquisite. The chapter includes an excellent summary of development and the use of mutants for studying development. Mullins, M. C., et al. 1996. Genes establishing dorsoventral pattern formation in the zebrafish embryo: the ventral specifying genes. Development 123: 81–93. This paper discusses six genes involved in

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specifying ventral regions in the early zebrafish embryo. It compares mechanisms of establishing dorsal-ventral patterning in the zebrafish embryo with what is known about dorsal-ventral patterning in the amphibian embryo. Saunders, A. 1996. Exposure of embryonic cells to alcohol: contrasting effects during preimplantation and postimplantation development. Semin Perinatol. 20(2): 127–139. This paper covers in succinct manner the effects of alcohol on human development. Soprano, D. 1995. Retinoids as teratogens. Ann. Rev. Nutrition 15: 111–132. This is a useful review of the detrimental effects that retinoids can have on human development. Stachel, S. E., D. J. Grunwald, and P. Z. Myers. 1993. Lithium perturbation and goosecoid expression identify a dorsal specification pathway in the pregastrula zebrafish. Development 117: 1261–1274. In this paper, the authors demonstrate that lithium treatment disrupts dorsal-ventral patterning in the zebrafish embryo. They obtain similar results whether the embryos are treated with or without their chorions intact. Strähle, U., and S. Jesuthasan. 1993. Ultraviolet irradiation impairs epiboly in zebrafish embryos: evidence for a microtubule-dependent mechanism of epiboly. Development 119: 909–919. This paper discusses the damaging effects of ultraviolet radiation in the UV-C range on early development in zebrafish embryos. The results indicate that epiboly is disrupted through the disruption of microtubules within the egg. Weinberg, E. S. 1992. Analysis of early development in the zebrafish embryo. In Early Embryonic Development of Animals. W. Hennig (ed.). Spinger-Verlag, Berlin, Germany. pp. 91–150. This chapter gives an excellent detailed account, along with photographs, of embryonic development of the zebrafish. It also includes detailed descriptions of how to create mutants and explains what is known about a number of important developmental mutants. Westerfield, M. 1995. The Zebrafish Book, A Guide for the Laboratory Use of Zebrafish (Danio rerio). University of Oregon Press, Eugene, OR. This classic is now out of print, but is in the process of being reissued. It is the authoritative reference on all things zebrafish, and no lab studying zebrafish can afford to be without a copy of it. Zhang, Z. 1996. Specific teratogenic effects of different retinoic acid isomers and analogs in the developing anterior central nervous system of zebrafish. Dev. Dyn. 206: 73–86. In this paper, the author compares the teratogenic effects of all-trans retinoic acid with that of 9-cis retinoic acid. The results show that 9-cis retinoic acid is a more potent teratogen than all-trans retinoic acid.

Suppliers From most pet stores or a biological supply house such as: Carolina Biological Supply Co. 2700 York Road Burlington, N.C. 27215 Tel. 1-800-334-5551 www.carolina.com Zebrafish (Danio rerio) Fish food Fish tanks, tank lights, timers,

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Tank filters such as Tetra Whisper® Bio-Bags; change filter approx. every 1-2 months. Aquarium testing kits for pH, ammonia, and nitrite

Any good biological supply company or chemical supplier, such as: Fisher Scientific 585 Alpha Dr. Pittsburgh, PA 15238 Tel. 1-800-766-7000 www.fishersci.com

Sigma Chemical Company P.O. Box 14508 St. Louis, MO 63178 Tel. 1-800-325-3010 www.sigma-aldrich.com Alcian blue Alizarin red S Lithium chloride Pronase Retinoic acid Salts for Hank’s solution Standard histological supplies Tricaine

Glossary Alcian blue: A dye that, when in an acidic solution, stains glycosaminoglycans (e.g., mucus and chondroitin sulfate of cartilage). Alcohol dehydrogenase: An enzyme that breaks down alcohol. It is present in the liver and detoxifies alcohol. The liver of a human fetus lacks this enzyme, and alcohol ingested by the pregnant mother will enter the amniotic fluid and be present, bathing the developing fetus, for a far longer time than it is present in the mother’s system. Alizarin red: A dye that, when in a basic solution, stains the calcium of bone. Amniote egg: The type of egg typical of the amniotes, and represents an egg that can survive on land. It consists of a hard outer shell that protects the embryo from physical damage and dehydration, a large yolk and a watery albumen that provides the nutrients and water necessary till hatching, and the embryo develops four extraembryonic membranes, the chorion, amnion, yolk sac, and allantoic membranes. Animal pole: The point on the surface of the egg where polar bodies are given off. Eggs usually orient to gravity with their animal pole oriented uppermost. Anlage (anlagen, pl): An embryonic primordium determined to form a specific organ or part of organ, an organ rudiment.

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Axis formation: The determination of the axes of the embryo. In a bilateral organism, there are three axes: anterior-posterior, dorsal-ventral, and proximal-distal. Typically, these are determined early in development. Beta-carotene: A precursor to vitamin A, found in yellow, red, orange, and green leafy vegetables. Vitamin A is a necessary component to anyone’s diet, though in overdoses it is dangerous and acts as a teratogen. In its precursor form, as beta-carotene, however, it is safe even in large quantities. Blastoderm: The layer of cells formed in the embryo during cleavage. In Drosophila this layer is formed in the periphery of the yolky egg. In the chick and fish, it is formed as a disc-shaped area at the animal pole of the yolky egg. Blastodisc: In a chick or fish egg, the puddle of non-yolky cytoplasm that sits atop of the vast quantity of yolk. The nucleus of the egg sits here, and it is the blastodisc where development begins. Blastomeres: The cells that are created through cleavage during early embryonic development. Blastula stage: The stage in embryonic development following cleavage and preceding gastrulation. The embryo may form a hollow blastula (coeloblastula) or a solid blastula (stereoblastula). The cavity formed in a coeloblastula is called the blastocoel. Branchial arches: A synonym for pharyngeal arches. Also called visceral arches. Brine shrimp: Artemia salina. A small, salt-water branchiopod crustacean, often used in the laboratory as a food source for other organisms; it can survive in salinities 0.1 to 10 times the salt concentration of normal seawater. The fertilized eggs of these are commonly sold in pet stores as “sea monkeys.” Carotenoid pigments: The yellow-orange pigments found in leafy green and yellow-to-red vegetables, from which the body makes vitamin A. The yellow color of yolk is from carotenoid pigments, derived from the bird’s food. Cerebrum: In a vertebrate, the anterior-most region of the brain, also called the telencephalon. It is associated with olfactory organs and the sense of smell, as well as being the center of intelligence. Chaoborus: A dipteran insect called a phantom midge. Its aquatic larval stage lives in freshwater and preys upon mosquito larvae, serving as a natural mosquito control. The larvae can be found in pet stores as frozen “white worms” or “glass worms,” sold as fish food. Chordata: The phylum that includes the vertebrates and two invertebrate groups, the urochordata and the cephalochordata. Diagnostic features of the phylum are the presence of a notochord (hence its name), a dorsal hollow nerve cord, and the presence of pharyngeal pouches during development. Chorion: A tough, protective eggshell. In Drosophila and other terrestrial insects, it is specially designed for maximal gas exchange with minimal water loss. In fish, it is a clear, tough enveloping layer. Clearing: The process in a histological preparation in which tissues become clear and translucent. Common clearing agents include xylene and toluene, which are immiscible with water, and require that the tissues be dehydrated prior to clearing. Glycerin is a clearing agent that is miscible with water, and can be used without dehydrating the specimen. Cleavage: The stage in embryonic development, normally following fertilization, in which the egg, or portions of the egg, divide by mitosis into a number of cells. Cleavage results in the formation of the blastula. Cortical granule reaction: The reaction in the cytoplasm of an egg upon fertilization in which the cortical granules of the egg fuse with the egg cell membrane, releasing their contents to the exterior.

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The extracellular matrix around the egg is modified by these contents such that further sperm are unable to get to the egg cell surface. Cortical granules: Vesicles found in the cortical cytoplasm of an unfertilized egg. These contain components that are released to the exterior upon fertilization or activation of the egg. These components modify the extracellular substances surrounding the egg such that sperm are no longer able to approach the egg cell surface. Craniofacial defects: Defects of the head and face. Cyclopia: A developmental defect in which only one eye develops. Danio rerio: The species name for the zebrafish, previously known as Brachydanio rerio. Daphnia: A freshwater branchiopod crustacean, commonly known as a water flea. These organisms can be easily raised in the laboratory and are often used as a food source for other organisms such as zebrafish. DDT: Dichloro-diphenyl-trichloroethane, a chlorinated organic pesticide. DDT initially seemed to be an inexpensive, effective and relatively non-toxic compound, but it has been shown instead to be highly toxic to many organisms. It accumulates in fatty tissue. In humans, it can cause liver damage, liver cancer, damage to the reproductive system and reduced reproductive success, among other things. Its use was banned in the United States in 1972, but it is still used in other parts of the world. Dechorionation: Removal of the chorion. The chorion is a tough extracellular coat that surrounds the eggs of many species (e.g., Drosophila, zebrafish). A chorion can usually be removed either physically with fine forceps or chemically with an enzyme capable of dissolving the chorion (e.g., Clorox®, in the case of the Drosophila chorion, and pronase in the case of the zebrafish chorion). Deep cells: In the fish blastula and gastrula, the middle layer of cells of the blastoderm. These cells form the body of the embryo. Dehydration: The removal of water. In histological procedures, a specimen is usually dehydrated through a graded series of alcohol prior to embedding or mounting. Demersal: Living or found at the bottom of a body of water. Diencephalon: The region of the forebrain posterior to the telencephalon. Its lateral evaginations form the optic vesicles, a ventral evagination forms the infundibulum, and a dorsal evagination forms the epiphysis. Dioxin: One of the most toxic group of chemicals known, dioxin is formed as a by-product of many industrial processes involving the burning of chlorine-based chemical compounds with hydrocarbons. It is produced by waste incineration, chemical and pesticide manufacturing, polyvinyl chloride plastics production, and pulp and paper bleaching. It was the primary toxic agent of Agent Orange. It is a carcinogen, and a teratogen, it disrupts the immune system, and acts as an endocrine disruptor. There is no “safe” level of exposure to dioxin. Dominance hierarchies: A hierarchy established within a social group of animals, usually through aggressive behavior. Individuals establish dominance over others, and thereby gain preferred access to food and other resources. Dorsalized: A developmental abnormality in which no ventral structures develop in the embryo. Drosophila: A fruit fly, a genus of dipteran insects. The common fruit fly, Drosophila melanogaster, is a model organism in developmental biology. Ebstein anomaly: A congenital heart defect in which the tricuspid valve is displaced downward in the right ventricle. The ventricle is less muscular and the tricuspid valve is leaky. Babies with this condition are usually blue at birth and may require immediate surgery.

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Ectoderm: The germ layer in an embryo that gives rise to the epidermis and nervous system. In many cases, as in vertebrates, an anterior and posterior invagination of ectoderm gives rise to the stomodeum and proctodeum, respectively. Egg water: A very dilute salt solution that is suitable for maintaining fish embryos. It can be made from instant ocean (60 mg/liter distilled water) or as a 10% solution of Hank’s solution. Embryonic shield: In fish development, the region of the germ ring where cells are converging. It is analogous to the dorsal lip of the blastopore in the amphibian embryo, and it is here that the body of the embryo will be organized. Endocrine disruptors: Exogenous substances that interfere with the normal function of hormones. Environmental endocrine disruptors (such as DDT, dioxin, phthalates, PCBs) are considered to be a general threat to species and have been the focus of a great deal of recent research. Endoderm: The germ layer in an embryo that gives rise to the epithelium (lining) of the gut and gut derivatives. Enveloping layer (EVL): In the fish blastula and gastrula, the outer-most layer of cells of the blastoderm. These cells protect the developing embryo by becoming the periderm, an outer layer of cells that is eventually sloughed. Epiblast: The uppermost layer of the blastoderm formed during gastrulation. In the fish embryo, involution within the deep cells separates the hypoblast from the upper epiblast; the epiblast will form the ectoderm. Epiboly: A type of gastrulation movement in which a sheet of cells spreads to cover a greater surface area. Epidermal ectoderm: The region of ectoderm that forms the epidermis (the epithelial layer of the skin). Epidermis: The epithelium of the skin. It is typically a stratified squamous epithelium, with outer keratinized layers of cells that protect against dehydration and physical abrasion. Epithelium: A major type of tissue in an organism, found lining body cavities and covering the outside of the body. The epidermis is an example of a specific type of epithelium. Estrogenic: Having an effect similar to that of estrogen, either by mimicking estrogen or inhibiting the effectiveness of androgen. EVL: The enveloping layer of the fish blastula and gastrula. Exogastrula: A deformity in developing embryos in which the archenteron, instead of invaginating, evaginates, forming an external archenteron. Fertilization: Fusion of egg and sperm. Fertilization cone: In certain eggs such as the sea urchin and zebrafish, a structure that forms during fertilization at the point of sperm entry. Microvilli elongate and wrap themselves around the entering sperm. These appear as a small cone-shaped structure on the surface of the egg. Fetal alcohol effect: Developmental defects caused by moderate exposure to alcohol. A milder condition than fetal alcohol syndrome, it consists primarily in slowness in learning. It can be caused by a pregnant mother consuming only small amounts of alcohol. It is now considered that there are no safe levels of alcohol that can be consumed during pregnancy. Fetal alcohol syndrome: A well-recognized collection of developmental defects caused by exposure to alcohol; these include facial abnormalities, heart defects, and mental retardation and can be caused by 1–2 alcoholic drinks/day consumed by the mother during pregnancy. Fin bud: The precursor to the pectoral and pelvic fins, it consists of an outgrowth of mesenchyme covered by epidermis, much like the limb bud of a tetrapod vertebrate.

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Fin fold: In a fish embryo, a median fold of the skin that gives rise to the dorsal, caudal, and anal fins. Fixation: The process of fixing a tissue. Fixation kills the tissue, as well as any bacteria present, and coagulates (cross-links) proteins. Fixative: Anything that fixes a living organism so that it can’t be degraded by bacteria or other microorganisms. “Floppy infant” syndrome: A birth defect that can be caused by toxic levels of lithium and is characterized by weakness, lethargy, a weak cry, low body temperature, and a poor appetite. Footed coverslip: A coverslip that has a small spacer on each corner. It can be made by nicking the corners of the coverslip against some hard paraffin, so that crumbs of paraffin remain attached at each corner. Alternatively, vacuum silicone grease can be used. This can give just enough spacer between the slide and the coverslip to avoid crushing the specimen on the slide. Forebrain: The anterior-most region of the brain. In the embryonic vertebrate, this is the prosencephalon, which gives rise to the telencephalon and diencephalon. Formaldehyde: A pungent gas, HCHO, used in solution as a fixative. It is sold as a solution (about 40% formaldehyde in water) called formalin. OSHA has determined that formaldehyde is carcinogenic, so care should be taken to avoid fumes and gloves should be worn to avoid contact with the skin. Galen: A Greek physician from the first century A.D. Gamete: An egg or sperm. Gastrulation: The stage in embryonic development that follows cleavage. During gastrulation, cells rearrange themselves, with the endoderm and mesoderm cells moving inward and the ectoderm cells spreading around the outside. Some ectoderm invaginates to form such structures as the stomodeum and proctodeum. Germ layers: The ectoderm, mesoderm, and endoderm of a developing embryo. Germ ring: In a fish embryo, the thickened ring of deep cells created by the involution of deep cells during early gastrulation. Glycerin jelly: A mounting medium used in making whole mounts of specimens that have not been dehydrated. It contains glycerin, gelatin, and water. It is heated before use to make it liquid; when it cools, it becomes a firm gel. Glycogen synthase kinase: An enzyme that phosphorylates glycogen synthase and thereby helps to regulate its activity. Glycogen synthase is an enzyme that causes the synthesis of glycogen from glucose. Glycogen is the principle storage form of glucose, and is found mainly in the liver and in muscle. Glycosaminoglycan (GAG): A long, unbranched polysaccharide (formerly called mucopolysaccharide) made up of repeating units of a disaccharide in which one of the sugars is an amino sugar. Complexes with protein to make proteoglycans. Out in the extracellular matrix, they cause extensive swelling of spaces by absorbing water. Examples of GAGs are hyaluronic acid (hyaluronate), chondroitin sulfate, heparan sulfate. Hank’s salt solution: A physiological, balanced salt solution. Can be easily made in the laboratory. Also available premade through biological supply houses. Hindbrain: The posterior-most region of the brain. In the embryonic vertebrate, this is the rhombencephalon, which gives rise to the metencephalon and myelencephalon. Hox genes: Developmentally important genes, containing a homeobox region, that are important for patterning the anterior-posterior axis in bilaterian organisms. Hyoid arch: The second pharyngeal arch. The hyoid arch forms supports for the jaws and tongue.

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Hypoblast: The lowermost layer of the blastoderm formed during gastrulation. In the fish embryo, involution within the deep cells separates the hypoblast from the upper epiblast; the hypoblast will form the endoderm and mesoderm. IACUC: Institutional Animal Care and Use Committee. U.S. Federal law requires that all institutions that use laboratory animals for research must establish an Institutional Animal Care and Use Committee (IACUC) to oversee and evaluate all aspects of the institution's animal care and use program. Inositol triphosphate: IP3. This is created when the signaling molecule PIP2, found in the cell membrane, is split into DAG and IP3. IP3 causes the release of calcium ions from intracellular stores, raising cytoplasmic levels of calcium. Involution: A type of gastrulation movement in which a sheet of cells folds inward and spreads over an inner surface. Iridophores: A type of chromatophore, a pigment containing cell. Iridophores contain light-reflecting crystals, usually of guanine. Isotretinoin: 13-cis-retinoic acid. A synthetic form of retinoic acid used orally for the treatment of severe acne. It is highly teratogenic, and can have numerous side effects in adults as well. Kaiser’s glycerin jelly: A specific recipe for glycerin jelly, containing gelatin, glycerin, water, and a mold retardant such as thymol. It is used as a mounting medium for whole mounts. Lateral line system: A system of canals or grooves in the skin containing sensory structures called neuromasts that are designed to detect low-frequency motions within the water. Lateral line systems are found in fish and aquatic and larval amphibians. Lens placode: In the vertebrate embryo, a thickened disc of epidermal ectoderm cells that form next to the optic vesicle. The lens placode later invaginates to form the lens vesicle, and ultimately forms the lens of the eye. Lithium: A silver-white metal, the lightest of the metal elements. It is toxic to a number of organ systems. Embryos exposed to lithium show severe defects. It is prescribed as lithium carbonate to patients with manic depressive (bipolar) disorder. Macrolecithal egg: An egg with a very large yolk, as in the eggs of birds, fish, and insects. Mandibular arch: The first pharyngeal arch. The mandibular arch forms the jaws, the upper portion forming the maxilla and the lower portion forming the mandible. Manic depressive disorder: Also called bipolar disorder, this is a disease in which the patient suffers from severe mood swings consisting of both manic phases, in which the patient has an excessive amount of energy, and phases of depression. Lithium is commonly used for treating this disorder since it helps to stabilize the mood swings. Maternal mRNA: Maternal messenger RNA. This is the messenger RNA that is stored in the egg during the primary oocyte stage while the egg is still developing within the ovary. This stored mRNA will direct development until the genes of the embryo are turned on. Medulla oblongata: In a vertebrate, the posterior-most region of the brain, also called the myelencephalon. In embryonic development, it is characterized by a series of neuromeres, each of which will become associated with a specific set of motor and sensory nerves. Meroblastic cleavage: Incomplete cleavage. Meroblastic cleavage occurs in macrolecithal eggs, such as that of the fish or bird, during early cleavage. The cleavage planes cutting down through the blastodisc toward the dense yolk are stopped when they reach the yolk. The egg, therefore, does not divide completely. Mesencephalon: The midbrain, the middle region of the brain that will serve primarily for processing data from the eyes and ears.

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Mesoderm: The germ layer in an embryo that gives rise to the muscular and circulatory systems and most of the skeletal and urogenital systems. Metasynchronous: Not precisely synchronous. Metencephalon: Forms the cerebellum dorsally and the pons ventrally. The cerebellum primarily coordinates stimuli concerning body position and movement; the pons is a bundle of connective pathways shunting information between the cerebrum and cerebellum. Methoprene: An insecticide that inhibits maturation of insects, known as an insect growth regulator. Though initially considered safe, it has been found to break down into harmful retinoids when exposed to UV-radiation. It has been linked to deformities in frogs, including missing limbs, extra limbs, and extra toes. Microcephaly: A birth defect in which the head fails to develop to the normal size due to microencephaly, a small brain. It is often accompanied by mental retardation. Microphthalmia: A birth defect in which one or both eyes are smaller than normal. This usually results in a restricted field of vision, which may be accompanied by other impairments to vision. Micropyle: Found in the eggs of certain organisms, such as the fruit fly and zebrafish, a small channel through the chorion of the egg that provides an entryway for sperm at fertilization. Microtubules: Thin tubules found in the cytoplasm of a cell. Microtubules are made up of tubulin subunits. Microtubules function both as a cytoskeletal support and for transport within the cell. Motor-molecules, associated with the microtubules, are able to ferry such things as organelles along the microtubules. Mid-blastula transition: The time during the blastula stage at which the genes of the embryo begin to be transcribed. Up until this point, development of the embryo is orchestrated by maternal messenger RNA stored within the cytoplasm of the egg. In the zebrafish embryo, the mid-blastula transition begins at the 512-cell stage (ninth cleavage) after 2 3 / 4 hours of development (at 28ºC). Midbrain: The middle region of the brain. In the vertebrate embryo, this is the mesencephalon, which serves primarily for processing data from the eyes and ears. Model organism: An organism that has been used extensively for scientific inquiry such that a large body of information is accumulated on the organism. Because further work can build on this backlog of information, the organism becomes a preferred species for laboratory investigations, with the assumption that it will serve as a model for other organisms. Mounting medium: The medium in which a specimen is mounted for observation under the microscope. A mounting medium can be water soluble, such as glycerin jelly, or immiscible with water, such as a plastic polymer dissolved in a solvent. When a mounting medium dries, it is firm and transparent, and typically has the same refractive index as glass. Mutagenic screen: A procedure for isolating new and novel mutants that are of specific interest. A major mutagenic screen, done on zebrafish, was successful in isolating a number of mutants affecting embryonic development. In this screen, male zebrafish were mutagenized with a potent mutagen (ethyl nitrosurea) and then outcrossed. Subsequent incrossings among individuals of F1 and F2 generations were then conducted. Pairs from these crosses that produced mutant embryos were then outcrossed to carry the mutations to subsequent generations. Myelencephalon: Also called the medulla oblongata, it is associated with a series of enlargements called neuromeres, each of which becomes associated with a specific set of motor or sensory nerves. Negative phototaxis: Directed movement away from light. Neural ectoderm: The region of ectoderm that forms the neural tube. Neural fold: The neural ectoderm that is folding upward to form a neural tube.

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Neural plate: The neural ectoderm prior to its invagination to form the neural tube. Neural tube: The tube formed by the neural ectoderm. The precursor to the brain and spinal cord. Neuromasts: The sensory structures found in the lateral line system of fish and aquatic and larval amphibians. They consist of a gelatinous cupula that sits on sensory cells. Deflection of the cupula by water currents in turn moves the ends of the sensory cells, causing the transmission of a neural signal. Neuromeres: In a vertebrate embryo, the series of enlargements or the developing neural tube, marked off by constrictions, that correspond to different regions of the brain. Neurula: The stage in embryonic development when neurulation is occurring. Neurulation: The morphogenetic movements of the neural ectoderm that form a neural tube out of the neural plate. Neutral buffered formalin: A commonly used fixative containing 10% formalin and buffered with sodium phosphate. Olfactory placodes: Also called nasal placodes. A pair of epidermal thickenings at the anterior region of the head that will invaginate to form the lining of the nasal cavities and the olfactory nerve. Ooplasmic segregation: The segregation of specific informational molecules within the egg cytoplasm to different regions of the egg, so that as the egg cleaves, different cells receive different types of information. Optic cup: In a vertebrate embryo, the precursor to the neural and pigmented retinas of the eye. During eye development, optic vesicles that form as outpocketings of the diencephalon invaginate to form the optic cups. Optic vesicles: Lateral evaginations of the diencephalon. They later form the optic cups that will become the neural and pigmented retinas of the eyes. Organogenesis: The stage during embryonic development following gastrulation when organs are forming. Otic placode: Epidermal thickening that will invaginate to form the inner ear. Paramecia: A ciliated freshwater protozoan. Periderm: In a developing embryo, an outer layer of cells that is protective. It does not participate in forming any structures for the embryo and is eventually sloughed. Pharyngeal arches: Also called branchial arches and visceral arches. These are the bars of tissue in the chordate embryo that separate the pharyngeal pouches. A pharyngeal arch contains a visceral cartilage, muscle associated with this cartilage, a blood vessel known as an aortic arch, and a cranial nerve. Pharyngeal pouches: Also called visceral pouches. Typical of chordates. These form embryonically. In organisms with gills, they will form the gill slits. In higher vertebrates, they are transformed into structures such as the eustachian tube and cavity of the middle ear, the epithelium of the tonsils, thymus, parathyroid, and ultimobranchial bodies. Pharyngeal region: The region of the pharynx. Phosphatidylinositol bisphosphate (PIP) cycle: A signal transduction pathway that is initiated by the splitting of phosphatidylinositol bisphosphate (PIP2), a phospholipid within the cell membrane. PIP2 splits into inositol triphosphate (IP3) and diacylglycerol (DAG). IP3 causes the release of calcium ions from intracellular stores, raising cytoplasmic levels of calcium. DAG is involved with activating an ion pump that moves hydrogen ions out of the cell and sodium ions into the cell, raising the intracellular pH. Phototaxis: Directed movement towards light.

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Phthalates: A class of widely used industrial compounds known as dialkyl or alkyl aryl esters of 1,2-benzenedicarboxylic acid. Used widely as softeners of plastics, oily substances in perfumes, additives to hairsprays, lubricants, and wood finishers, and causing the “new car small” when it volatilizes from a hot plastic dashboard, they are thought to cause reproductive damage in both males and females. PIP cycle: Phosphatidylinositol bisphosphate cycle. A signal transduction pathway that is initiated by the splitting of phosphatidylinositol bisphosphate (PIP2), a phospholipid within the cell membrane. PIP2 splits into inositol triphosphate (IP3) and diacylglycerol (DAG). IP3 causes the release of calcium ions from intracellular stores, raising cytoplasmic levels of calcium. DAG is involved with activating an ion pump that moves hydrogen ions out of the cell and sodium ions into the cell, raising the intracellular pH. Placode: A flat, round plate of cells. P-nonylphenol: A product contained in industrial and household detergents. It is an environmental pollutant that has estrogenic effects. Polyspermy: Fertilization of an egg with more than one sperm. Posteriorizing signal: A developmental signal that specifies posterior structures along a posterioranterior axis. Pronase: A mixture of endo- and exo-proteinases that is prepared from bacteria and cleaves almost any peptide bond. It is available from biological and chemical supply houses. Prosencephalon: The forebrain. In the vertebrates, this consists of an anterior telencephalon and a posterior diencephalon. RA: An abbreviation for retinoic acid. Retinal: A form of vitamin A from which the pigment rhodopsin is made. Retinoic acid: A derivative of vitamin A, it is a posteriorizing signal in many developing systems and is known to influence Hox gene expression. It is highly teratogenic. Retinol: An alternate name for vitamin A. Rhombencephalon: The hindbrain. In the vertebrates, this consists of a metencephalon anteriorly and a myelencephalon posteriorly. Rhombomeres: The neuromeres of the hindbrain (rhombencephalon). Somites: Blocks of mesoderm that become subdivided into a sclerotome, myotome, and dermatome. Sclerotomes form vertebrae and ribs, myotomes form skeletal muscles for the back and appendages, and dermatomes give rise to dermis. Somitic: Of the somite. Spawning: The release of gametes (eggs or sperm). Sperm: The male gamete. Spina bifida: A congenital defect in which the neural plate doesn’t fully close when making the neural tube, and parts of the neural tube remain open. Spinal cord: The posterior continuation of the neural tube. Staging series: An embryonic timing series that identifies stages of development. Complete staging series are valuable, and once a good one for an organism has been published, this is usually used for many years. Stained: In histological procedures, this means dyed with histological dyes to contrast various components of cells and tissues. Syncytium: A multinucleate tissue in which cell membranes don't completely separate the nuclei. Tail bud: In an embryo, the posterior bud that will form the tail.

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Telencephalon: The anterior region of the forebrain. It is associated with olfaction and is the center of intelligence. More commonly called the cerebrum. Teleost: The higher bony fish, in the subclass Actinopterygii (the ray-finned fish) and the class Osteichthyes (bony fish). Telolecithal: Having yolk concentrated toward the vegetal pole of an egg. Teratogen: Any substance that can cause birth defects. Tetralogy of Fallot: A developmental defect in which there are four problems in the development of the heart: a ventricular septal defect in which there is a hole in the septum between the two ventricles, a pulmonary stenosis in which there is a narrowing of the right ventricular outflow, a defect in which the aorta is misaligned so that it sits over the ventricular septal defect, and a defect in which the right ventricle is more muscular than normal. Tetraploid: Having four sets of chromosomes. Most organisms are diploid, having two sets of chromosomes. Tretinoin: All-trans retinoic acid, a synthetic form of retinoic acid that is used topically for superficial pustular acne. When exposing embryos directly to the substance, it is highly teratogenic; when applied topically on the skin of a pregnant mother, it is not considered a threat to the fetus. Vertebrata: The class of chordates that includes the fish, amphibians, reptiles, birds and mammals. Diagnostic features of the vertebrates are a vertebral column (hence its name), neural crest cells during development, and the presence of bone. Visceral arches: Synonym for pharyngeal arches. Vitamin A: A fat-soluble vitamin; retinol. It is used by the body to make visual pigments, and is important for the health of the skin, eyes, and epithelia in general. It is needed for lysosomal stability and glycoprotein synthesis. It is the precursor to two active forms of the vitamin, retinal and retinoic acid. Vitelline envelope: An extracellular coat surrounding an ovum. Washing: In histological procedures, this is the step following fixation in which the fixative is washed out of the tissue. Whole mounts: A histological preparation of a specimen in which the whole specimen is mounted. The specimen is usually mounted on a microscope slide, either in a water-mountant such as glycerin jelly, or a non-water mountant such as the plastic mounting media used routinely for stained paraffin sections. β -catenin pathway: A signal transduction pathway that involves the binding of the Wnt protein Wnt/β at the cell surface, causing the release of β-catenin inside the cell so that β-catenin can then enter the nucleus as a transcription factor, bind with other transcription factors, and activate various genes. Xenopus laevis: The South African clawed frog or African clawed frog (previously commonly called the South African or African clawed toad). It is a species of frog that has become a model organism for amphibian development because of the ease with which it can be raised and maintained in the laboratory. Yolk syncytial layer (YSL): In the fish blastula and gastrula, the syncytial layer of nuclei within the yolky cytoplasm that forms the lower-most layer of the blastoderm. It is formed by the fusion of the lower-most blastoderm cells with the yolk below. The yolk syncytial layer leads the rest of the blastoderm down around the yolk sphere during gastrulation. YSL: The yolk syncytial layer of the fish blastula and gastrula.

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Zebrafish: Danio rerio, previously known as Brachydanio rerio. A small, freshwater teleost fish originally identified in the Ganges River in northeastern India. It is now commonly used as a model organism in studies of embryonic development.

Zebrafish Development Questions Name: ______________________

Date: ___________________

Please print these sheets and then answer all questions prior to the laboratory period. This will help prepare you for the laboratory. Information necessary for answering the questions can be found on Vade Mecum2 and in the Laboratory Manual that is included on the CD-ROM. 1. How can you distinguish an adult male from an adult female zebrafish?

2. At what stage is the zebrafish embryo after 24 hours of development at 24ºC. List the structures that you can see in the embryo at this point.

3. When does the larva hatch from its chorion, and at what stage must you start feeding the larva?

4. How do you remove the chorion from an embryo?

Name: _______________________

Zebrafish - 2

5. What effects do low temperatures have on the development of the embryo? What effect does low temperatures have on microtubules?

6. What forms of vitamin A are known to have teratogenic effects? What forms are considered safe for developing embryos?

8. What developmental abnormalities can be caused by alcohol? Is there any recognized safe limit of alcohol that a pregnant woman can imbibe?