Patterns and Experiments In Developmental Biology

ISBN: 0-07-237965-0 Description: ©2001 / Spiral Bound/Comb / 256 pages Publication Date: January 2001 Overview A labora...

Author: Leland G. Johnson | E. Peter Volpe

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ISBN: 0-07-237965-0 Description: ©2001 / Spiral Bound/Comb / 256 pages Publication Date: January 2001

Overview A laboratory manual for developmental biology offering basic, easy to use, laboratory investigations (18 experiments) spanning various models including echinoderm (Sea Urchin), amphibian (Frog), chick embryo, and fern gametophyte.

Johnson:Johnson & Volpe’s Patterns & Experiments in Developmental Biology, 3/e

Front Matter

Preface

© The McGraw−Hill Companies, 2004

Preface

As with the earlier editions, the goal of this edition of Patterns and Experiments is to facilitate and encourage developmental biology and embryology laboratory experiences that bring students together with fascinating and dynamic developing systems. Professional biologists and nonbiologists both often relate that the study of some aspect of development of a living organism has been a memorable highlight in their educational experience. How fascinating it is to watch those tiny clusters of cells as one makes that first marathon set of observations of a batch of developing sea urchin embryos. How exciting it is to return to the lab to find a vigorously beating heart in an in vitro cultured chick embryo where there had been no visible heart and a much simpler form only twenty-four hours earlier. My own view of biology and my career plans changed when I had that experience. I want to say to students who will use this manual that I envy you the excitement that comes with those first opportunities to experiment with living, developing organisms. I hope that a few of you might be inspired to go on to careers researching developmental processes and sharing the fascination of development with your own students. This is a truly exciting time in developmental biology because we are now able to investigate directly many of the genetic mechanisms underlying various developmental processes. However, as you begin your study of developmental biology, whether you pursue that study only in this course or study development for many years to come, I would like to offer one bit of advice from the perspective of many years in developmental biology.As intently as you may study certain individual developmental processes,please try not to lose sight of the whole developing organism and the still broader picture of the role of development in the perpetuation of species. Much of the fascination and beauty of development is to be found at those levels. This third edition of Patterns and Experiments includes a number of additions and new features. Several of the additions are to the considerably expanded section on echinoderm development. There are much more detailed directions for caring for sea urchin and sand dollar embryos and larvae ( Laboratory 1 and Appendix A). Several colleagues have reported that their students have been frustrated with their inability to observe development beyond the earliest stages, and I think that these directions will make it much easier for students to observe additional parts of development.The simpler and more effective procedure for blastomere separation that has been incorporated into Laboratory 2 should make it easier for students to conduct “twinning” experiments like those that have such a rich history in developmental biology’s past. Laboratory 2 also includes a fascinating new experiment on the somewhat surprising, but very adaptive, capacity of echinoderm embryos and larvae to regenerate lost cilia. Also, reorganization of the echinoderm portion of the manual led to creation of a new part ( Laboratory 3) that includes investigation of differentiation of an enzyme system. This investigation provides students a chance to study specific localized genetic activation in differentiation.Also,“Suggestions for Further Investigation of Echinoderm Development” was reorganized and substantially rewritten. vii


Front Matter

Preface


There is an important addition to the chick embryo section as well. In Laboratory 11, an earlier brief suggestion about investigating heart duplication has been expanded to a full experiment on heart rudiment separation and heart tube duplication that includes informative new illustrations. Numerous other updates and additions, including several added illustrations, have been made throughout the manual. Well over one hundred new references have been incorporated into the “Suggestions for Further Investigation” that appear at the ends of the portions of the manual. Each set of references has been updated, and the majority of the new references are to works that have been added to the very dynamic literature of developmental biology since publication of the second edition of Patterns and Experiments in 1995. I’ve also added citations to a number of the useful websites, many of which have come into being since 1995 as well. I’ve tried for a modest mix of specialized websites as well as general ones that provide links to many more of the valuable resources now available on the World Wide Web and which are likely to incorporate additional links to important sites that surely will be developed in the coming years. Developmental biology is not a discipline isolated from other aspects of biology. This is particularly evident, for example, in regard to the worldwide ecological problem of declining populations of numerous amphibian species recognized during the 1980s and 1990s. Appendix G contains some suggestions concerning responsible use of amphibians in teaching that are relevant to this problem. That appendix also contains suggestions of strategies for teaching developmental biology without sacrificing adult vertebrate animals, which is an option that a number of biologists, including me, prefer to choose. I thank the colleagues and students who have used the earlier editions of this manual and have taken the time to share some of their experiences in developmental biology. They have made insightful comments about the manual and have offered helpful suggestions and criticisms.A number of those suggestions led to additions to the second edition, and others have influenced the development of this third edition. I also warmly thank the many colleagues from colleges and universities across the United States and Canada who have participated over the years in my summer workshops on the Developmental Biology Teaching Laboratory at the University of Maine’s Darling Marine Center. Those developmental biologists have brought their own individual perspectives and expertise to the workshop sessions, and we’ve shared some remarkable learning experiences in that beautiful setting. I owe them and my Darling Center colleagues a great deal. Finally, I wish once again to offer my thanks to Peter Volpe who was my colleague and mentor in preparation of the first edition of this manual. Several of the amphibian development labs, especially Laboratories 4, 5, 6, and parts of Laboratory 8 have been only slightly updated and have remained largely as Peter conceived them, as has Appendix B. Some years ago, Peter’s main interests moved into the areas of human development, medical genetics, and biomedical ethics, and he turned his full energy and attention to those pursuits. Thus, his direct involvement with this manual ended with the first edition, but his influence remains evident in a number of places. The manual’s current title stands as a recognition of his original contributions. Leland G. Johnson

viii

Preface


I. Echinoderm Development

1. Fertilization and Early Development of Sea Urchins and Sand Dollars


L A B O R A T O R Y

1 Fertilization and Early Development of Sea Urchins and Sand Dollars

Echinoid echinoderms (sea urchins and sand dollars, which are also known as irregular urchins) have been the subjects of many investigations of fertilization and early development, and much of our understanding of developmental processes in animals has come from this research. Sea urchin and sand dollar gametes are readily obtained just before, and during, the breeding season and their developing embryos can be cultured in seawater or salt solutions that approximate the osmotic and ionic properties of seawater. Eggs and embryos of many species are quite translucent, so it is possible to observe a number of cell activities during early development, using a light microscope. In this laboratory, you will have opportunity to observe development from fertilization through assembly of the pluteus larva, which is the swimming, feeding larval form that is characteristic of many of the echinoid echinoderms. Techniques Please read and understand the techniques for obtaining gametes for fertilization and for the observation of embryos before you begin this laboratory. Obtaining Gametes As in other echinoderms, the sexes are separate in sea urchins and sand dollars. In nature, gametes are discharged into the water, and the sperm swim freely until they reach an egg. Since the sexes are difficult or impossible to distinguish by external features, sex of an individual animal must be determined by observing the gametes that it sheds. Injection of a small amount of potassium chloride into the coelom will induce an urchin to shed its gametes.The sex of the animal can then be determined by observing the color of gametes that are extruded from gonopores of the aboral (dorsal) surface of the animal within a few minutes after injection. The eggs of most sea urchins and sand dollars range in color from translucent yellow to pale orange, but eggs of some species are darker and may have a reddish cast. Sperm, when shed in mass, appear white or very light gray.

1





You should be very careful about conditions under which gametes and embryos are maintained. Temperature control is especially important, and your instructor will provide information concerning temperatures that are appropriate for the species you are studying. 1. Gently blot excess water off an adult urchin and place it on a clean surface with its aboral (opposite to, or away from, the mouth) surface down. Induce shedding of gametes by injecting 1 or 2 ml of 0.5 M KCl through the membrane surrounding the mouth opening (perioral membrane). Sand dollars should be injected with a finegauge needle inserted at a very shallow angle.To enhance effectiveness of the KCl injection, it is advisable to divide the injected dose of KCl among two or three sites in the perioral membrane. Several websites demonstrate these techniques (see Materials, p. 8). It is very important to avoid possible contamination of eggs with sperm.This can be accomplished by using a separate syringe and needle for each animal, but that usually isn’t practical.An alternative technique is to retain enough KCl solution in the syringe so that some can be expelled after each injection to flush the needle. Then rinse the needle surface with distilled water and dry it with a clean Kimwipe before refilling the syringe and injecting the next animal. 2. Collect eggs by inverting a female over a beaker or a finger bowl containing seawater.The water level in the beaker should be such that the female’s gonopores are in the seawater. The eggs will flow out of the gonopores and settle to the bottom of the beaker.After the eggs have been shed, they should be washed by decanting the supernatant water and replacing it with clean seawater.This washing removes coelomic fluid, broken spines, and body surface debris from the water. Eggs should be washed twice if time permits. Alternatively, it is possible to collect shed gametes directly from the body surface with a pipette, which helps avoid contamination by debris and extraneous fluids. Direct collection by pipette often is the best technique to use when only a few gametes are shed, as is sometimes the case with sand dollars. It is best to proceed with fertilization immediately, but if necessary, the eggs of some species can be refrigerated at 5° C for several hours and still respond fairly well in fertilization. 3. Active sperm, unlike eggs, are viable for only a few minutes in seawater.Thus, it is necessary to keep the sperm quiescent by collecting them under “dry” conditions (that is, in an undiluted suspension). A small portion of the “dry” suspension can be diluted in seawater each time active sperm are needed. When an animal has been identified as a male, wipe away excess moisture from among the spines on the aboral surface. Invert the male over a clean, dry petri dish or Syracuse dish.After several large drops of the white sperm suspension are in the dish, remove the animal and snugly cover the dish with parafilm or aluminum foil. The sperm should be kept concentrated until just prior to use, when they are activated by dilution in seawater. Collected sperm may be stored “dry” at a cool room temperature for an hour or so, but they should be stored in a 5° C refrigerator if longer storage is required. Sperm of some species can be stored in a refrigerator for up to a day. 4. Observe suspensions of eggs and sperm microscopically and record your observations.To observe active sperm, add 1 drop of “dry” sperm to about 100 ml of seawater in a small container. Sperm are best observed using phase contrast, a dark-field technique, or some other type of microscopy that increases contrast or otherwise enhances visibility of very small objects. If you don’t have available phase-contrast optics or a dark-field arrangement on the microscope that you are using, close down the iris diaphragm of the microscope’s condenser. This will add some artificial contrast that will facilitate these observations. The newly shed echinoid egg is surrounded by a transparent jelly coat that has a refractive index similar to that of seawater. If you wish to observe the eggs’ jelly coats, mix a drop of India ink with a small quantity of seawater and observe eggs in the suspension. Since the India ink particles do not penetrate the jelly coat, each egg should appear to be surrounded by a clear area (the jelly coat) containing no ink particles.

Fertilization 1. The fertilization procedure involves mixing drops of a diluted sperm suspension with eggs in seawater. A dilute sperm suspension is prepared by placing 1 drop of the undiluted (“dry”) sperm in a beaker containing 100 ml of seawater. Mix with a clean pipette to obtain a uniform, faintly cloudy suspension.The “dry” sperm suspension is quite viscous so it is sometimes difficult to control the amount transferred to the beaker of seawater. The final diluted sperm suspension should be only slightly cloudy, not milky, in appearance, because use of an excessively dense sperm suspension can lead to polyspermy. Polyspermy, the entry of more than one sperm into an egg, results in abnormal, arrested development. Since sperm activation requires several minutes, the dilute sperm suspension should be allowed to stand for 5 to 8 minutes before use.

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Laboratory 1





TABLE 1.1 Timing of Some Fertilization Events 0 seconds 30– 40 seconds 35– 50 seconds

60– 70 seconds 65– 80 seconds 2 minutes 5 minutes

Insemination Exocytosis of cortical granules Initiation of fertilization membrane elevation (5–10 seconds following cortical granule exocytosis) Completion of cortical granule exocytosis Completion of fertilization membrane elevation Hyaline layer formed Fertilization membrane hardened

Transfer several drops of washed eggs to a container with about 100 ml of clean seawater.A thinly scattered layer of eggs on the bottom of a beaker or finger bowl is an appropriate egg density. Eggs, when they have settled, should cover no more than about one-third of the area of the bottom of the container.Then, add 2 or 3 drops of the dilute sperm suspension to the beaker or dish containing the eggs. Mix the sperm and eggs by stirring very gently with a clean pipette. 2. Transfer a sample of the suspension of eggs and sperm to a slide and observe it with a compound microscope.The most conspicuously observable event is the formation of the fertilization membrane (fertilization envelope), which is a visible indication that the union of sperm and egg has occurred. If fewer than two-thirds of the eggs display fertilization membranes after 2 or 3 minutes, add several more drops of dilute sperm suspension and stir gently. Repeat if necessary. The fertilization membrane gradually rises away from the surface of the egg, beginning in the area of sperm penetration and spreading outward around the entire egg. Fertilization membrane elevation is usually complete within 1 to 2 minutes. Elevation of the fertilization membrane is associated with the exocytosis of the contents of cortical granules that are located just below the surface of the egg.The fertilization membrane, which initially is thin and soft, hardens within a few minutes after its elevation.The translucent hyaline layer that forms just over the surface of the egg also develops within a few minutes. These processes are quite temperature dependent, and their timing varies among species, but table 1.1 shows the approximate sequence of events following the addition of sperm to eggs. Note that several of the listed processes cannot be observed with a light microscope. 3. Since the fertilization membrane is elevated rather quickly, you may miss its formation and wish to use another technique to observe the process directly. One means of direct observation consists of placing a drop of eggs and a drop of sperm side by side on a slide. With the microscope focused on the eggs, the 2 drops can be connected by pushing them together with a needle. This technique will permit you to observe sperm swarming around the eggs. Another method that facilitates direct observation involves placing a pair of 1-mm thick threads of modeling clay parallel to one another on a microscope slide.After laying down the clay, place a drop of unfertilized eggs on the slide between the two strips of modeling clay and add a coverslip. Focus on a group of eggs and, without moving the slide, add a drop of sperm at one edge of the coverslip.You should be able to observe the arrival of sperm in your field of view and the elevation of fertilization membranes on the eggs as you watch them.You may also be lucky enough to observe an egg in which you can see the fertilization cone form at the point where sperm entry is occurring. If you can produce dark-field or Rheinberg illumination on your microscope, you will find these dark-field techniques especially helpful in making these observations.

Caring for Embryos and Larvae 1. Once you are satisfied that most of the eggs have fertilization membranes, leave the beaker undisturbed until the zygotes have largely finished settling to the bottom.Then pour off the supernatant water and add clean seawater. This step eliminates many of the extra sperm that can degenerate later and foul the water around the developing embryos.

Laboratory 1

3





(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

FIGURE 1.1 Cleavage stages in the sand dollar, Dendraster excentricus. (a) Zygote shortly after fertilization. The fertilization membrane (fertilization envelope) has been elevated. Pigment granules in the jelly coat are visible outside the fertilization membrane. (b) 2-cell stage. (c) 4-cell stage. (d ) 8-cell stage, lateral view. (e) 16-cell stage, vegetal view. Note the presence of four micromeres that were produced during the fourth cleavage. (f ) 32-cell stage, lateral view. Note the cluster of micromeres in the vegetal region at the right of this photo. (g) 64-cell stage, lateral view. (h) Blastula shortly before hatching. (The egg cell in a is approximately 120 ␮m in diameter, and all other photos are printed at the same magnification.) Photos by R. B. Emlet.

4

Laboratory 1





2. Put a loose-fitting aluminum-foil cover over the beaker and set it aside or put it in an appropriate constant temperature chamber until you are ready to make the observations described in the following. Developing embryos and larvae must be maintained at a temperature appropriate for the particular species (See Appendix A, p. 201). 3. When blastulae eventually hatch, you will be able to see them swimming near the surface of the water in the upper part of the beaker. Sometimes shining a flashlight or microscope illuminator through the culture can help you to spot the swimmers. Once a substantial number of blastulae have hatched, pour the swimmers from the upper part of the culture into a clean beaker. Avoid pouring over the unhatched or nonswimming embryos that remain at the bottom of the original culture.These should be discarded so that they do not foul the water in the culture when they die and degenerate.While 100-ml beakers are very useful for some lab manipulations, longerterm cultures should be maintained in 250-ml or larger beakers. 4. Aerate the cultures twice daily by repeatedly and gently pipetting air to the bottom of the cultures. However, you need to be careful to avoid sucking embryos in and out the pipette.This means that once you have expelled air from the pipette into the culture, you should keep the pipette bulb compressed until you have lifted the tip of the pipette above the water’s surface. Cultures also may be aerated with a very slow stream of air bubbles from an air line or an air pump, but this needs to be done very cautiously because vigorous bubbling can damage swimming embryos and larvae. 5. If you wish to extend the time that you maintain cultures and feed the developing larvae, you will eventually need to exchange the water in the cultures. A convenient technique for water exchange is described on p. 13 in Laboratory 2. For some hints on feeding larvae, see Appendix A (p. 208).

Embryonic Development 1. Cleavage of echinoid echinoderm embryos (fig. 1.1) is holoblastic; that is, the entire cell is divided at cytokinesis during each cleavage division. The first cleavage, which is meridional, produces a two-cell embryo. The second cleavage division is also meridional and yields a four-cell embryo. In the third cleavage, the plane of division is at right angles to the first two cleavages and the product of the division is an eight-cell embryo with upper and lower quartets of cells. During the fourth cleavage, the four blastomeres of the upper (animal) quartet divide equally to form a single tier of eight medium-sized cells called mesomeres. However, the divisions of the other four (vegetal) blastomeres are very unequal, producing a middle tier of four larger macromeres and a lower tier of four much smaller micromeres that lie at the vegetal pole of the embryo.As cleavage proceeds, the embryo becomes organized as a single-layered, hollow ball of cells surrounding a cavity that is known as the blastocoel.The embryo at this stage of development is called a blastula. 2. An embryo will hatch as a blastula, and just before hatching, the blastula begins to rotate within its fertilization membrane as a result of ciliary activity. Hatching involves enzymatic digestion of the membrane, which becomes fainter in appearance as it thins. Eventually, the membrane opens at one side, allowing the blastula to roll out. The time from fertilization to hatching varies among species and also is strongly influenced by the temperature at which development takes place. Once embryos have hatched, they are harder to observe because they swim more or less continuously. Some individuals eventually become “beached” near the edge of drops on slides, but it is also possible to take active steps to slow or stop them. If Poly-L-Lysine-coated coverslips or slides are available,embryos will settle out of a drop onto the coated surface and be held still while you observe them.Alternatively, embryos and larvae can be anesthetized.To do this, mix about 8 drops from the culture with 1 drop of saturated MgCl2 solution in a small container before transferring the embryos or larvae to a slide for observation. 3. Gastrulation is a set of processes by which embryonic cells are repositioned as the basic body organization of the larva is established. It is somewhat difficult to observe details of gastrulation because embryos swim actively during these stages of development, but patient viewing of several embryos will permit you to see at least some of the interesting cellular activities that are involved. Review a description of gastrulation in your textbook or in references provided in the lab before observing various gastrulation processes for yourself. Just before gastrulation begins, one side of the blastula wall flattens and thickens to form a prominent vegetal plate. Cells of the vegetal plate play major roles in gastrulation. Gastrulation begins with the separation of the primary mesenchyme cells (fig. 1.2a) from the vegetal plate portion of the blastula wall and their subsequent inward movement (ingression). These cells are among the descendants of the micromeres that originally were established during the fourth cleavage. The primary mesenchyme cells move over the inner surface of the blastula wall to new positions where they form clusters of cells. Cells in these clusters will begin to assemble the crystalline spicules of the larval skeleton.

Laboratory 1

5


(a)




(b)

(c)

FIGURE 1.2 Gastrulation in sea urchin embryos. (a) Mesenchyme blastula of a sea urchin embryo. Primary mesenchyme cells have entered the blastocoel and are beginning to migrate over the blastocoel’s inner surface. Note the vegetal plate made up of relatively taller cells. (b) Scanning electron micrograph (SEM) of an external view of a gastrulating sea urchin embryo showing the invagination of the vegetal plate at the beginning of archenteron formation. Cilia have been removed from the surface of this embryo. (c) Composite of SEM photos showing the interior of an embryo during archenteron invagination. Note the primary mesenchyme cells migrating over the surface of the blastocoel wall. (a) is a differential interference contrast photo by R. B. Emlet; (b and c) are SEM photos by John B. Morrill. (c) is from Morrill and Santos, 1985, in R. H. Showman and R. M. Sawyer, eds., The Cellular and Molecular Biology of Invertebrate Development, Univ. S.C. Press.

mouth

anus skeletal rod

(a)

(b)

FIGURE 1.3 (a) Midgastrula stage of sea urchin development. As the archenteron lengthens, its wall thins. Some of the secondary mesenchyme cells at the tip of the archenteron have filopodia extending to the blastocoel wall. (This gastrula is approximately 120 ␮m long.) (b) A sea urchin pluteus larva. Propelled by cilia, a pluteus swims with its mouth and arms directed upward. This pluteus measures approximately 200 ␮m from apex to arm tip. (a) is a differential interference photo by Jeff Hardin, Dept. of Zoology, University of Wisconsin; (b) is a light micrograph by John B. Morrill.

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Laboratory 1





Movements of the primary mesenchyme cells involve rather complex individual cell behavior, but the shape changes that initiate development of the archenteron ( primitive gut) depend upon the collective activity of a number of cells.Archenteron development begins with the invagination (inward sinking or “in-pocketing”) of the vegetal plate (fig. 1.2b and c).You should be able to observe some parts of this invagination process. Once it has been established by invagination, the archenteron lengthens, and its wall thins appreciably.The final phase of extension involves activity of the secondary mesenchyme cells, a group of cells that become evident at the archenteron’s tip when it has extended about halfway across the blastocoel. Some secondary mesenchyme cells extend long, thin projections known as filopodia that reach out to touch various sites on the inside of the blastocoel wall (fig. 1.3a). It is often possible to observe extended filopodia. Some of the contacts made by filopodia are temporary, and the cells retract these filopodia, but other filopodia remain attached if they have contacted the region of the oral ectoderm where the mouth will form. These attached filopodia contract, pulling the tip of the archenteron over into contact with the oral ectoderm, and the larval mouth develops in the contact area. The oral surface becomes flattened, giving the embryo an angular appearance that characterizes the prism stage of development. The angularity of prism-stage embryos contrasts distinctly with the spherical shape of the blastula and gastrula stages. Skeletal spicules are clearly evident in prism-stage embryos. Over the next day or two, you will be able to observe the differentiation of the pyramid-shaped, four-armed pluteus larva. First, two arms (the postoral arms), and slightly later, two more (the anterolateral arms) are extended (fig. 1.3b). These arms, along with the main portion of the body of the pluteus, are supported by skeletal rods whose development began with formation of the skeletal spicules. If you have a microscope with polarizing optics, it would be interesting for you to examine the developing skeleton using polarized light. A pluteus larva swims with its arms directed upward and its beating cilia set up currents that sweep small food particles into its mouth. Observe as much detail of gut structure, skeleton organization, and other features of the pluteus larva as time permits. For example, differentiation of esophagus, stomach, and intestine can easily be seen. Watch for muscular contractions in the digestive tract. Pluteus larvae will swim for some time but usually will not develop beyond the four-armed stage unless they are fed. Unfed larvae eventually starve, fall to the bottom of the culture container, and degenerate. Further development can be observed only if larvae are fed, and feeding usually requires availability of cultures of appropriate marine algae. (See comments on feeding pluteus larvae in Appendix A.)

Laboratory 1

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Materials EQUIPMENT

GLASSWARE

AND

Beakers, various sizes, but 100-ml and 250-ml beakers are especially useful Clean syringes and hypodermic needles Clean Pasteur pipettes Petri dishes Aluminum foil or parafilm Microscope slides and coverslips Depression slides Compound microscope Dark-field stop for microscope, if available (see p. 206) Rheinberg filter for microscope, if available (see p. 207) Polarizing optics, if available (see p. 207) Poly-L-Lysine-coated coverslips or slides (optional) India ink (optional) Modeling clay SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) 0.5 M KCl solution Saturated MgCl2 solution (optional) ANIMALS Sea urchins or sand dollars

Some Useful Information Sources VIDEO —A DOZEN EGGS

This video includes several nice video sequences of sea urchin development photographed by Rachel Fink, Mount Holyoke College, and Seth Ruffins and Charles Ettensohn, Carnegie Mellon University. The video was produced under the auspices of the Society for Developmental Biology and is available from: Sinauer Associates, Inc., P. O. Box 407, 23 Plumtree Road, Sunderland, MA 01375-0407 WEBSITES www.stanford.edu/group/Urchin/ This is a website developed by teachers and Stanford University researchers that contains information about sea urchin development and techniques for studying urchins. http://sdb.bio.purdue.edu/ This is the very useful website of the Society for Developmental Biology. It includes a number of links to sites containing information about echinoderm development. http://worms.zoology.wisc.edu/urchins/SUmainmenu.html This website prepared by Jeff Hardin of the University of Wisconsin contains a number of instructive illustrations and micrographs as well as a lot of useful information about sea urchin development. CD-ROM—SEA URCHIN EMBRYOLOGY

This CD-ROM gives access to the same material available on the urchin website listed previously. It is inexpensive and is available from Hopkins Marine Station, Stanford University, Dept. of Biol. Sci., Pacific Grove, CA 93950-3094. CD-ROM—VADE MECUM: AN INTERACTIVE GUIDE

TO

DEVELOPMENTAL BIOLOGY

This CD-ROM, prepared by Mary Tyler and R. N. Kozlowski of the University of Maine, contains much information about development and techniques, including a good deal of information about echinoderm development. It is available from Sinauer Associates, Inc., P. O. Box 407, 23 Plumtree Road, Sunderland, MA 01375-0407.

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2. Experimental Investigation of Early Development in Echinoderms


L A B O R A T O R Y

2 Experimental Investigation of Early Development in Echinoderms

Literally thousands of experimental investigations in developmental biology, as well as in cell and molecular biology, have been done using the gametes and embryos of echinoid echinoderms.The results of those many studies have greatly expanded knowledge in several areas of biology. It is quite easy to obtain sea urchin and sand dollar gametes, and the requirements for culturing embryos, at least through early phases of development, are fairly simple. Thus, a number of relatively simple experiments on these cells and embryos can be done in the teaching laboratory. In this laboratory, you will have opportunity to conduct several of these experiments.

SPERM CLUMPING Early in the twentieth century, F. R. Lillie studied sea urchin gametes extensively. Among his many observations was the discovery of a sperm clumping response induced by extracts from eggs’ jelly.When unfertilized eggs are separated from the seawater in which they have been standing for an extended period and the supernatant (or “egg water”) is added to a sperm suspension, the sperm form clusters. Later, the clusters disperse, but sperm that have reacted in this way are no longer capable of participating in fertilization interactions. These observations became the basis for the fertilizin-antifertilizin hypothesis of sperm-egg interaction developed by Lillie and others.This hypothesis suggested that “egg water” contains a soluble agglutinating factor from the jelly surrounding mature sea urchin eggs that might play a role in fertilization or possibly in polyspermy prevention. The fertilizin-antifertilizin hypothesis was later reexamined when the responses of sperm in “egg water” were analyzed further (Collins 1976; see “Suggestions for Further Investigation of Echinoderm Development” on p. 26). It was observed that “egg water” induces sperm to undergo the acrosome reaction, including extrusion of the acrosomal process, and that the sperm clumping seen in “egg water” isn’t an agglutination process such as that caused by antibodies or other cross-linking factors. Rather, sperm actively swim into rosettelike clusters where their acrosomal processes adhere. It is now clear that the inability of sperm previously subjected to “egg water” to participate in fertilization is due to the condition of their acrosomes rather than to an agglutinating factor from egg jelly coats that blocks binding sites.

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Techniques 1. The jelly coat surrounding sea urchin eggs will slowly dissolve in seawater, so seawater in which eggs have been stored for a few hours can be used as “egg water” in making these observations.To more quickly prepare an “egg water” solution, vigorously shake a sample of unfertilized eggs in about 30 ml of seawater in a covered test tube. (Shaking helps to dissolve the jelly coats.) Filter the test tube’s contents and collect the filtrate (“egg water”). Place a drop of “egg water” on a slide or in a watch glass. Prepare a milky, diluted sperm suspension and add one drop of it to the “egg water.”Within 1 minute, the sperm suspension will take on a granular, or flocculent, appearance. Microscopically, it can be seen that the sperm are clustered. 2. You can test the functional capacity of the previously clumped sperm by tipping the suspension off the slide into a sample of fresh eggs on a depression slide. Check for fertilization membrane formation and, if you do observe any, determine the fertilization percentage. 3. If the fertilization percentage is very low, what conclusions might you draw? What are some alternative explanations of the results? What additional experimental step could you take to provide control results that would help to clarify interpretation of the results? Do the additional test. What are the results? What conclusions might you draw?

Materials EQUIPMENT Basic equipment and supplies for sea urchin and sand dollar experiments as listed in Laboratory 1 Test tube and stopper or cover for the tube during shaking Filter paper and funnel SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) 0.5 M KCl solution LIVING MATERIAL Sea urchins or sand dollars

ARTIFICIAL PARTHENOGENESIS The process of fertilization involves a complex set of cellular responses and interactions, and many aspects of the egg cell’s physiology change as a result of the activation occurring during and after sperm contact and entry. Some of these changes can be initiated artificially by various treatments. Egg activation without sperm contact is called parthenogenesis, and artificial parthenogenesis in sea urchin eggs has been investigated by many biologists since it was first studied by Oscar and Richard Hertwig during the 1880s. Many experimental treatments have been found to cause some, or even many, of the activation responses to occur in sea urchin eggs. In this experiment, you will have opportunity to investigate the effects of one of these treatments— immersion in seawater made hypertonic to egg cells by addition of 30 grams of sodium chloride per liter. This procedure is one recommended by the great early twentieth-century American developmental biologist Ethel Browne Harvey, who, near the end of her career, summarized and reinvestigated many of the experimental procedures used to investigate sea urchin development up to the 1950s. Techniques 1. Transfer a sample of freshly shed eggs to hypertonic seawater (HSW) by pipette and allow them to settle to the bottom of the beaker or dish. 2. After 20 minutes, wash the eggs by pouring off as much of the HSW as you can and resuspending them in normal strength seawater. (Since eggs of different species, and even different batches of eggs of a single species, respond differently, you might consider trying several additional treatment times.)

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Laboratory 2


3. centage 4. 5.




Check for fertilization membrane elevation, and if fertilization membranes are present, determine the perof eggs that have them. After 10 minutes, wash the eggs again as in Step 2. Check again for fertilization membranes and percentage of eggs possessing them.

Analysis of Results 1. If fertilization membranes have been found, continue to observe the culture at intervals and be alert for signs of further development. If you detect developmental progress such as cleavage divisions, compare timing with control embryos. 2. Follow your culture long enough to observe hatching and gastrulation should they occur in any embryos developing from parthenogenetically activated eggs. However, parthenogenetically activated development only rarely proceeds beyond completion of a few cleavages.

Materials EQUIPMENT Basic equipment and supplies for sea urchin and sand dollar experiments as listed in Laboratory 1 SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) 0.5 M KCl solution Hypertonic seawater (seawater with 30 g NaCl added per liter) LIVING MATERIAL Sea urchins or sand dollars

BLASTOMERE SEPARATION One of the most famous experiments in the early history of developmental biology was performed on the sea urchin embryo. In 1892, Hans Driesch vigorously shook two-cell-stage sea urchin embryos in a test tube of seawater until some of the blastomeres (cleavage cells) separated from one another. Driesch followed the development of these isolated individual cells and found that, in at least some cases, such cells could give rise to quite normally proportioned, though undersized, larvae. Driesch concluded that each of the blastomeres at the two- and four-cell stages has the capability to develop into a whole embryo. Though the developmental capacity of early cleavage cells may not be quite so dramatic or extensive as Driesch thought it to be, his results are still regarded as an important milestone in the investigation of animal development. Driesch’s procedure is tricky to repeat, but in this laboratory, you will have an opportunity to investigate the results of blastomere separation using another technique. The technique that we will use may not be quite so rough as Driesch’s vigorous shaking, but it is harsh, and appropriate controls should be added to your experimental design if you have the opportunity to do so.Whether or not you carry them out, think about appropriate sets of control observations for each of the steps in the technique that follows. Techniques 1. Place a sample of sea urchin eggs in a 10 mM solution of p-aminobenzoic acid (PABA) in seawater.Then proceed with fertilization by adding a sperm suspension to the eggs, using the same techniques that you used in Laboratory 1. PABA prevents the hardening of the fertilization membrane that would normally occur within a few minutes after its formation and facilitates later removal of the membrane. 2. After zygotes have settled to the bottom, gently pour off the PABA solution and replace it with calciumfree seawater containing 50 ␮⌴ EGTA.

Laboratory 2

11



(a)



(b)

(c)

FIGURE 2.1 Development of twinned embryos of the sea urchin, Lytechinus pictus. (a) Twinned embryos after two cleavage divisions. (These cells are slightly flattened. Each is about 60 ␮m in diameter.) (b) Twinned early blastulae (magnification same as in a). (c) A dark-field photo of a control pluteus (right ) and a small pluteus that developed from an embryo twinned at the two-cell stage. (The control pluteus is about 200 ␮m long.) Photos by L. G. Johnson.

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Laboratory 2





3. Pour off the supernatant after the zygotes once again have settled and add fresh calcium-free seawater solution. Repeat this washing step. 4. Use a simultaneous control culture to monitor developmental progress and sample your experimental culture only when cytokinesis of the first cleavage is proceeding in the control culture. 5. Following completion of the first cleavage division, pour the embryos through a 70-␮m mesh Nitex filter. Check a drop sample from your culture to see what percentage of the embryos have been separated into individual blastomeres. If most are still in the two-cell configuration, it may be necessary to repeat this filtering step. If this second pass through the filter does not result in blastomere separation, it may be necessary to use a filter with a smaller mesh size. 6. Once blastomeres have been successfully separated, allow the cells to settle and pour off the supernatant. Resuspend in normal seawater and proceed to the Analysis of Results section that follows.

Analysis of Results 1. Observe cleavage in your culture over the next few hours if you are able to do so. Watch for differing numbers of cells in the embryos. After two more cleavage divisions,“twinned” embryos will consist of four cells, while embryos whose blastomeres were not separated will contain eight cells. On the basis of what you know about results of the fourth cleavage in control sea urchin embryos, what cell-size classes might you expect in “twinned” embryos completing a third division subsequent to blastomere separation at the two-cell stage? 2. If you can return to check your experimental culture when the embryos have hatched and are swimming, try to determine whether you have embryos of more than one size in your culture. If you can, measure several embryos in your culture and make comparative measurements of embryos from a control culture. 3. If you can maintain your cultures long enough to do so, look for pluteus larvae in your experimental culture. Check for size differences among the plutei that you find and compare them with plutei from control cultures (fig. 2.1).

Materials EQUIPMENT Basic equipment and supplies for sea urchin and sand dollar experiments as listed in Laboratory 1 Nylon filters with appropriate mesh size (see Appendix A) Cylinders to hold filters, for example, 50-ml syringes with tip end cut off or 7.5-cm (3-inch) or 10-cm (4-inch) pieces of PVC pipe with an inside diameter of about 2.5 cm (1 inch) SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) 0.5 M KCl solution 10 mM p-aminobenzoic acid (PABA) in seawater (use the sodium salt of PABA, not the free acid) Calcium-free seawater containing 50 ␮⌴ EGTA Hypertonic seawater (seawater with 30 g NaCl added per liter) LIVING MATERIAL Sea urchins or sand dollars

Exchanging Seawater in Cultures Once embryos have hatched and are swimming, it is very difficult to exchange the water in the culture simply by pouring.The following technique permits removal of a large percentage of the water from a culture while leaving the swimming embryos or larvae behind. It also can be used when the medium around early, nonswimming embryos must be changed and there is not time to allow them to settle to the bottom of the container. 1. Submerge the filter-covered (35 to 40 ␮m mesh size) end of a piece of PVC pipe in the culture. It may take a moment for the filter to be wetted by the seawater. ( You can pre-wet the filter using a squirt bottle filled with seawater.)

Laboratory 2

13





PVC pipe

turkeybaster

embryo culture

filter

FIGURE 2.2 Diagram showing the use of a turkey baster and a filter-covered pipe to remove liquid from a beaker containing a suspension of embryos or larvae. The filter should have a mesh size between 35 ␮m and 40 ␮m (see Appendix A).

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Laboratory 2





2. Squeeze the bulb of an ordinary kitchen turkey baster and insert the baster’s tip into the piece of pipe (fig. 2.2). 3. Gently draw seawater into the baster by slowly releasing pressure on the bulb. This removes fluid from the culture while the filter at the end of the pipe blocks passage of embryos or larvae so they are left behind. 4. Discard the water in the baster and repeat Steps 1–3 until you have removed most of the water from the culture container. 5. Gently pour replacement water into the culture to the desired level.

DEVELOPMENTAL EFFECTS OF REDUCED CALCIUM ION CONCENTRATION Calcium ions are important in some cellular responses to external signals, in the intracellular regulation of various processes, and also are required for some cell movements. Therefore, it is not surprising that calcium ions must be present at or near the normal seawater concentration for development to proceed normally. In this experiment, you will have an opportunity to experiment with the effects of reduced calcium ion availability. Techniques 1. Concentrate a culture of embryos between the hatching blastula and very early mesenchyme blastula stages by drawing off almost all of the seawater in the beaker using a filter-covered piece of PVC pipe and a turkey baster (see p. 13). Be especially careful not to use strong suction as you near the end of this process because embryos can be damaged if they are drawn forcefully against the filter. 2. Fill the beaker with calcium-free seawater.Then reduce the volume again and refill the beaker with calciumfree seawater once more. 3. Observe samples of embryos from control (ordinary seawater) and experimental cultures at intervals over the next several days. Make morphological comparisons and developmental rate comparisons between control embryos in ordinary seawater and embryos developing in the calcium-free seawater. Watch especially for differences in gastrulation movements.

Materials EQUIPMENT Basic equipment and supplies for sea urchin and sand dollar experiments as listed in Laboratory 1 PVC pipe with filter-covered end—mesh size 35 to 40 ␮m Turkey baster SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) Calcium-free seawater containing 50 ␮⌴ EGTA LIVING MATERIAL Sea urchins or sand dollars

CILIA REGENERATION Development of larval cilia is an interesting subroutine in the echinoderm developmental program. Cilia are essential for normal swimming and thus vital to the life and further development of the larva, but, like most parts of the larval body, they are temporary features in the developmental sequence leading to the adult. It is especially interesting that developing echinoid echinoderms can regenerate even large numbers of lost cilia and that they do so without significant delay to general developmental progress. This ability may represent an adaptive response to possible cilia loss in the natural environment. Larvae can be completely deciliated by one of several chemical treatments. Thus, they can serve as an experimental system for investigating synthesis and assembly processes involved in cilia development. In this experiment, you will have opportunity to observe the process of deciliation following very brief hypertonic shock and to follow the progress of cilia regeneration.

Laboratory 2

15





Techniques 1. Concentrate a culture of swimming blastula or later-stage embryos by drawing off almost all of the seawater in the beaker using a filter-covered piece of PVC pipe and a turkey baster (see p. 13). Be especially careful not to use strong suction as you near the end of this process because embryos can be damaged if they are drawn forcefully against the filter. 2. Transfer some of the embryos to a second beaker. They will serve as controls during the process. Then add normal seawater to the control beaker and add hypertonic seawater to the original beaker. 3. The total period of exposure to hypertonic seawater should be only about 30 seconds so you need to begin removing the hypertonic seawater almost immediately. Remove almost all of the water in the beaker using a filter pipe and turkey baster. Immediately fill the beaker with normal seawater and begin your observations.

Alternative Method 1. Concentrate a culture of swimming blastula or later-stage embryos by drawing off almost all of the seawater in the beaker using a filter-covered piece of PVC pipe and a turkey baster (see p. 13). Be especially careful not to use strong suction as you near the end of this process because embryos can be damaged if they are drawn forcefully against the filter. 2. Transfer some of the embryos to a second beaker containing seawater.They will serve as controls during the process. Fill the beaker with normal seawater. 3. Submerge the tip of another filter-covered piece of PVC pipe in a small quantity of normal seawater in a beaker. Slowly and gently pour the remainder of concentrated culture of swimming blastulae through the pipe so that the embryos are held against the filter inside the pipe. 4. Expose the embryos to hypertonic seawater by slowly lifting the filter-covered pipe and immersing its filter-covered end in hypertonic seawater for 30 seconds. 5. Slowly lift the filter-covered pipe out of the hypertonic seawater, invert it over a beaker containing a little seawater, and gently pour seawater on the filter, thereby washing the embryos out of the pipe into the beaker. Immediately fill the beaker with normal seawater.

Analysis of Results 1. Make a gross comparison between the experimental and control cultures. Do you see evidence of a difference in swimming behavior? Do you observe a tendency for the experimental embryos to settle out of the culture? 2. Collect samples from the experimental and control cultures and examine them microscopically. It is possible to observe differences in ciliation with an ordinary light microscope if the iris diaphragm is closed down, but you will get better results using some form of phase-contrast microscopy or the dark-field or Rheinberg techniques (see p. 206). The Rheinberg technique is especially useful for observing cilia. Do you see differences in swimming? In ciliation? 3. Repeat the observations in Steps 1 and 2 every few minutes for the next 60 to 90 minutes. If you have seen almost complete settling of the experimental embryos, watch for them to resume swimming. Try to correlate resumption of swimming with reappearance of cilia that you might observe directly. How much time do you estimate is required for complete cilia regeneration?

Possible Further Observations 1. You might be able to observe the process of cilia loss directly if you mix a drop of seawater containing some embryos with several drops of hypertonic seawater on a slide. Can you see cilia that have fallen off ? Try to pipette away the hypertonic seawater and replace it with normal seawater. If you succeed in doing so, place the slide on a piece of wet paper towel in a petri dish and cover the dish to reduce evaporation. How many minutes pass before you can see regenerating cilia? How long does it take for these small cilia to double in length? When do the blastulae begin to swim? 2. If you have a culture of embryos that have successfully regenerated their cilia, you might wish to do another experiment to test the ability to regenerate cilia several times in succession.

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Laboratory 2





Materials EQUIPMENT Basic equipment and supplies for sea urchin and sand dollar experiments as listed in Laboratory 1 PVC pipe with filter-covered end—35 to 40 ␮m mesh size (two are needed for the alternative method) Turkey baster SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) 0.5 M KCl solution Hypertonic seawater (seawater with 30 g NaCl added per liter)

Laboratory 2

17







3. Experimental Investigation of Organization and Differentiation in Echinoderms


L A B O R A T O R Y

3 Experimental Investigation of Organization and Differentiation in Echinoderms

Embryos of various echinoderms have been used in literally hundreds of investigations of organization and differentiation during early development. This research has stretched from studies early in the twentieth century that gave rise to the historical double-gradient hypothesis to intense investigation of specific gene expression studies that has continued into this century.This lab introduces several of the types of experiments that have been part of this long series of investigations.

DEVELOPMENTAL EFFECTS OF LITHIUM CHLORIDE The results of the experimental exposure of echinoderm embryos to lithium ions played an important role in development of the historical double-gradient hypothesis of developmental control. Although the double-gradient hypothesis has undergone considerable reevaluation, the sometimes striking effects of lithium ions on major morphogenetic events in echinoderm embryos remain of interest. For example, abnormal development of structures derived from the vegetal area can result in formation of embryos with a proportionately large archenteron or even with an archenteron that bulges outward from the surface rather than invaginating properly into the blastocoel. This phenomenon is called exogastrulation. In light of lithium’s known effects on cellular signaling systems (see Suggestions section), it is not surprising that lithium has significant effects on developmental processes that depend upon cellular interactions. Lithium has been used more recently as a tool for the investigation of major organizational processes in early development. Techniques 1. Use a 60 mM solution of LiCl in seawater to prepare 30 mM and 15 mM solutions as well. 2. Fertilize sea urchin or sand dollar eggs in a 250 ml or larger beaker, using the standard techniques introduced in Laboratory 1. Once you are satisfied that most of the eggs have fertilization membranes, swirl the culture and pour off samples into several smaller beakers. Cover each culture loosely with aluminum foil and leave the beakers undisturbed until most of the zygotes have settled to the bottom. Then pour off as much of the supernatant water in each culture as you can, and replace it with normal seawater. Set the cultures aside or return them to an appropriate culture chamber. 3. When the embryos reach the two-cell stage, pour off as much of the supernatant water in each culture as you can and replace it with seawater containing LiCl at each of the concentrations that you have chosen to test. Keep one culture as a control in which you replace the discarded seawater with normal seawater.

19





4. Sometime between the time when hatching begins and the mesenchyme blastula stage, replace the lithium-containing seawater in the cultures with normal seawater. (It is probably best to transfer Lytechinus species while they are hatching; transfer other species closer to the mesenchyme blastula stage.) If you have enough separate culture containers, leave some embryos in lithium solutions. To replace the lithium-containing seawater with normal seawater, concentrate each culture of embryos by drawing off almost all of the seawater in the beaker using a filter-covered piece of PVC pipe and a turkey baster (see p. 13). Be especially careful not to use strong suction during this process because embryos can be damaged if they are drawn forcefully against the filter. Replace the discarded lithium-containing seawater with normal seawater. Repeat the process to further wash the embryos.

Analysis of Results 1. Observe samples of embryos from a control culture (embryos maintained continuously in normal seawater) and each of the experimental cultures at intervals over the next several days. 2. Make morphological comparisons and developmental rate comparisons between control embryos and embryos in the several experimental groups. Do you observe differences in gut (archenteron) development between experimental and control embryos? Among the experimental embryos, is there evidence of guts that are larger in proportion to body size than are those of control embryos? Are there any embryos that have a developing gut projecting outward rather than into the interior (exogastrulae)?

Materials EQUIPMENT Basic equipment and supplies for sea urchin and sand dollar experiments as listed in Laboratory 1 PVC pipe with filter-covered end—mesh size 35 to 40 ␮m SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) 0.5 M KCl solution 60 mM LiCl solution in seawater LIVING MATERIAL Sea urchins or sand dollars

DEVELOPMENTAL EFFECTS OF NICKEL IONS Development of the larval skeleton involves an easily observed series of events.After primary mesenchyme cells enter the blastocoel, they migrate to establish a ring of cells. At two points within the ring, primary mesenchyme cells aggregate in two ventrolateral clusters and form the syncytial clumps that produce the first two skeletal spicules.This cell activity, along with expression of specific genes involved in differentiation of ventral and dorsal ectoderm, is involved in converting the previously radially patterned embryo into the bilaterally arranged prism-stage larva. Some ions, such as nickel, are known to affect this characteristic patterning process. In this experiment, you’ll be experimenting with this nickel effect. Techniques 1. Use a 2 mM stock solution of NiCl2 in seawater to prepare 1 mM and 0.1 mM solutions in seawater by dilution. If your experiments are limited by beaker availability or culture space and you can test the effect of only one concentration of nickel ions, consult classmates who test the other two concentrations so that you have opportunity to share observations of the effects of the several concentrations. 2. Fertilize sea urchin eggs in a 250 ml or larger beaker, using the standard techniques introduced in Laboratory 1. Once you are satisfied that most of the eggs have fertilization membranes, swirl the culture and pour off samples into several smaller beakers. Leave the beakers undisturbed until most of the zygotes have settled to the bottom. Then pour off as much of the supernatant water in each culture as you can and replace it with seawater containing NiCl2 at each of the concentrations that you have chosen to test. Keep one culture as a control

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Laboratory 3





in which you replace the discarded seawater with normal seawater. Cover each culture loosely with aluminum foil and set the cultures aside or return them to an appropriate culture chamber. 3. Leave the developing embryos in the nickel-containing seawater until an early gastrula stage when you can observe some evidence of archenteron invagination. Concentrate each culture of gastrula embryos by drawing off almost all of the seawater in the beaker using a filter-covered piece of PVC pipe and a turkey baster (see p. 13). Be especially careful not to use strong suction during this process because embryos can be damaged if they are drawn forcefully against the filter. Replace the discarded nickel-containing seawater with normal seawater. Listen carefully for instructions regarding disposal of the nickel-containing solutions. 4. Repeat this washing step at least twice and refill the beakers with normal seawater each time. Listen carefully for instructions regarding disposal of the nickel-containing solutions. 5. Re-cover the beakers and maintain the cultures for further observation.

Analysis of Results 1. Periodically collect samples from the experimental and control cultures and examine them microscopically. Compare the developing embryos with embryos in your control culture. 2. Is the clustering of primary mesenchyme cells similar in the experimental and control cultures? When two skeletal spicules form in your control embryos, do you see evidence of spicule formation in the experimental cultures? Polarizing optics are very useful in making these observations if you have them available. If you do not have polarizing optics available, close down the iris diaphragm of your microscope’s condenser to provide added artificial contrast. 3. When you observe definitely bilaterally symmetrical prism-stage embryos in your control cultures, carefully examine embryos from your experimental cultures.Check for differences in continuing skeletal spicule growth and patterns of archenteron growth. Does the archenteron in the nickel-treated embryos bend to contact a flattened oral ectoderm area? Do the nickel-treated embryos appear to be bilaterally symmetrical?

Materials EQUIPMENT Basic equipment and supplies for sea urchin and sand dollar experiments as listed in Laboratory 1 PVC pipe with filter-covered end—mesh size 35 to 40 ␮m Turkey baster SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) 0.5 M KCl solution 2 mM solution of NiCl2 in seawater; Please consult your institution’s Hazardous Material Officer for instructions regarding disposal of nickel-containing solutions

ENZYME SYSTEM DIFFERENTIATION A number of developmental biologists are investigating patterns of gene expression in echinoderm embryos and larvae. They detect specific gene activation in particular areas of embryos by techniques such as in situ hybridization, using specific probes for gene expression. An inexpensive and relatively simple approach to studying gene activation is to detect enzyme systems characteristic of areas or structures in embryos and larvae as they become functionally differentiated. One such enzyme system is the alkaline phosphatase that is specifically expressed in the developing gut.You will have an opportunity to test for alkaline phosphatase activity in this experiment. Techniques Preparing for staining: 1. Rear sea urchin or sand dollars to the gastrula, prism, and early pluteus stages using the techniques introduced in Laboratory 1.

Laboratory 3

21




FIGURE 3.1 An early pluteus larva stained with an indicator that is converted from colorless to colored in conjunction with reactions catalyzed by alkaline phosphatase. The colored substance indicates sites where genes for the alkaline phosphatase enzymes have been expressed and the enzyme system is functioning actively.

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Laboratory 3






2. Concentrate an embryo or larva culture to a very small volume by drawing off almost all of the seawater in the beaker using a filter-covered piece of PVC pipe and a turkey baster (see p. 13). Be careful not to use strong suction as you near the end of this process because embryos can be damaged if they are drawn forcefully against the filter. 3. Position several coverslips coated with Poly-L-Lysine near the edge of a petri dish cover or inverted bottom set on a dark background. Shine a bright light through the culture that you have concentrated so that you can see the swimming individuals clearly. Draw as many of them as you can into a Pasteur pipette. 4. Transfer a couple of drops from the pipette to each coated coverslip. Be careful to avoid adding so much seawater that it flows off the edge of the coverslip. Wait several minutes until the embryos have begun to settle and adhere to the coated coverslip. Shine a bright light on the coverslip so that you can see the swimming individuals clearly.Then carefully draw off seawater from the coverslip, using a clean Pasteur pipette.As the volume of seawater on the coverslip decreases, more individuals will tend to stick to the surface. 5. If there are ten or twelve individuals stuck to the coverslip, proceed to the staining procedure that follows. If there are only five or six or fewer individuals on the coverslip, repeat Steps 3 and 4 before beginning staining. It is important to have a number of individuals on the coverslip because some will be lost during transfers in the staining process.

Staining procedure: 1. At each step in the enzyme reaction staining procedure, make transfers carefully. Hold coverslips by their edges and slide them very slowly into each staining jar. Remove coverslips from staining jars with similar caution because individuals are most likely to be washed off the coverslips as they pass through the surface films of the various solutions. It is wise to mark a front on each staining jar and transfer the coverslips consistently so that the embryo side is always directed forward. 2. Place the coverslips in ice-cold methanol (4° C or colder) in a staining jar and put them in the refrigerator for 5 minutes. 3. Carefully remove each coverslip from the jar, hold it vertically, allowing the methanol to drain by touching the bottom edge to absorbent paper (paper towel or blotting paper). 4. Transfer the coverslips to ice-cold seawater,* leave for 30 seconds, withdraw them, drain them by blotting the edges as in Step 3. 5. Repeat Step 4 by transferring the coverslips to a second staining jar containing ice-cold seawater. Drain and blot. 6. Transfer the coverslips to pH 9.1 buffer (room temperature) for 1 minute. Drain and blot. 7. Transfer to alkaline phosphatase substrate solution (room temperature) for 25 minutes.** Drain and blot. 8. Transfer to phosphate buffered saline (PBS) for 3 minutes.*** Then mount on a microscope slide (make certain that the embryo/larva side of the coverslip is toward the slide) and seal the edges.

Analysis of Results 1. Examine each slide carefully, looking for the bluish purple color that indicates alkaline phosphatase activity (fig. 3.1).

*Replace the seawater in the first seawater-containing staining jar after each use. **You may need to vary the time of exposure to alkaline phosphatase substrate depending on your results. ***Replace the PBS after each use. It is possible to skip the PBS wash and mount the coverslips immediately after removal from the alkaline phosphatase substrate.You might consider deleting this step if you are having problems with embryos/ larvae washing off coverslips. However, if you delete the PBS wash, reactions will continue, and stain eventually will diffuse out of the gut into other parts of the embryo larva.

Laboratory 3

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Materials EQUIPMENT Basic equipment and supplies for sea urchin and sand dollar experiments as listed in Laboratory 1 PVC pipe with filter-covered end—mesh size 35 to 40 ␮m Turkey baster Pasteur pipettes 22 ⫻ 22 mm coverslips Watchmaker’s forceps Coverglass (coverslip) staining jars Petri dishes Blotting paper or paper towels Microscope slides SOLUTIONS

AND

CHEMICALS

Seawater or appropriate salt mixture (artificial seawater—see Appendix A) 0.5 M KCl solution Methanol pH 9.1 buffer Alkaline phosphatase substrate Phosphate-buffered saline Water-miscible mounting medium or nail hardener or New Skin Liquid Bandage Poly-L-Lysine solution

Useful Information Source http://worms.zoology.wisc.edu/urchins/SUpattern_Ni.html This website, developed by Jeff Hardin of the University of Wisconsin, contains information about development of embryonic patterns, especially the roles of primary and secondary mesenchyme cells, and the effects of nickel ions.

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Laboratory 3



Suggestions for Further Investigation of Echinoderm Development


L A B O R A T O R Y

4 Observations and Experiments on the Living Frog Embryo

The continuous change of form in developing embryos has always provoked interest and curiosity, and amphibian eggs, which are large and fairly easy to collect and observe, long ago became popular subjects for those interested in studying animal development. Early investigators focused their attention on describing the orderly, normal course of developmental events, and a wealth of information was gathered by direct observation of the development of frog, toad, and salamander embryos. Later, this observational and descriptive work began to be supplemented by experimental studies. The distinguished German embryologist, Wilhelm Roux, who studied amphibian development in the 1880s, was one of the first to take an experimental approach to investigation of animal development. From the time that Roux pricked frog cleavage cells with a hot needle, the amphibian embryo has been subjected to a variety of experimental manipulations and to numerous alterations of its environment. In the United States, embryos of the common leopard frog Rana pipiens have been employed in many experimental studies of vertebrate animal development. Rana pipiens embryos are also widely used as subjects in biology courses dealing with development.This laboratory introduces the Rana pipiens embryo and some of the basic techniques used in obtaining and studying frog embryos. We will begin our study of the frog with a descriptive and experimental examination of fertilization and the early stages of development. In addition to uniting the haploid nuclei of the gametes to produce a diploid zygote, fertilization also initiates a complex set of activation responses in the egg. However, in some cases, an egg can develop in the absence of sperm; such development is known as parthenogenesis. The drone-producing egg of the honey bee is an excellent example of naturally occurring parthenogenic development. Other eggs that do not ordinarily develop parthenogenetically can be induced to do so by a variety of experimental manipulations. For example, the eggs of frogs, salamanders, and toads may be stimulated to complete early stages of development in the absence of the nuclear events of normal fertilization if they are inseminated with pretreated sperm that have been irradiated or chemically modified. Such treatment disrupts the genetic material of the sperm but does not interfere with its mobility or the ability of the sperm to penetrate and activate the egg.

33




testis

testis

FIGURE 4.1 The position of the testes in relation to the other internal organs of the frog.

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Laboratory 4






Your initial experience with the embryo of the leopard frog, Rana pipiens, will be a study of normal diploid embryos accompanied by a parallel examination of parthenogenetically activated embryos. An embryo deprived by artificial means of one set of parental chromosomes exhibits markedly impaired development. The African clawed frog, Xenopus laevis, is now widely used in developmental studies. Xenopus zygotes can be obtained readily, and the embryos can be experimentally manipulated as easily as those of the leopard frog. Although the leopard frog is the organism suggested for this laboratory, Xenopus can be substituted. However, there are some differences between the methods used for obtaining and handling Xenopus gametes and embryos and those used with Rana pipiens. Thus, you should consult information about Xenopus development in Laboratory 9 and Appendix C if you are using Xenopus for this laboratory. Techniques Preparation of Sperm Suspension Mature sperm are present in the testes of male leopard frogs throughout the year, although there is a period of relatively lower spermatogenic activity from late June to mid-September. 1. Pith (or anesthetize) a male frog and use a scissors and forceps to dissect out the pair of testes. ( If you are not adept at pithing, that is, destroying the central nervous system of the frog with a needle, then anesthetize the frog in a sealed container with a wad of cotton saturated with ether or chloroform.) The testes are yellowish, ovoid, paired bodies held to the kidneys by means of mesentery folds (fig. 4.1). 2. After removing the testes, roll each testis gently on paper toweling to free the organ of adhering blood and mesentery. Using blunt forceps, mince the pair of testes thoroughly in 10 ml of spring water or in a diluted (10%) Amphibian Ringer’s solution in a finger bowl. Tilt the finger bowl so that the testes can be macerated easily in the pool of fluid at one side of the bowl. 3. Allow the suspension to stand for 15 minutes, during which time the sperm will become active. Place a drop of the suspension on a glass slide and examine under the high-power objective of a compound microscope. Observe the shape of the sperm and check for motility.

Irradiation of Sperm Nuclear damage is one of the demonstrable effects of ultraviolet ( UV ) irradiation of the sperm cell. In particular, the energy of U V radiation induces abnormal bonding of the pyrimidine bases of nucleic acids. Chromatin material of the sperm cell can be disrupted by U V radiation without diminishing the sperm cell’s capacity to enter the egg. A phenomenon observed in cases of U V exposure is that of photoreactivation, in which the effect of the U V irradiation is perceptibly lessened by the presence of intense visible light (for example, overhead illumination).Accordingly, it is advisable to irradiate in a dimly lit room.An inexpensive U V source is a 15-watt germicidal lamp mounted in a fluorescent fixture.The inverse square law operates for U V radiation—the greater the distance, the longer the exposure time required to accomplish the desired effect. 1. Transfer a small quantity of the sperm suspension to each of two petri dishes. The sperm suspension should be spread thinly over the bottom of each dish. Label one petri dish “control” and the other “experimental”. Set aside the control dish. 2. Position the ultraviolet lamp 38 to 40 cm (15 to 16 in) above the table top. Place the uncovered experimental petri dish beneath the lamp and expose the sperm suspension to the rays of the lamp for 15 minutes. Occasionally swirl the sperm suspension gently to ensure equal exposure of all sperm to the rays. Do not expose your skin to direct UV radiation and do not look directly at the lamp.

You are now ready to inseminate eggs with the control and experimental sperm.

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FIGURE 4.2 Two different views showing how to hold the female frog while “stripping” her eggs.

Side view

A

A⬘ gray crescent

View from vegetal pole

B

B⬘

FIGURE 4.3 Formation of the gray crescent in the frog’s egg. A and B represent unfertilized eggs; A⬘ and B⬘ represent eggs shortly after fertilization.

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Ovulation and Fertilization Ovulation can be induced by injecting frog pituitary extracts or fragmented pituitaries into mature, healthy female frogs. The pituitary hormones cause ovulation within 24–48 hours at room temperature (20–24° C) or within 4–5 days at 10° C. Injected female frogs will be available in the laboratory. (Pre-injected females can be purchased from commercial suppliers, or female frogs can be injected to provide eggs when needed in the laboratory.The procedures for inducing ovulation in Rana pipiens are described in Appendix B.) The female frog need not be sacrificed to release her eggs. Eggs are removed from the oviducts by the technique of “stripping.” 1. Hold the female frog with her back against the palm of one hand. Grasp and extend the frog’s hind limbs with your other hand. Position the frog’s back in your hand so that your fingers partially encircle the body just posterior to the forelimbs.The tips of your fingers will come to rest on the ventral surface of the frog (fig. 4.2). 2. Eggs can be forced from the cloaca by initially applying gentle pressure to the anterior part of the body and then progressively closing the hand toward the cloaca region. First, squeeze the female over paper toweling until she releases several eggs. Usually, they will be accompanied by cloacal fluid. Discard the first few eggs issued from the oviduct and wipe the cloacal region dry.Then proceed to strip 100 or more eggs into each of the petri dishes containing the unirradiated or irradiated sperm suspensions. When stripping the eggs, squeeze gently and move the female around over dishes to produce several chains of eggs rather than a single, heaped mound.To assure complete exposure to sperm, repeatedly draw sperm suspension into a clean pipette and squirt the sperm over the eggs. Observe the orientation of the black-pigmented area (animal pole) and the creamy white vegetal pole of the eggs. 3. Allow the eggs to remain in the sperm suspension for 15 minutes. After the 15-minute period, pour the sperm suspensions out of the petri dishes and flood the eggs with spring water (or 10% Ringer’s solution). Note: A mature female frog can release approximately 2000 eggs. A “stripped” female can be stored at 4° C, and she will subsequently yield viable eggs each day for about 4 days.When removed from the 4° C storage each day, she should be allowed to sit at room temperature for 30 minutes to effect temperature equilibration. 4. After fertilization, the egg becomes free to rotate slowly within the space (the perivitelline space) located between the egg itself and the vitelline membrane. Gravity causes the vegetal hemisphere containing the relatively heavier yolk to rotate to the underside. Soon after fertilization, and before the first cleavage begins, a light crescent-shaped area, the “gray crescent,” appears on one side of the egg in the boundary between the animal and vegetal regions (fig. 4.3), although it is often difficult to locate.The gray crescent is formed by the shifting of cytoplasmic components in the egg, and the position of the crescent is related to the penetration path of the sperm cell; the gray crescent appears on the side of the egg opposite the sperm entry point. One of the earliest observations made by embryologists is that the gray crescent establishes the bilateral symmetry of the frog egg. The plane of the first cleavage of the frog egg, in most cases, passes through the gray crescent.Thus, the first cleavage effectively divides the egg into bilaterally equal halves. 5. The jelly envelope, which initially is dense and viscous, absorbs water and swells to several times its original thickness.This thick envelope of jelly, comprised of two or three concentric layers, protects the egg from mechanical injury.The jelly layers swell maximally about 1 hour after eggs enter water. The jelly mass generally sticks to the glass bottom of the petri dishes, so use a clean scalpel or section lifter to free the jelly from the glass, and then gently lift the cluster of eggs from the bottom of the dish. With sharp scissors, cut the mass of eggs into small clusters of 5 to 10 eggs. You need not be hesitant or overly cautious in cutting the mass as it is almost impossible to shear an egg since the jelly-coated eggs are very resistant to mechanical injury. 6. Lift the small egg clusters with forceps and place the clusters in several finger bowls (4 inches in diameter) containing spring water (or 10% Ringer’s solution). Development will proceed well with 15 to 20 eggs in 250 ml of solution, but more crowded conditions generally should be avoided if it is practical to do so. To reduce evaporation, cover the finger bowls with loose aluminum-foil covers. No change of solution is required

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AP

(b) VP (a)

(c) FIGURE 4.4 Scanning electron micrographs of frog embryo cleavage stages. (a) First cleavage division. The cleavage furrow deepens more quickly in the animal pole (AP) than in the yolky vegetal pole (VP). (b) The 8-cell stage. The size difference between the smaller animal pole blastomeres and the larger, yolky vegetal pole blastomeres is apparent. (c) Early blastula stage viewed from above and to one side. SEM photographs of Xenopus laevis embryos courtesy of Dr. Robert E. Waterman.

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throughout embryonic development. At room temperature (20–24° C), first cleavage of the egg occurs within 2 or 3 hours after fertilization. The first cleavage furrow is evident first at the animal pole and later at the vegetal pole, and it divides the egg into two equal blastomeres (fig. 4.4). Subsequent cleavages partition the egg into an increasing number of smaller blastomeres.

External Features of Development Various stages of development can be made available at different times by distributing both the control (diploid) and experimental (parthenogenetically activated) eggs to various temperature-control cabinets set at different temperatures, if they are available.The eggs of the leopard frog can tolerate temperatures as low as 6° C and as high as 28° C. However, for optimum development, restrict the range to temperatures between 12° C and 26° C. 1. Repeated reference to figure 4.5 and table 4.1 will aid in identifying the stages of development.The stage numbers are those originally assigned by Waldo Shumway in 1940. Plan to make frequent observations during the course of the week to witness the continuous change in the form of the embryo.

TABLE 4.1 Embryonic Development of the Frog

Stage*

Description

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25

Unfertilized Gray crescent 2-cell 4-cell 8-cell 16-cell 32-cell Mid-cleavage Late cleavage Dorsal lip Midgastrula Late gastrula Neural plate Neural folds Rotation Neural tube Tail bud Muscular response Heart beat Gill circulation Mouth open Tail fin circulation Opercular fold Operculum closed on right Operculum complete

Age (at 18° C) 0 1 3.5 4.5 5.5 6.5 7.5 16 21 26 34 42 50 62 67 72 84 96 118 140 162 192 216 240 284

in

Hours (at 25° C) 0 0.5 2.5 3.5 4.5 5.5 6.5 11 14 17 20 32 40 48 52 56 66 76 96 120 138 156 180 210 240

*Stages are numbered after Shumway (1940).

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1. Unfertilized

5. 8-cell

9. Late cleavage

13. Neural plate

2. Gray crescent

6. 16-cell

10. Dorsal lip

13. Neural folds

3. 2-cell

7. 32-cell

11. Midgastrula

15. Rotation

4. 4-cell

8. Mid-cleavage

12. Late gastrula

16. Neural tube

17. Tail bud

21. Mouth open

18. Muscular reponse

22. Tail fin circulation

19. Heart beat

23. Opercular fold

20. Gill circulation

24. Operculum closed on right

25. Operculum complete

FIGURE 4.5 Normal stages in the development of the frog embryo (Rana pipiens), based on Waldo Shumway’s series.

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The process of development takes place in a series of stages that occur in a regular sequence. Cleavage of the fertilized egg leads to the blastula stage (stages 8–9), which is followed by the process of gastrulation (stages 10–12). Gastrulation is a complex set of cell movements that results in the establishment and positioning of the three germ layers (ectoderm, endoderm, and mesoderm). Subsequently, the gastrula elongates, and there is development of neural folds and the neural tube (the neurula stages of development, stages 13–16). Various outpocketings, inpocketings, thickenings, and other changes produce the sundry organs of the body. The developmental history of internal structures will be considered in the next laboratory; only the externally visible changes during development are considered here. 2. Cleavage of the frog egg is total (holoblastic); cytokinesis divides the entire egg, notwithstanding the relatively large amount of yolk. Notice that the first cleavage is meridional (vertical); the second is also meridional, but at right angles to the first.The third is latitudinal, separating four smaller, upper animal hemisphere cells from four larger vegetal hemisphere cells. After the first few divisions, cleavage of the cells in the vegetal hemisphere lags behind that of the animal hemisphere cells that contain relatively little yolk and continue to divide quite synchronously. Throughout early development, yolky vegetal cells are larger than the animal pole cells. 3. Notice the externally visible changes that occur during gastrulation.An essential feature is epiboly, a spreading overgrowth by animal hemisphere cells that eventually envelop and enclose the vegetal hemisphere cells. Gastrulation movements are first indicated by the appearance of a crescent-shaped groove, or depression (stage 10), on one side of the blastula below the equator.The depression itself is known as the blastopore, and the rim above the depression is referred to as the dorsal lip of the blastopore (fig. 4.6). The tips of the blastopore progressively extend around until they meet—passing through a succession of shapes: quarter circle (stage 10), half circle (stage 11), and full circle (stage 12). At stage 12, only a small area of vegetal pole cells is visible. This remaining visible yolk area is called the yolk plug. The ring-shaped blastopore will soon close, that is, narrow to a barely visible slit (stage 13). 4. The body elongates during neurulation, and the neural folds are conspicuously elevated at stage 14 (fig. 4.6).At the tail-bud stage (stage 17 ), the embryo can easily be removed from its surrounding jelly coats and vitelline membrane. Use two pairs of fine forceps to grasp the vitelline membrane and rip it apart without harming or distorting the embryo. Leave some of the embryos in their membranes so that you can determine when normal hatching occurs. Become familiar with some of the important landmarks of the embryo at the tail-bud stage (fig. 4.7). At the anterior end is the prominent oral sucker (cement gland), a V-shaped groove with prominent lips. Between the lips of the sucker may be seen a depression called the stomodaeum, or mouth invagination.An olfactory pit and the optic bulge are evident at each side of the head.The gill plate has become subdivided by transverse furrows into three bars: the first, second, and third branchial (visceral ) arches. Behind the gill plate, a lateral swelling marks the position of the pronephros, or early larval kidney.The tail bud appears as an outgrowth of the posterior end of the body. Note that the embryo rotates continuously within its jelly coat, propelled by the cilia that cover its body. 5. Try to detect the beating of the heart at stage 19 by using a bright light source and focusing carefully on the ventral surface immediately posterior to the V-shaped sucker. Look for circulation of blood through the external gills developed as branched filaments on the branchial arches of a stage 20 embryo. In late embryonic development, the tail is differentiated into a dorsal and ventral fin, and myotomes (muscle segments) become clearly visible. On the ventral side, at the base of the tail, the proctodaeum is evident in the area where the blastopore closed. In the final stages of embryonic development (stages 21–25), a mouth forms, and rows of horny teeth develop. Observe the development of an operculum on each side as the external gills are resorbed and replaced by internal gills.These membranous operculum folds grow back from the visceral arches and become fused with the trunk, leaving only one aperture, the spiracle, on the left side. The spiracle provides the means by which water taken through the mouth passes through the gill region to the exterior. The completion of operculum development on both sides (stage 25) marks the end of embryonic development.

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DL YP

Early gastrula

Late gastrula

Neural folds

FIGURE 4.6 Selected stages during gastrulation and neurulation of the frog embryo (DL: dorsal lip of blastopore, and YP: yolk plug). Photographs from R. G. Kessel and C. Y. Shin, Scanning Electron Microscopy in Biology, 1976, Springer-Verlag.

gill plate pronephros optic bulge tail bud olfactory pit

stomodaeum oral sucker

FIGURE 4.7 A tail-bud stage (stage 17) frog embryo.

(a)

(b)

FIGURE 4.8 Mitotic figures as seen in aceto-orcein squash preparations of tail-region cells of (a) haploid (n ⫽ 13) and (b) diploid (2n ⫽ 26) embryos.

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Haploid Development Compare the haploid (parthenogenetically activated) embryos with diploid (control) embryos and describe any differences you observe in detail. You may wish to confirm the chromosome number in the haploid embryos. Accurate counts of metaphase chromosomes can be obtained from squash preparations of tail tips of embryos at stages 22–23. Representative haploid (n ⫽ 13) and diploid (2n ⫽ 26) metaphase configurations are shown in figure 4.8. 1. Immerse an embryo at stage 22 or 23 in a 1⬊3000 solution of ethyl m-aminobenzoate methane sulfonate until it becomes immobile. Cut off the distal third of the embryo’s tail tip with a razor blade or scalpel. Transfer the tail tip to a clean slide with a wide-mouth pipette. 2. Remove the solution around the tail tip and add several drops of distilled water. Leave the tail tip in distilled water for 2 to 10 minutes. Distilled water causes osmotic swelling of the nucleus that helps to untangle the chromosomes from each other. 3. Draw off the distilled water and replace it with a large drop of a 2% aceto-orcein solution. Quickly add a clean coverslip over the tissue to prevent crystal formation in the stain drop. 4. After 5 minutes, place a paper towel above the coverslip and exert strong pressure on the coverslip with your thumb. Seal the edges of the coverslip with melted glycerine jelly applied with a fine brush. (A comparable semipermanent preparation can also be prepared by using a nonresinous mounting medium to ring the edges of the coverslip.) Examine the preparation microscopically for mitotic figures, especially cells in metaphase of mitosis. 5. The slides may be stored in a refrigerator at 2–4° C.The preparation will last for several days, and indeed, the quality of staining might actually improve with time.

Materials EQUIPMENT Wooden-handled probe (dissecting needle) for pithing Scissors and forceps for dissection Clean microscope slides and coverslips “Frog pipettes” Disposable Pasteur pipettes, other clean pipettes, or medicine droppers Petri dishes Clean scalpel or section lifter Watchmaker’s forceps 4-inch finger bowls or other containers for developing eggs UV light source (see Section B) Compound microscope Dissecting microscope Illuminator SOLUTIONS

AND

CHEMICALS

Spring water or 10% Amphibian Ringer’s solution (see Appendix B) Ethyl m-aminobenzoate methane sulfonate solution (1⬊3000 in spring water or 10% Amphibian Ringer’s solution) Distilled water 2% aceto-orcein solution Petroleum jelly ( Vaseline), nonresinous mounting medium, nail hardener, or New Skin Liquid Bandage. LIVING MATERIAL Pituitary-injected female Rana pipiens and male Rana pipiens ( hormone-injected Xenopus laevis adults may be used instead) PRESERVED MATERIAL OPTION Some biological supply companies provide sets of preserved Rana pipiens embryos at a number of stages of development. Embryos at each stage are supplied in a small separate container that can be conveniently opened for direct observation.

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Some Useful Information Sources WEBSITES http://sdb.bio.purdue.edu/ This is the very useful website of the Society for Developmental Biology. It includes several sources of information about amphibian development, which can be found by following links to “Virtual LibraryDevelopmental Biology” and “Education.” http://worms.zoology.wisc.edu/frogs/welcome.html This website, prepared by Jeff Hardin, University of Wisconsin, is a good source of information on several aspects of amphibian development. www.ucalgary.ca/UofC/eduweb/virtualembryo/ This is a comprehensive website prepared by Leon Browder, University of Calgary, that has several links to sources of information on amphibian development. www.luc.edu/depts/biology/dev.htm This is Bill Wasserman’s Developmental Biology Page from Loyola University of Chicago. It contains a listing of Web resources, several of which provide information about amphibian development. www.utexas.edu/courses/zoo321/ This is a website prepared by Klaus Kalthoff, University of Texas, for his developmental biology course. Information about amphibian development can be found by following the links to “Movies” and “Related Web Sites.” VIDEO —A DOZEN EGGS This video includes a video sequence of frog gastrulation photographed by Ray Keller and John Shih, University of California, Berkeley.The video was produced under the auspices of the Society for Developmental Biology and is available from Sinauer Associates, Inc., P. O. Box 407, 23 Plumtree Road, Sunderland, MA 01375-0407.

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II. Amphibian Development

4. Observations and Experiments on the Living Frog Embryo


L A B O R A T O R Y

5 Patterns of Frog Development

The embryonic development of amphibians, like that of other animal embryos, begins with a series of mitotic cell divisions called cleavage. During cleavage, chromosomal replication, mitosis, and cytokinesis occur repeatedly without any intervening cell growth. This pattern of repeated division, but no growth, establishes a large population of cells, each of which contains a small part of the cytoplasm of the original, very large zygote.At the end of the cleavage phase of development, these many cells are organized as the blastula, a sphere of cells enclosing an internal fluid-filled cavity. Subsequent development involves the extensive cell migrations of gastrulation that reposition the cells of the embryo so that they are prepared to proceed with further development and differentiation. Gastrulation establishes the three basic body layers (germ layers): ectoderm, mesoderm, and endoderm. In some developing animals (for example, sea urchins) that produce eggs containing very little yolk, the cells of the blastula are arranged in the form of a relatively thin-walled, hollow ball.The process of gastrulation consists of the pushing inward of the cells of the vegetal hemisphere (prospective endoderm and mesoderm) in a manner comparable to pushing in one side of a hollow rubber ball. In amphibians, however, the presence of a large mass of yolk-filled cells precludes such a simple inpushing process.The eventual interior positioning of the endoderm and mesoderm of the amphibian embryo is effected by more complex processes. Gastrulation establishes the three germ layers in their final positions relative to one another. Each of the three germ layers is defined by its developmental fate.The ectoderm contains cells that will produce the epidermal covering of the body as well as the nervous system and sense organs; the endoderm produces the lining of the digestive tract and its various derivatives (for example, the lungs, digestive glands, and bladder); and the mesoderm, which lies between the other two, gives rise to a variety of tissues (for example, muscular, skeletal, and circulatory systems).

45


(a)




Animal pole

(b) pigment vitelline membrane

blastocoel

blastocoel

yolk-laden blastomeres

dorsal lip of blastopore

Vegetal pole (c) ectoderm (presumptive neural plate) roof of archenteron (chordamesoderm) archenteron floor of archenteron (endoderm) dorsal lip of blastopore

Animal pole

yolk plug ventral lip of blastopore blastocoel

FIGURE 5.1 Diagrams of sections of the frog blastula (a), early gastrula (b), and late gastrula (c).

Animal pole blastocoel

Vegetal pole blastocoel



archenteron blastocoel

archenteron yolk plug blastocoel

dorsal lip of blastopore lateral lip of blastopore

dorsal lip of blastopore yolk plug ventral lip of blastopore

FIGURE 5.2 Morphogenetic movements during gastrulation in the frog embryo; diagrams of longitudinal sections (left ) and oblique sections showing surface features (right ).

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Observations The external changes in the developing embryo’s form that you observed in the previous laboratory reveal only a small fraction of the events occurring internally.The internal processes can be studied in detail when the embryo is prepared as follows.The embryo is embedded in paraffin and/or various organic polymers, and cut with a microtome into a continuous series of slices or sections. The sections are stained and mounted in sequential order for microscopic study. This laboratory will be devoted to examining prepared slides of various stages of development of the frog embryo. Familiarity with the internal changes is essential to fully comprehending the developmental processes and for understanding any experimental studies that you may conduct later. Blastula As cleavage occurs in the frog, the embryo comes to consist of a large number of cells, or blastomeres, which enclose a hemispherical cavity that lies wholly in the animal hemisphere.This cavity is the blastocoel, and the embryo at this stage is called a blastula. The blastocoel does not arise abruptly; in fact, its beginnings may be detected as small spaces among the blastomeres as early as the eight-cell stage. Examine a microscopic slide of a section of the frog blastula. (Fig. 5.1a is drawn from stained sections of preserved embryos.) The jelly layers have been removed, but the vitelline membrane may be discernible as a faint line surrounding the embryo.The nuclei of the cells appear as dark spots.The cells themselves are sharply defined in the animal hemisphere but are only faintly outlined in the vegetal hemisphere. Note the eccentric location of the blastocoel.The roof of the blastocoel is formed by about four layers of small animal cells.The outermost cell layer is deeply pigmented. Below the blastocoel are a number of layers of larger cells, heavily laden with granules of yolk. (Note: the blastocoel fluid may have been coagulated into a compact mass by the chemicals used in the preparation of the slide.) Gastrulation Study figure 5.2 carefully (also consult fig. 5.1).The illustrations on the left in figure 5.2 represent the developing embryo cut in the median plane; those on the right are stereodiagrams of the same embryos. The process of gastrulation in the frog embryo is heralded by the appearance of a deeply pigmented, pitlike depression. The depression itself is known as the blastopore, and its rim is referred to as the dorsal lip of the blastopore. The animal hemisphere cells that have been actively migrating, and have actually extended downward below the equator, roll around the lip of the blastopore into the interior. Laterally, to the right and left of the dorsal lip, animal hemisphere cells roll in, or tuck in, to establish the lateral lips of the blastopore, causing the blastopore to become crescent shaped.As animal hemisphere cells that have spread downward become folded under at the ventral lip of the blastopore, the blastopore becomes circular and is filled with a mass of yolky vegetal cells known as the yolk plug. The process of gastrulation involves several types of cell movements. The downward movement of animal hemisphere cells to envelop the yolk-laden cells of the vegetal hemisphere is known as epiboly. The inward movement of cells at the blastopore lips is referred to as involution. There is a considerable streaming of surface cells into the interior of the embryo, and simultaneous internal reorganization of the embryo involves substantial shifting of yolky vegetal cells. The cell movements during gastrulation were first investigated using vital stains that color, but do not damage, the embryonic cells. In a series of experiments in the 1920s,Vogt briefly placed small agar chips saturated with Nile Blue sulphate against various areas of amphibian embryos at the beginning of gastrulation. He then traced the movement of the stained cells and determined their eventual position

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neural plate ectoderm

epidermal ectoderm

notochord dorsal (somite) mesoderm

dorsal lip of blastopore lateral mesoderm endoderm


epidermal ectoderm



lateral mesoderm

endoderm FIGURE 5.3 A fate map of an early gastrula stage frog embryo, based on Vogt’s experiments with vital stains.

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in the embryo at the end of gastrulation.This permitted construction of fate maps of the early gastrula, showing the eventual roles of cells in various regions (fig. 5.3). More recent research using injected intracellular dyes and other kinds of cell markers has confirmed the essential accuracy of these early fate maps.There are, however, variations among amphibian species in the details of fate maps. For example, fate maps of the clawed frog (Xenopus laevis) show that the fates of cells in some areas of the early gastrula depend upon whether these cells are at the surface or lie in deeper layers. It is also important to note that complex interactions among the cells of the early embryo are involved in determining the many different parts of the embryo. Your textbook will provide detailed information on what is now known about these various cell interactions. 1. Study a prepared section of an early gastrula in which the blastopore appears as a small dark notch and the blastocoel is conspicuous (fig. 5.1b). The blastopore is situated about 25° below the equator, in the boundary between the pigmented and unpigmented regions of the embryo. Some involution of cells is evident at the dorsal lip of the blastopore. Consult your textbook regarding the roles of cell shape change and cell movement in initiating and continuing inward migration of cells through the blastopore, beginning at the dorsal lip. 2. Study a longitudinal section through a late gastrula in which the blastocoel is still prominent, but the archenteron has formed (fig. 5.1c). Gastrulation movements have led to the formation of the slitlike archenteron, or primitive gut, which is lined by both future mesoderm and future endoderm cells. Cells that have moved to the interior through the dorsal lip and occupy the roof of the archenteron are future mesoderm cells. Cells along the midline of the archenteron roof are identified as the chordamesoderm and are destined to produce the notochord, while those to the right and left will produce the somites. Yolky endodermal cells form the floor and sidewalls of the archenteron. Later, the cells of the sides will spread upward to fuse beneath the chordamesoderm, thereby producing a complete, endodermally lined cavity, a true gut. At the same time, lateral mesoderm spreading and the differentiation of various portions of the mesoderm proceeds. The blastocoel becomes reduced to a small space on the ventral side and will eventually be obliterated. Identify the prominent yolk plug, bounded by the blastopore lips.

Study figure 5.4, which summarizes the movements of cells during gastrulation and identifies developmental fates of cells in various regions. It should be noted that considerable shifting of the yolk mass occurs, with a concomitant displacement of the center of gravity.The original animal pole of the egg becomes the anterior side of the future embryo, while the blastopore marks the posterior end of the future embryo. With the exception of the yolk plug, the outer surface of the egg is now covered with ectoderm. Two ectodermal regions are distinguishable: the epidermal ectoderm and the neural plate ectoderm. The future notochord and dorsal (somite) mesoderm are derived from the cells that involute at the dorsal and lateral lips of the blastopore. Future lateral mesoderm involutes at the lateral and ventral lips of the blastopore. Generalized Organogenesis in Vertebrates Once cells have migrated to their definitive locations in the gastrula, differentiation of the major organ systems proceeds. Before examining slides that show some aspects of organogenesis in the frog, it would be useful to familiarize yourself with a simplified and diagrammatic account of the development of the major parts of a generalized vertebrate embryo, depicted in figure 5.5. Note that the neural tube has its origin from ectodermal cells along the dorsal surface of the embryo. The mesodermal bands on either side of the neural tube and notochord, called the dorsal mesoderm, become divided transversely into components known as somites. The somites ultimately develop into the segmental muscles of the body. The mesodermal sheets adjacent to the developing gut constitute the lateral mesoderm. Each lateral mesodermal sheet splits into an outer somatic layer, adjacent to the ectoderm, and an inner splanchnic layer, next to the endoderm.The new cavity thus formed, which lies wholly within the mesoderm, is the coelom, or body cavity. The intermediate mesoderm (nephrotome) that lies between the developing somites and the spreading lateral sheets produces excretory and reproductive structures.The positions of the developing kidney and the genital ridge that initiates gonad development are indicated in figure 5.5.

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An.

D

V

D

V

Veg.

D

D

posterior anterior

V

V

Epidermal ectoderm

An. Animal pole

Neural plate ectoderm

Veg. Vegetal pole

Endoderm

D

Dorsal

Notochord and mesoderm

V

Ventral

FIGURE 5.4 Diagrams of several stages of gastrulation in the frog embryo, including the developmental fates of cells in various regions.

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neural groove

neural plate

neural tube

notochord

notochord mesoderm mesoderm

endoderm

endoderm archenteron

ectoderm

ectoderm

spinal cord

notochord vertebra

dorsal mesoderm

ectoderm

spinal cord

somite

notochord

somatic layer lateral mesoderm

coelom primitive kidney

splanchnic layer genital ridge coelom mesenchyme

dorsal mesentery digestive tract

FIGURE 5.5 A series of generalized diagrams showing major features of developing body organization in vertebrate embryos.

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(a)

(b) neural fold

dorsal mesoderm

neural plate

endoderm

notochord

dorsal mesoderm endoderm

gut lateral mesoderm

lateral mesoderm

epidermal ectoderm

epidermal ectoderm

(d)

(c) neural crest neural tube dorsal (somite) mesoderm

lateral mesoderm epidermal ectoderm

FIGURE 5.6 Neural tube formation in the frog embryo.

notochord

rhombencephalon

spinal cord somite

mesencephalon infundibulum

prosencephalon

anal evagination

hypophysis stomodaeum

yolk heart

midgut

oral evagination pharynx (foregut)

FIGURE 5.7 Median longitudinal section of a 3-mm frog embryo.

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liver diverticulum





The lateral mesodermal plates of the two sides grow toward the midline, both dorsal and ventral to the gut, until they come into contact with one another to form the two-layered dorsal and ventral mesenteries. Although the dorsal mesentery persists in the adult animal, nearly all of the ventral mesentery disappears, so that the two previously separate coelomic cavities join to become continuous. Although not represented in the illustration, mesodermal tissue that lies beneath the pharyngeal floor becomes involved in the formation of the heart. This brief account should assist you in understanding the relationships of the structures seen in sections of embryos at later stages of frog development. Neurulation in the Frog You observed earlier in intact embryos that the forerunner of the neural tube is the neural plate (stage 13), which appeared as a faintly outlined thickening of a wide strip of ectoderm above the notochord. The right and left borders of the neural plate subsequently became elevated as neural folds (stage 14). Finally, the neural folds curve inward and converge along the midline to form the neural tube (stages 15 and 16). 1. Compare a section of the neural plate stage, in which the ectoderm on the dorsal side of the embryo is flattened (fig. 5.6a), with a section of the neural fold stage, in which the margins of the neural plates are elevated, but the folds have not yet fused (fig. 5.6b).A neural fold is actually composed of two layers—an outer, pigmented epidermal layer and an inner neural layer. The former layer is thin, while the neural layer is thicker and is composed of tall columnar cells. 2. Note that by this time there has been a separation of the dorsal mesoderm that leaves the notochord as a distinct median rod. The future coelom, the cavity within the lateral mesoderm, is not yet evident at this stage. Note also that the endodermal roof of the gut has developed by the upward spreading of the archenteron’s sides. 3. Study a section in which the neural folds are in the process of fusing (fig. 5.6c). Cells that proliferate from the lateral inner margin of each neural fold are neural crest cells, the progenitors of pigment cells and a variety of other cell types. Laboratory 6 includes an experiment on the origin and the differentiation of neural crest cells. 4. Observe a cross section of an embryo in which formation of the neural tube is completed (fig. 5.6d). When the folds have finally closed, the outer layer of epidermal (skin) ectoderm is separated from the roof of the neural tube. The mesodermal tissue between the ectoderm and endoderm has differentiated dorsally into somites on each side of the neural tube and into lateral mesoderm enclosing the archenteron. Note the appearance of a cavity within each mesodermal somite. This cavity will close in later development as the somite differentiates into segmental muscle and other structures. The lateral mesoderm is composed of two layers of cells that, when separated, will enclose the body cavity (coelom). The coelomic cavity is still not conspicuous at this stage.

Tail-Bud Embryo (Stage 17) The tail-bud stage of the frog exemplifies the main features of the basic body plan of vertebrate embryo.The anterior region of the neural tube expands to form the brain.Three primary enlargements— the forebrain ( prosencephalon), midbrain (mesencephalon), and hindbrain (rhombencephalon)— characterize the cephalic part of the neural tube in all vertebrate embryos. Later subdivisions result in a brain with five well-defined regions characteristic of all vertebrate animals. The rudiments of the paired eyes (optic vesicles) are lateral outgrowths of the forebrain, and the future lens of the eye is derived from the neighboring superficial ectoderm. The auditory (otic) vesicle, which will later develop into the internal ear, is formed from a thickened patch of superficial ectoderm opposite the hindbrain. Other conspicuous features, characteristic of all vertebrate embryos, are the stomodaeum and the proctodaeum.The former is a shallow ectodermal depression that meets an endodermal out-pocketing (the oral evagination) at the extreme anterior end of the alimentary canal. The resulting two-layered Laboratory 5

53





(a)

(b)

epiphysis epidermal ectoderm

olfactory placode

prosocoel

retinal layer of optic cup

lens placode

prosencephalon (neural ectoderm)

prosocoel

prosencephalon

epidermal ectoderm optic cup

optic stalk hypophysis oral evagination

mesenchyme

oral membrane oral sucker stomodaeum

(c) mesencephalon mesocoel

optic cup infundibulum hypophysis

pharynx (foregut) oral sucker

(d)

(e) spinal cord rhombocoel

rhombencephalon notochord auditory vesicle

notochord

pronephros gut somatic layer of lateral mesoderm

pharynx yolk

dorsal mesocardium pericardial cavity oral sucker

endocardium myocardium

liver diverticulum

parietal pericardium

FIGURE 5.8 Representative cross sections of the 3-mm frog embryo.

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somite

splanchnic layer of lateral mesoderm epidermal ectoderm





oral membrane will break through to form the definitive mouth.At the base of the tail, an ectodermal pit, the proctodaeum, will open into the hind end of the alimentary cavity to form the posterior aperture, the cloaca. As you study the slides of the 3- or 4-mm frog embryo, you will encounter rudiments of several other organs and systems. 1. Study a median longitudinal section of a 3- or 4-mm frog embryo (fig. 5.7 ). The three primary divisions of the brain ( prosencephalon, mesencephalon, and rhombencephalon) are evident.The prosencephalon is chiefly below, and in front of, the anterior end of the notochord; the mesencephalon is anterodorsal to the end of the notochord; and the rhombencephalon lies entirely over the notochord. An out-pushing in the ventral portion of the forebrain represents the infundibulum, the forerunner of the posterior lobe of the pituitary gland.The rudiment of the anterior lobe of the pituitary, the hypophysis, may be seen as a mass of ectodermal cells dorsal to the stomodaeal invagination. The -shaped markings are the somites, which formed from the dorsal mesoderm lateral to the notochord. Differentiated parts of the endodermal gut include the pharynx (foregut), liver diverticulum, midgut, and hindgut (not readily apparent).The mesodermal rudiments of the heart appear below the pharynx. 2. Study serial cross sections of the 3- or 4-mm frog embryo (fig. 5.8).The whole embryo has been cut transversely into a continuous series of slices, or sections, and mounted in an anteroposterior sequence on one slide. The most-anterior slices appear at the upper left-hand corner, and each row of sections is “read” from left to right. As you observe the structures in each section, try to build a mental reconstruction of the whole embryo from the various slices. a. The most-anterior sections pass through the forebrain, or prosencephalon. The conspicuous cavity of the forebrain is the prosocoel (fig. 5.8a). Notice that the wall of the forebrain, derived from neural ectoderm cells, is clearly delimited from the outer covering of heavily pigmented epidermal (skin) ectoderm. Ventrolaterally, on either side of the prosocoel, the epidermal ectoderm has thickened and invaginated to form the olfactory placodes, or nasal pits. In later development, these pits will enlarge to form the nasal cavities and will shift away from the surface of the head. The nasal cavities will, however, remain connected to the surface by tubes whose outer openings form the external nares. In many specimens at this stage, you may observe a narrow dorsal out-pushing of the brain vesicle.This dorsal evagination is the epiphysis, the forerunner of the pineal adult body. b. As you trace the sections posteriorly, observe that the olfactory placodes disappear and a cleft appears in the midventral line.This ventral ectodermal invagination is the stomodaeum (fig. 5.8b).The stomodaeal wall meets and fuses with an endodermal out-pocketing of the pharynx, the oral evagination.The region of contact between the stomodaeal invagination and the oral invagination is the oral membrane, or oral plate (fig. 5.8b). When the oral membrane becomes perforated, the stomodaeal cavity (mouth) opens into the pharynx. Notice the prominent paired ectodermal elevations on each side of the stomodaeum, called the oral suckers, or mucous glands. Identify the optic cups, which developed from the optic vesicles, two lateral evaginations of the wall of the prosencephalon (more specifically, of the diencephalon region of the prosencephalon).The optic cup has two layers; the thick (most lateral) layer is sensory and will form the retina of the eye.This thick sensory layer also induces the adjacent epidermal ectoderm layer to form the lens placode. Beneath the floor of the prosencephalon and dorsal to the foregut, notice the thickened stalk of ectodermal cells, the hypophysis, which is one of the rudiments of the pituitary gland. The spaces between the superficial ectoderm and the wall of the brain are filled with scattered mesodermal cells known as mesenchyme. Mesenchyme cells will develop into connective tissue, muscle, bones, and blood vessels of the head. c. At the level shown in figure 5.8c, the brain appears to have divided into two separate parts. Actually, this section passes through the thick-roofed midbrain (mesencephalon), represented by the dorsal component, and through the ventrally located infundibulum, an out-pocketing of the diencephalon region of the prosencephalon. Consulting figure 5.7 again will help clarify this relationship. The infundibulum together with the hypophysis will give rise to the pituitary gland.The previously observed, rodlike aggregation of cells comprising the hypophysis is still evident just ventral to the infundibulum. d. Continue to trace the sections posteriorly, and notice that the cavity of the brain becomes reduced in size and the roof becomes thinner (fig. 5.8 d ). The thin-walled roof serves to identify the rhombencephalon (the future metencephalon and myelencephalon). Beneath the brain is the conspicuous notochord with its large vesicular cells. Lateral and slightly ventral to the brain are the auditory vesicles, which later differentiate into the membranous labyrinth of the inner ear. The auditory vesicles originated from paired, thickened lateral depressions in the superficial ectoderm.

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FIGURE 5.9 The exposed genital ridges of the frog tadpole, showing the lightcolored primordial germ cells (indicated by arrows). The illustration shows two tadpoles in parabiotic union; the technique of parabiosis is considered in Laboratory 6.

Notes and Observations

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Beneath the expanded pharynx, splanchnic mesodermal elements have formed the embryonic heart.The heart is contained in a large semicircular cavity, the pericardial cavity. The thicker layer of the developing heart is the myocardium; it represents the future muscular wall of the heart. In the cavity inside the myocardium, mesodermal cells are becoming organized into the continuous endothelial lining of the heart known as the endocardium. The outer thin layer of the pericardial cavity comprises the future parietal wall of the adult pericardial cavity. e. Examine serial sections more posteriorly until the gut is very reduced dorsally and the rudiment of the liver is visible as a fingerlike evagination of the ventral wall of the foregut.The liver diverticulum extends into the yolk mass (fig. 5.8 e). Consult figure 5.7 again in order to clarify the physical relationship between the anterior portion of the gut and the liver rudiment. On either side of the notochord, the dorsal mesoderm has segmented to form the somites. Notice the double-layered condition of the lateral mesoderm: the outer layer adjacent to the ectoderm is the somatic mesoderm, while the inner layer adjacent to the endoderm is the splanchnic layer. The cavity between these two layers is the coelom. Between the dorsal mesoderm (somite) and the lateral mesoderm is an area known as the intermediate mesoderm, or nephrotome (sometimes also called mesomeric mesoderm). At this stage, the intermediate mesoderm has enlarged and differentiated into the pronephros, the functional kidney of the larva.The pronephric duct, with its hollow cavity, may also be identifiable. Far posteriorly, locate the proctodaeum, which originates as a ventral invagination of the epidermal ectoderm.

Dissection of the 11-mm Embryo (Stage 25) During stages 23 to 25, a fold of integument, the opercular fold, grows back to enclose and cover the external gills on each side. This is completed sooner on the right side (stage 24) than on the left side (stage 25). The resulting gill chamber remains in communication with the exterior through a single, short funnel on the left side, known as the spiracle. The dissection described in the following for stage-25 embryos may be performed equally as well on stage 24 embryos. 1. Use a wide-mouth pipette to transfer a tadpole to the anesthetic solution. This is a 1⬊3000 solution of ethyl m-aminobenzoate methanesulfonate in either spring water or 10% Amphibian Ringer’s solution. Wait until the tadpole stops all swimming and other movements. Then test it for responsiveness to touch by gently poking it with a blunt instrument. When the tadpole shows no response to touch, transfer it to an operating dish.There should be at least enough of the anesthetic solution in the dish to prevent excessive drying. 2. Place the operating dish on the stage of a dissecting microscope and illuminate it from the side with a focusing illuminator. Turn the tadpole on its back in a shallow groove that you have prepared in your operating dish. Grasp the skin in the anal region (at the base of the tail) with two pairs of watchmaker’s forceps and tear open the ventral body wall. Observe the coiled intestine and attempt to trace its course.The tadpole feeds largely on plants and, like other herbivores, requires a large absorbing surface, provided by the long intestinal tract. Note that great length without great bulk is accomplished by the coiling of the tract into a compact spiral. Note also other abdominal structures such as the liver. Finish exposing the heart if it is not uncovered and examine it and other structures near it. 3. Return to the abdominal area and remove all digestive organs from the body cavity.You should now have exposed the dorsal aspect of the lining of the body cavity.Along the midline, you can observe the area of the developing gonads, the genital ridges (fig. 5.9). Switch your microscope to a higher magnification and carefully observe this area. The two ridges project into the coelomic cavity and are covered by coelomic epithelium (peritoneum). Note the large, light-colored protuberances on the ridges. These are primordial germ cells that have migrated into this area to colonize the gonads. 4. Make sketches of your observations on the internal anatomy of the tadpole.You may have to dissect several tadpoles in order to complete your observations on tadpole morphology.

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Materials EQUIPMENT Compound microscope Dissecting microscope Illuminator Watchmaker’s forceps Operating dish (see Appendix B) “Frog pipette” SOLUTIONS

AND

CHEMICALS

Ethyl m-aminobenzoate methanesulfonate solution (1⬊3000 in spring water or 10% Amphibian Ringer’s solution) LIVING MATERIAL Stage 25 (11-mm) frog embryos COMMERCIALLY PREPARED SLIDES Frog blastula section Early gastrula sagittal section Late gastrula or yolk plug sagittal section Neural plate or early neural groove representative cross sections (rep. c.s.) Neural fold or late neural groove rep. c.s. (some companies supply slides labeled “groove-tube transition”) Early neural tube rep. c.s. Late neural tube rep. c.s. Representative longitudinal sections of 3- or 4-mm frog embryos (selected sections on a “serial sagittal sections” slide can be used as well) Serial cross sections of 3- or 4-mm frog embryos

Some Useful Information Sources WEBSITES http://zygote.swarthmore.edu/index.html This comprehensive site, assembled by Scott Gilbert, Swarthmore College, includes a good deal of information about developmental processes in amphibians and has links to other relevant sites. www.uoguelph.ca/zoology/devobio/dbindex.htm This site contains Stephen Scadding’s supplemental materials for his developmental biology course at the University of Guelph, Ontario. It includes a set of labeled cross sections of frog embryos plus other information about frog development. http://sdb.bio.purdue.edu/SDBEduca/qt_embryos1.html This site, assembled by Laurie Iten, Purdue University, contains movies of serial sections and discussion of the value of serial section study.

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5. Patterns of Frog Development


L A B O R A T O R Y

5 Patterns of Frog Development

The embryonic development of amphibians, like that of other animal embryos, begins with a series of mitotic cell divisions called cleavage. During cleavage, chromosomal replication, mitosis, and cytokinesis occur repeatedly without any intervening cell growth. This pattern of repeated division, but no growth, establishes a large population of cells, each of which contains a small part of the cytoplasm of the original, very large zygote.At the end of the cleavage phase of development, these many cells are organized as the blastula, a sphere of cells enclosing an internal fluid-filled cavity. Subsequent development involves the extensive cell migrations of gastrulation that reposition the cells of the embryo so that they are prepared to proceed with further development and differentiation. Gastrulation establishes the three basic body layers (germ layers): ectoderm, mesoderm, and endoderm. In some developing animals (for example, sea urchins) that produce eggs containing very little yolk, the cells of the blastula are arranged in the form of a relatively thin-walled, hollow ball.The process of gastrulation consists of the pushing inward of the cells of the vegetal hemisphere (prospective endoderm and mesoderm) in a manner comparable to pushing in one side of a hollow rubber ball. In amphibians, however, the presence of a large mass of yolk-filled cells precludes such a simple inpushing process.The eventual interior positioning of the endoderm and mesoderm of the amphibian embryo is effected by more complex processes. Gastrulation establishes the three germ layers in their final positions relative to one another. Each of the three germ layers is defined by its developmental fate.The ectoderm contains cells that will produce the epidermal covering of the body as well as the nervous system and sense organs; the endoderm produces the lining of the digestive tract and its various derivatives (for example, the lungs, digestive glands, and bladder); and the mesoderm, which lies between the other two, gives rise to a variety of tissues (for example, muscular, skeletal, and circulatory systems).

45


(a)




Animal pole

(b) pigment vitelline membrane

blastocoel

blastocoel

yolk-laden blastomeres


Vegetal pole (c) ectoderm (presumptive neural plate) roof of archenteron (chordamesoderm) archenteron floor of archenteron (endoderm) dorsal lip of blastopore

Animal pole

yolk plug ventral lip of blastopore blastocoel

FIGURE 5.1 Diagrams of sections of the frog blastula (a), early gastrula (b), and late gastrula (c).

Animal pole blastocoel

Vegetal pole blastocoel



archenteron blastocoel

archenteron yolk plug blastocoel

dorsal lip of blastopore lateral lip of blastopore

dorsal lip of blastopore yolk plug ventral lip of blastopore

FIGURE 5.2 Morphogenetic movements during gastrulation in the frog embryo; diagrams of longitudinal sections (left ) and oblique sections showing surface features (right ).

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Observations The external changes in the developing embryo’s form that you observed in the previous laboratory reveal only a small fraction of the events occurring internally.The internal processes can be studied in detail when the embryo is prepared as follows.The embryo is embedded in paraffin and/or various organic polymers, and cut with a microtome into a continuous series of slices or sections. The sections are stained and mounted in sequential order for microscopic study. This laboratory will be devoted to examining prepared slides of various stages of development of the frog embryo. Familiarity with the internal changes is essential to fully comprehending the developmental processes and for understanding any experimental studies that you may conduct later. Blastula As cleavage occurs in the frog, the embryo comes to consist of a large number of cells, or blastomeres, which enclose a hemispherical cavity that lies wholly in the animal hemisphere.This cavity is the blastocoel, and the embryo at this stage is called a blastula. The blastocoel does not arise abruptly; in fact, its beginnings may be detected as small spaces among the blastomeres as early as the eight-cell stage. Examine a microscopic slide of a section of the frog blastula. (Fig. 5.1a is drawn from stained sections of preserved embryos.) The jelly layers have been removed, but the vitelline membrane may be discernible as a faint line surrounding the embryo.The nuclei of the cells appear as dark spots.The cells themselves are sharply defined in the animal hemisphere but are only faintly outlined in the vegetal hemisphere. Note the eccentric location of the blastocoel.The roof of the blastocoel is formed by about four layers of small animal cells.The outermost cell layer is deeply pigmented. Below the blastocoel are a number of layers of larger cells, heavily laden with granules of yolk. (Note: the blastocoel fluid may have been coagulated into a compact mass by the chemicals used in the preparation of the slide.) Gastrulation Study figure 5.2 carefully (also consult fig. 5.1).The illustrations on the left in figure 5.2 represent the developing embryo cut in the median plane; those on the right are stereodiagrams of the same embryos. The process of gastrulation in the frog embryo is heralded by the appearance of a deeply pigmented, pitlike depression. The depression itself is known as the blastopore, and its rim is referred to as the dorsal lip of the blastopore. The animal hemisphere cells that have been actively migrating, and have actually extended downward below the equator, roll around the lip of the blastopore into the interior. Laterally, to the right and left of the dorsal lip, animal hemisphere cells roll in, or tuck in, to establish the lateral lips of the blastopore, causing the blastopore to become crescent shaped.As animal hemisphere cells that have spread downward become folded under at the ventral lip of the blastopore, the blastopore becomes circular and is filled with a mass of yolky vegetal cells known as the yolk plug. The process of gastrulation involves several types of cell movements. The downward movement of animal hemisphere cells to envelop the yolk-laden cells of the vegetal hemisphere is known as epiboly. The inward movement of cells at the blastopore lips is referred to as involution. There is a considerable streaming of surface cells into the interior of the embryo, and simultaneous internal reorganization of the embryo involves substantial shifting of yolky vegetal cells. The cell movements during gastrulation were first investigated using vital stains that color, but do not damage, the embryonic cells. In a series of experiments in the 1920s,Vogt briefly placed small agar chips saturated with Nile Blue sulphate against various areas of amphibian embryos at the beginning of gastrulation. He then traced the movement of the stained cells and determined their eventual position

Laboratory 5

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epidermal ectoderm


dorsal lip of blastopore lateral mesoderm endoderm


epidermal ectoderm



lateral mesoderm

endoderm FIGURE 5.3 A fate map of an early gastrula stage frog embryo, based on Vogt’s experiments with vital stains.

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in the embryo at the end of gastrulation.This permitted construction of fate maps of the early gastrula, showing the eventual roles of cells in various regions (fig. 5.3). More recent research using injected intracellular dyes and other kinds of cell markers has confirmed the essential accuracy of these early fate maps.There are, however, variations among amphibian species in the details of fate maps. For example, fate maps of the clawed frog (Xenopus laevis) show that the fates of cells in some areas of the early gastrula depend upon whether these cells are at the surface or lie in deeper layers. It is also important to note that complex interactions among the cells of the early embryo are involved in determining the many different parts of the embryo. Your textbook will provide detailed information on what is now known about these various cell interactions. 1. Study a prepared section of an early gastrula in which the blastopore appears as a small dark notch and the blastocoel is conspicuous (fig. 5.1b). The blastopore is situated about 25° below the equator, in the boundary between the pigmented and unpigmented regions of the embryo. Some involution of cells is evident at the dorsal lip of the blastopore. Consult your textbook regarding the roles of cell shape change and cell movement in initiating and continuing inward migration of cells through the blastopore, beginning at the dorsal lip. 2. Study a longitudinal section through a late gastrula in which the blastocoel is still prominent, but the archenteron has formed (fig. 5.1c). Gastrulation movements have led to the formation of the slitlike archenteron, or primitive gut, which is lined by both future mesoderm and future endoderm cells. Cells that have moved to the interior through the dorsal lip and occupy the roof of the archenteron are future mesoderm cells. Cells along the midline of the archenteron roof are identified as the chordamesoderm and are destined to produce the notochord, while those to the right and left will produce the somites. Yolky endodermal cells form the floor and sidewalls of the archenteron. Later, the cells of the sides will spread upward to fuse beneath the chordamesoderm, thereby producing a complete, endodermally lined cavity, a true gut. At the same time, lateral mesoderm spreading and the differentiation of various portions of the mesoderm proceeds. The blastocoel becomes reduced to a small space on the ventral side and will eventually be obliterated. Identify the prominent yolk plug, bounded by the blastopore lips.

Study figure 5.4, which summarizes the movements of cells during gastrulation and identifies developmental fates of cells in various regions. It should be noted that considerable shifting of the yolk mass occurs, with a concomitant displacement of the center of gravity.The original animal pole of the egg becomes the anterior side of the future embryo, while the blastopore marks the posterior end of the future embryo. With the exception of the yolk plug, the outer surface of the egg is now covered with ectoderm. Two ectodermal regions are distinguishable: the epidermal ectoderm and the neural plate ectoderm. The future notochord and dorsal (somite) mesoderm are derived from the cells that involute at the dorsal and lateral lips of the blastopore. Future lateral mesoderm involutes at the lateral and ventral lips of the blastopore. Generalized Organogenesis in Vertebrates Once cells have migrated to their definitive locations in the gastrula, differentiation of the major organ systems proceeds. Before examining slides that show some aspects of organogenesis in the frog, it would be useful to familiarize yourself with a simplified and diagrammatic account of the development of the major parts of a generalized vertebrate embryo, depicted in figure 5.5. Note that the neural tube has its origin from ectodermal cells along the dorsal surface of the embryo. The mesodermal bands on either side of the neural tube and notochord, called the dorsal mesoderm, become divided transversely into components known as somites. The somites ultimately develop into the segmental muscles of the body. The mesodermal sheets adjacent to the developing gut constitute the lateral mesoderm. Each lateral mesodermal sheet splits into an outer somatic layer, adjacent to the ectoderm, and an inner splanchnic layer, next to the endoderm.The new cavity thus formed, which lies wholly within the mesoderm, is the coelom, or body cavity. The intermediate mesoderm (nephrotome) that lies between the developing somites and the spreading lateral sheets produces excretory and reproductive structures.The positions of the developing kidney and the genital ridge that initiates gonad development are indicated in figure 5.5.

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An.

D

V

D

V

Veg.

D

D

posterior anterior

V

V

Epidermal ectoderm

An. Animal pole

Neural plate ectoderm

Veg. Vegetal pole

Endoderm

D

Dorsal

Notochord and mesoderm

V

Ventral

FIGURE 5.4 Diagrams of several stages of gastrulation in the frog embryo, including the developmental fates of cells in various regions.

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neural groove

neural plate

neural tube

notochord

notochord mesoderm mesoderm

endoderm

endoderm archenteron

ectoderm

ectoderm

spinal cord

notochord vertebra

dorsal mesoderm

ectoderm

spinal cord

somite

notochord

somatic layer lateral mesoderm

coelom primitive kidney

splanchnic layer genital ridge coelom mesenchyme

dorsal mesentery digestive tract

FIGURE 5.5 A series of generalized diagrams showing major features of developing body organization in vertebrate embryos.

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(a)

(b) neural fold

dorsal mesoderm

neural plate

endoderm

notochord

dorsal mesoderm endoderm

gut lateral mesoderm

lateral mesoderm

epidermal ectoderm

epidermal ectoderm

(d)

(c) neural crest neural tube dorsal (somite) mesoderm

lateral mesoderm epidermal ectoderm

FIGURE 5.6 Neural tube formation in the frog embryo.

notochord

rhombencephalon

spinal cord somite

mesencephalon infundibulum

prosencephalon

anal evagination

hypophysis stomodaeum

yolk heart

midgut

oral evagination pharynx (foregut)

FIGURE 5.7 Median longitudinal section of a 3-mm frog embryo.

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liver diverticulum





The lateral mesodermal plates of the two sides grow toward the midline, both dorsal and ventral to the gut, until they come into contact with one another to form the two-layered dorsal and ventral mesenteries. Although the dorsal mesentery persists in the adult animal, nearly all of the ventral mesentery disappears, so that the two previously separate coelomic cavities join to become continuous. Although not represented in the illustration, mesodermal tissue that lies beneath the pharyngeal floor becomes involved in the formation of the heart. This brief account should assist you in understanding the relationships of the structures seen in sections of embryos at later stages of frog development. Neurulation in the Frog You observed earlier in intact embryos that the forerunner of the neural tube is the neural plate (stage 13), which appeared as a faintly outlined thickening of a wide strip of ectoderm above the notochord. The right and left borders of the neural plate subsequently became elevated as neural folds (stage 14). Finally, the neural folds curve inward and converge along the midline to form the neural tube (stages 15 and 16). 1. Compare a section of the neural plate stage, in which the ectoderm on the dorsal side of the embryo is flattened (fig. 5.6a), with a section of the neural fold stage, in which the margins of the neural plates are elevated, but the folds have not yet fused (fig. 5.6b).A neural fold is actually composed of two layers—an outer, pigmented epidermal layer and an inner neural layer. The former layer is thin, while the neural layer is thicker and is composed of tall columnar cells. 2. Note that by this time there has been a separation of the dorsal mesoderm that leaves the notochord as a distinct median rod. The future coelom, the cavity within the lateral mesoderm, is not yet evident at this stage. Note also that the endodermal roof of the gut has developed by the upward spreading of the archenteron’s sides. 3. Study a section in which the neural folds are in the process of fusing (fig. 5.6c). Cells that proliferate from the lateral inner margin of each neural fold are neural crest cells, the progenitors of pigment cells and a variety of other cell types. Laboratory 6 includes an experiment on the origin and the differentiation of neural crest cells. 4. Observe a cross section of an embryo in which formation of the neural tube is completed (fig. 5.6d). When the folds have finally closed, the outer layer of epidermal (skin) ectoderm is separated from the roof of the neural tube. The mesodermal tissue between the ectoderm and endoderm has differentiated dorsally into somites on each side of the neural tube and into lateral mesoderm enclosing the archenteron. Note the appearance of a cavity within each mesodermal somite. This cavity will close in later development as the somite differentiates into segmental muscle and other structures. The lateral mesoderm is composed of two layers of cells that, when separated, will enclose the body cavity (coelom). The coelomic cavity is still not conspicuous at this stage.

Tail-Bud Embryo (Stage 17) The tail-bud stage of the frog exemplifies the main features of the basic body plan of vertebrate embryo.The anterior region of the neural tube expands to form the brain.Three primary enlargements— the forebrain ( prosencephalon), midbrain (mesencephalon), and hindbrain (rhombencephalon)— characterize the cephalic part of the neural tube in all vertebrate embryos. Later subdivisions result in a brain with five well-defined regions characteristic of all vertebrate animals. The rudiments of the paired eyes (optic vesicles) are lateral outgrowths of the forebrain, and the future lens of the eye is derived from the neighboring superficial ectoderm. The auditory (otic) vesicle, which will later develop into the internal ear, is formed from a thickened patch of superficial ectoderm opposite the hindbrain. Other conspicuous features, characteristic of all vertebrate embryos, are the stomodaeum and the proctodaeum.The former is a shallow ectodermal depression that meets an endodermal out-pocketing (the oral evagination) at the extreme anterior end of the alimentary canal. The resulting two-layered Laboratory 5

53





(a)

(b)

epiphysis epidermal ectoderm

olfactory placode

prosocoel

retinal layer of optic cup

lens placode

prosencephalon (neural ectoderm)

prosocoel

prosencephalon

epidermal ectoderm optic cup

optic stalk hypophysis oral evagination

mesenchyme

oral membrane oral sucker stomodaeum

(c) mesencephalon mesocoel

optic cup infundibulum hypophysis

pharynx (foregut) oral sucker

(d)

(e) spinal cord rhombocoel

rhombencephalon notochord auditory vesicle

notochord

pronephros gut somatic layer of lateral mesoderm

pharynx yolk

dorsal mesocardium pericardial cavity oral sucker

endocardium myocardium

liver diverticulum

parietal pericardium

FIGURE 5.8 Representative cross sections of the 3-mm frog embryo.

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somite

splanchnic layer of lateral mesoderm epidermal ectoderm





oral membrane will break through to form the definitive mouth.At the base of the tail, an ectodermal pit, the proctodaeum, will open into the hind end of the alimentary cavity to form the posterior aperture, the cloaca. As you study the slides of the 3- or 4-mm frog embryo, you will encounter rudiments of several other organs and systems. 1. Study a median longitudinal section of a 3- or 4-mm frog embryo (fig. 5.7 ). The three primary divisions of the brain ( prosencephalon, mesencephalon, and rhombencephalon) are evident.The prosencephalon is chiefly below, and in front of, the anterior end of the notochord; the mesencephalon is anterodorsal to the end of the notochord; and the rhombencephalon lies entirely over the notochord. An out-pushing in the ventral portion of the forebrain represents the infundibulum, the forerunner of the posterior lobe of the pituitary gland.The rudiment of the anterior lobe of the pituitary, the hypophysis, may be seen as a mass of ectodermal cells dorsal to the stomodaeal invagination. The -shaped markings are the somites, which formed from the dorsal mesoderm lateral to the notochord. Differentiated parts of the endodermal gut include the pharynx (foregut), liver diverticulum, midgut, and hindgut (not readily apparent).The mesodermal rudiments of the heart appear below the pharynx. 2. Study serial cross sections of the 3- or 4-mm frog embryo (fig. 5.8).The whole embryo has been cut transversely into a continuous series of slices, or sections, and mounted in an anteroposterior sequence on one slide. The most-anterior slices appear at the upper left-hand corner, and each row of sections is “read” from left to right. As you observe the structures in each section, try to build a mental reconstruction of the whole embryo from the various slices. a. The most-anterior sections pass through the forebrain, or prosencephalon. The conspicuous cavity of the forebrain is the prosocoel (fig. 5.8a). Notice that the wall of the forebrain, derived from neural ectoderm cells, is clearly delimited from the outer covering of heavily pigmented epidermal (skin) ectoderm. Ventrolaterally, on either side of the prosocoel, the epidermal ectoderm has thickened and invaginated to form the olfactory placodes, or nasal pits. In later development, these pits will enlarge to form the nasal cavities and will shift away from the surface of the head. The nasal cavities will, however, remain connected to the surface by tubes whose outer openings form the external nares. In many specimens at this stage, you may observe a narrow dorsal out-pushing of the brain vesicle.This dorsal evagination is the epiphysis, the forerunner of the pineal adult body. b. As you trace the sections posteriorly, observe that the olfactory placodes disappear and a cleft appears in the midventral line.This ventral ectodermal invagination is the stomodaeum (fig. 5.8b).The stomodaeal wall meets and fuses with an endodermal out-pocketing of the pharynx, the oral evagination.The region of contact between the stomodaeal invagination and the oral invagination is the oral membrane, or oral plate (fig. 5.8b). When the oral membrane becomes perforated, the stomodaeal cavity (mouth) opens into the pharynx. Notice the prominent paired ectodermal elevations on each side of the stomodaeum, called the oral suckers, or mucous glands. Identify the optic cups, which developed from the optic vesicles, two lateral evaginations of the wall of the prosencephalon (more specifically, of the diencephalon region of the prosencephalon).The optic cup has two layers; the thick (most lateral) layer is sensory and will form the retina of the eye.This thick sensory layer also induces the adjacent epidermal ectoderm layer to form the lens placode. Beneath the floor of the prosencephalon and dorsal to the foregut, notice the thickened stalk of ectodermal cells, the hypophysis, which is one of the rudiments of the pituitary gland. The spaces between the superficial ectoderm and the wall of the brain are filled with scattered mesodermal cells known as mesenchyme. Mesenchyme cells will develop into connective tissue, muscle, bones, and blood vessels of the head. c. At the level shown in figure 5.8c, the brain appears to have divided into two separate parts. Actually, this section passes through the thick-roofed midbrain (mesencephalon), represented by the dorsal component, and through the ventrally located infundibulum, an out-pocketing of the diencephalon region of the prosencephalon. Consulting figure 5.7 again will help clarify this relationship. The infundibulum together with the hypophysis will give rise to the pituitary gland.The previously observed, rodlike aggregation of cells comprising the hypophysis is still evident just ventral to the infundibulum. d. Continue to trace the sections posteriorly, and notice that the cavity of the brain becomes reduced in size and the roof becomes thinner (fig. 5.8 d ). The thin-walled roof serves to identify the rhombencephalon (the future metencephalon and myelencephalon). Beneath the brain is the conspicuous notochord with its large vesicular cells. Lateral and slightly ventral to the brain are the auditory vesicles, which later differentiate into the membranous labyrinth of the inner ear. The auditory vesicles originated from paired, thickened lateral depressions in the superficial ectoderm.

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FIGURE 5.9 The exposed genital ridges of the frog tadpole, showing the lightcolored primordial germ cells (indicated by arrows). The illustration shows two tadpoles in parabiotic union; the technique of parabiosis is considered in Laboratory 6.


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Beneath the expanded pharynx, splanchnic mesodermal elements have formed the embryonic heart.The heart is contained in a large semicircular cavity, the pericardial cavity. The thicker layer of the developing heart is the myocardium; it represents the future muscular wall of the heart. In the cavity inside the myocardium, mesodermal cells are becoming organized into the continuous endothelial lining of the heart known as the endocardium. The outer thin layer of the pericardial cavity comprises the future parietal wall of the adult pericardial cavity. e. Examine serial sections more posteriorly until the gut is very reduced dorsally and the rudiment of the liver is visible as a fingerlike evagination of the ventral wall of the foregut.The liver diverticulum extends into the yolk mass (fig. 5.8 e). Consult figure 5.7 again in order to clarify the physical relationship between the anterior portion of the gut and the liver rudiment. On either side of the notochord, the dorsal mesoderm has segmented to form the somites. Notice the double-layered condition of the lateral mesoderm: the outer layer adjacent to the ectoderm is the somatic mesoderm, while the inner layer adjacent to the endoderm is the splanchnic layer. The cavity between these two layers is the coelom. Between the dorsal mesoderm (somite) and the lateral mesoderm is an area known as the intermediate mesoderm, or nephrotome (sometimes also called mesomeric mesoderm). At this stage, the intermediate mesoderm has enlarged and differentiated into the pronephros, the functional kidney of the larva.The pronephric duct, with its hollow cavity, may also be identifiable. Far posteriorly, locate the proctodaeum, which originates as a ventral invagination of the epidermal ectoderm.

Dissection of the 11-mm Embryo (Stage 25) During stages 23 to 25, a fold of integument, the opercular fold, grows back to enclose and cover the external gills on each side. This is completed sooner on the right side (stage 24) than on the left side (stage 25). The resulting gill chamber remains in communication with the exterior through a single, short funnel on the left side, known as the spiracle. The dissection described in the following for stage-25 embryos may be performed equally as well on stage 24 embryos. 1. Use a wide-mouth pipette to transfer a tadpole to the anesthetic solution. This is a 1⬊3000 solution of ethyl m-aminobenzoate methanesulfonate in either spring water or 10% Amphibian Ringer’s solution. Wait until the tadpole stops all swimming and other movements. Then test it for responsiveness to touch by gently poking it with a blunt instrument. When the tadpole shows no response to touch, transfer it to an operating dish.There should be at least enough of the anesthetic solution in the dish to prevent excessive drying. 2. Place the operating dish on the stage of a dissecting microscope and illuminate it from the side with a focusing illuminator. Turn the tadpole on its back in a shallow groove that you have prepared in your operating dish. Grasp the skin in the anal region (at the base of the tail) with two pairs of watchmaker’s forceps and tear open the ventral body wall. Observe the coiled intestine and attempt to trace its course.The tadpole feeds largely on plants and, like other herbivores, requires a large absorbing surface, provided by the long intestinal tract. Note that great length without great bulk is accomplished by the coiling of the tract into a compact spiral. Note also other abdominal structures such as the liver. Finish exposing the heart if it is not uncovered and examine it and other structures near it. 3. Return to the abdominal area and remove all digestive organs from the body cavity.You should now have exposed the dorsal aspect of the lining of the body cavity.Along the midline, you can observe the area of the developing gonads, the genital ridges (fig. 5.9). Switch your microscope to a higher magnification and carefully observe this area. The two ridges project into the coelomic cavity and are covered by coelomic epithelium (peritoneum). Note the large, light-colored protuberances on the ridges. These are primordial germ cells that have migrated into this area to colonize the gonads. 4. Make sketches of your observations on the internal anatomy of the tadpole.You may have to dissect several tadpoles in order to complete your observations on tadpole morphology.

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Materials EQUIPMENT Compound microscope Dissecting microscope Illuminator Watchmaker’s forceps Operating dish (see Appendix B) “Frog pipette” SOLUTIONS

AND

CHEMICALS

Ethyl m-aminobenzoate methanesulfonate solution (1⬊3000 in spring water or 10% Amphibian Ringer’s solution) LIVING MATERIAL Stage 25 (11-mm) frog embryos COMMERCIALLY PREPARED SLIDES Frog blastula section Early gastrula sagittal section Late gastrula or yolk plug sagittal section Neural plate or early neural groove representative cross sections (rep. c.s.) Neural fold or late neural groove rep. c.s. (some companies supply slides labeled “groove-tube transition”) Early neural tube rep. c.s. Late neural tube rep. c.s. Representative longitudinal sections of 3- or 4-mm frog embryos (selected sections on a “serial sagittal sections” slide can be used as well) Serial cross sections of 3- or 4-mm frog embryos

Some Useful Information Sources WEBSITES http://zygote.swarthmore.edu/index.html This comprehensive site, assembled by Scott Gilbert, Swarthmore College, includes a good deal of information about developmental processes in amphibians and has links to other relevant sites. www.uoguelph.ca/zoology/devobio/dbindex.htm This site contains Stephen Scadding’s supplemental materials for his developmental biology course at the University of Guelph, Ontario. It includes a set of labeled cross sections of frog embryos plus other information about frog development. http://sdb.bio.purdue.edu/SDBEduca/qt_embryos1.html This site, assembled by Laurie Iten, Purdue University, contains movies of serial sections and discussion of the value of serial section study.

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6. Extirpation Experiments, Transplantations, and Parabiosis


L A B O R A T O R Y

6 Extirpation Experiments, Transplantations, and Parabiosis

After William Roux initiated experimentation on frog embryos, other scientists undertook a variety of experimental manipulations of amphibian embryos. Some experiments involved removal (extirpation) of small parts of embryos and careful analysis of the developmental defects that resulted. Other experiments included transplantations of various parts of embryos from place to place within the embryo and even the conjoining of two entire embryos ( parabiosis). Several developmental principles emerged from these “microsurgical” manipulations of amphibian embryos. One of the most important of these is the understanding that the developmental fates of various groups of cells are not rigidly determined early in development. In many cases, if a multicellular fragment of an early frog gastrula is transplanted from one region of the embryo to another, the cells will cooperate with the surrounding cells in their new environment to produce the structures that would normally develop in that part of the embryo. Later in development, however, cells of transplanted fragments develop differently. They tend to continue along the same developmental pathways that they would have followed in their original location. They differentiate as they would have had they not been transplanted and thus produce an anomalous patch of misplaced tissue. Clearly something has happened to the cells that has definitely established (determined) their developmental fates. Many experiments in developmental biology have demonstrated that developmental commitments of cells are determined through interactions with their cellular environments. One such interaction is embryonic induction, a form of cellular communication in which one group of cells transmits a chemical signal, or set of signals, that specifically influences the differentiation of a nearby group of cells.The induction of the lens of the eye by the optic cup is a well-known example of this phenomenon. One of the most famous series of experiments on embryonic induction employing microsurgical techniques and transplantations was performed by Hans Spemann and his brilliant (but tragically fated) student Hilde Mangold. Mangold and Spemann demonstrated the inductive role of archenteron roof tissue in organization of the main body axis in amphibians. More recent experimental analysis of amphibian development using microsurgical manipulation has focused on very early cellular interactions in amphibian embryos.Transplantation of individual cells of 64-cell stage Xenopus embryos have, among other things, clarified the developmental relationships of Spemann and Mangold’s organizer in amphibian embryos (see “Suggestions for Further Investigation” section on p. 90).

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FIGURE 6.1 Steps in the removal of the jelly coats of the frog embryo.

FIGURE 6.2 Steps in the microsurgical removal of part of a neural fold of a frog embryo.

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In this laboratory, you will have the opportunity to perform surgical manipulations on amphibian embryos. Specifically, you will attempt to (a) extirpate (surgically remove) an area of embryonic tissue, (b) transplant a piece of tissue to a new, or foreign, environment, and (c) graft two embryos together side by side. You will be using microsurgical needles and other instruments like those employed over the years in many important experiments on amphibian embryos. Techniques Extirpation of a Lateral Neural Fold This experiment consists of the extirpation (removal) of part of a single lateral neural fold from an embryo at a stage before the folds have come together (stage 14).This type of surgery is referred to as a “defect” experiment.The extirpation of one part, of course, affects the relationships of various other parts of the embryo to one another.What are the chances of survival of an embryo deprived of part of one lateral neural fold? Will a complete neural tube form from the other intact fold? Will the proliferation and invasion of cells adjacent to the wound surface restore or reconstitute the region from which tissue has been removed? 1. The operation should be done under aseptic conditions. The operating medium is Barth and Barth’s inorganic solution containing 0.1% gamma globulin.The composition of the operating medium is given in Appendix B. Perform all phases of the operation itself in Barth and Barth’s medium because it has been demonstrated to promote wound healing. After healing occurs, however, transfer the embryo to spring water or 10% Amphibian Ringer’s solution. 2. Aseptic conditions must be maintained. Before a glass needle is applied to the embryo, the tip of the needle should be dipped in 90% alcohol and then passed through a small beaker containing the operating medium. Some workers prefer to dip the glass needle in boiling water rather than in alcohol.Tungsten needles, as well as all other metal instruments, should be dipped in alcohol and then held over a flame.The stender dish, containing an agar bottom, should be covered with a sterile lid (except, of course, during the operation itself ). 3. With a wide-mouth “frog pipette,” transfer a frog embryo in the early neurula stage of development (stage 14) to an agar-bottomed stender dish containing the operating medium. Before you can operate on an embryo, you must remove its jelly coats and vitelline membrane. Use two pairs of watchmaker’s forceps and work under the low power of a dissecting microscope. Pierce the jelly envelopes with one prong of the left forceps as shown in figure 6.1. A firm grip on the jelly coat may be secured by closing the left forceps.Then, insert both prongs of the right forceps into the jelly coat, closely adjacent to the first prong. Pull the right pair of forceps in a quick motion away from the left pair of forceps.The embryo, still contained within its vitelline membrane, should “squirt” out of its ruptured jelly coats.The same procedure is used in removing the vitelline membrane—the vitelline membrane is firmly grasped with the left pair of forceps and torn apart with a quick jerk of the right pair of forceps. Afterwards, the denuded embryo must be handled carefully. 4. With a ball-tipped glass needle (or a watchmaker’s forceps), make a shallow depression in the agar in which the embryo can “sit” comfortably, dorsal side up. Using a fine-tipped glass needle, remove part of the neural fold on the right side as shown in figure 6.2.With a single complete motion, anterior to posterior, of your glass needle, make a thin tear in the pigmented epidermis very close to the outer edge of the right neural fold. Then tear carefully through the whitish mesodermal cells that lie underneath. When the grayish tan endodermal cells become visible, cut no deeper. Do not cut through these cells, which comprise the roof of the archenteron. If the roof of the archenteron is injured, discard the embryo and start anew with another embryo. Follow the first longitudinal cut or tear with a second longitudinal cut at the inner edge of the right neural fold (fig. 6.2). The subsequent two cuts are transverse cuts, anteriorly and posteriorly, to complete a rectangular area of neural fold tissue. ( Note:The epidermis has a tendency to peel away from the underlying mesoderm; care should be exercised to avoid loosening the epidermal layer.) 5. Lift the entire rectangular area of neural tissue away from the embryo with the tip of the glass needle. The excised piece should include a thick mass of whitish mesoderm with the overlying pigmented ectoderm. The grayish tan roof of the archenteron should not be punctured during the removal of the mesodermalectodermal mass.

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FIGURE 6.3 Postoperative appearance of donor and host at embryonic tail-bud stage 17 (top): a conspicuous feature of the host embryo is the dome-shaped elevation in the ventral surface, the site of the neural fold implant. Self-differentiation of the neural fold graft is evident in the comparisons of the donor (a) and host (b) larvae (bottom). The host larva bears a distinctive circular mass of graft pigment cells on its ventral surface.

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6. Leave the embryo in the operating medium until the wound heals over, usually within an hour.Watch the process of healing: What is the source of cells that spread over the wound surface? After complete healing, transfer the “half-fold” embryo, using a wide-mouth “frog pipette,” to spring water or 10% Amphibian Ringer’s solution in a glass dish (without agar). Maintain the embryo at cool temperatures (15°–18° C) to decrease susceptibility to bacterial infection. Allow the embryo to develop, and record observations of the external features of its development. Make comparisons with control embryos. 7. As indicated in the following, amphibian pigment is derived from the neural crest.The extirpation of one neural fold has resulted in the ablation of an appreciable supply of neural crest. Carefully examine each developing tadpole to detect any effect of neural crest removal on tadpole pigmentation. If facilities are available, rear some of the experimental animals through metamorphosis and note the possible effect on the postmetamorphic pigment pattern.

Transplantation Classic experiments by DuShane in the 1930s revealed that the neural crest elements are precursors of pigment cells.To demonstrate that the pigment cells are derived from the neural crest, we may transplant the crest material to another region of the body. Neural crest cells form a dorsally placed wedge between the lateral wall of the neural fold and the ectoderm on each side of the embryo, so the extirpation of a lateral neural fold also removes a portion of the neural crest. The lateral neural crest elements can then be implanted into the ventral (abdominal) surface of another embryo. 1. As previously described, remove the jelly coat and vitelline membrane from two embryos at stage 14. Place the two embryos, one dorsal side up (donor) and the other ventral side up ( host), in shallow, closely adjacent depressions in an agar-bottomed stender dish. The operating medium, once again, is Barth and Barth’s solution. 2. Prepare a small “pocket” in the ventral surface of the host embryo. Do not attempt to cut a rectangular piece of ventral ectoderm to receive the rectangular piece of neural fold.All that is required is a mere slit or slight hole that can be created by carefully slicing the abdominal ectoderm with the glass needle. The pocket can be enlarged to receive the transplant by carefully removing some of the mesoderm cells with the tip of the glass needle. 3. After you have prepared the host, use the same procedures employed in the preceding section to extirpate part of one lateral neural fold from the right side of the donor embryo. 4. Transfer the neural fold transplant on the tip of the glass needle to the ventral slit, or pocket, of the recipient embryo. It is not necessary to orient the transplant in any particular direction. Gently, yet firmly, push the whole mass through the narrow incision and anchor it against the yolk. It may be necessary to enlarge the pocket slightly to accommodate the transplant, but a tight-fitting transplant is desirable. 5. Once the implant has been pushed into the slit, quickly cover it with the flat surface of a piece from a thin-grade coverslip.The weight of the coverslip should hold the implanted tissue in place. 6. After an hour, the piece of coverslip should be carefully removed. If the transplant has not healed properly, place it under pressure again, with a piece of coverslip, for an additional half hour.After healing is complete, transfer both the donor and the host embryos to a glass dish containing spring water or 10% Amphibian Ringer’s solution. 7. Observe the differentiation of the implanted tissue using figure 6.3 as a guide. You will observe that in the early postoperative stage (stage 18), the relatively bulky implant becomes evident externally as a moderate bulge that will flatten, and the graft will become the center of a circular patch of chromatophores, especially melanophores.The melanophore-laden patch will stand in sharp contrast to the silvery adjacent areas of the host. If you examine the differentiated graft under a dissecting microscope, you will see that the circular graft area has all the aspects of a transparent piece of dorsal skin.

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FIGURE 6.4 Dorsal (a) and ventral (b) views of two tail-bud-stage embryos joined in their developing gill regions.

FIGURE 6.5 Postmetamorphic parabiotic frogs that were united at the tail-bud stage of embryonic development.

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Parabiosis Biologists have been able to take advantage of the remarkable healing powers of amphibian embryos to fuse the cut surfaces of fragments of embryos and produce a variety of artificially combined embryos. It has even proven possible to fuse the front half of one embryo with the rear half of another. Many experiments, however, have involved fusion of two entire embryos in order to produce a parabiotic union in which the two circulatory systems communicate. In grafting two embryos together, various attachments can be made: back-to-back, front-to-front, and side-to-side. However, the most satisfactory position is side-to-side, since this union interferes least with feeding and permits more normal activity. The most suitable time for this operation is shortly after the closure of the neural folds, when the tail bud is well formed (stage 17). Fine glass needles are used for cutting, and the operation is performed under a dissecting microscope. 1. Prepare two grooves in an agar-bottomed stender dish containing the operating medium: the first should be a shallow depression in which an embryo can be placed on its lateral surface.The second groove should be a deep rectangular depression with steep vertical walls, constructed so that the two embryos can fit tightly together (fig. 6.4).The first depression is to be used in creating the wound surfaces in the gill regions of each of the two embryos; the second depression is to be used to hold the two embryos together. 2. After the necessary grooves have been prepared, obtain two embryos in the tail-bud stage of development and remove the jelly coats and vitelline membrane from each. The removal of the membranes should be done in a separate dish, not in the operating dish. 3. Transfer the denuded pair of embryos to the operating dish. With each embryo, in turn, placed right or left side up in the shallow depression, use a glass needle to remove an oval area of ectoderm from the right gill region of one embryo and from the left gill region of the other.The extirpated area should cover the entire protuberance of the gill plate. Care should be exercised to avoid cutting the pronephric swelling just posterior to the gill plate.The wounds in both embryos should be approximately the same size and shape. 4. The cut surfaces will tend to heal rapidly, so immediately after the patches of gill ectoderm have been removed, slip the two embryos into the rectangular depression with the cut surfaces pressed together.The embryos should be dorsal side down and should fit so tightly together that the wound surfaces are in intimate contact.The abdomens of the pair of embryos should protrude slightly above the surface of the rectangular depression.Apply a piece of thin coverslip on top of the pair of embryos.The weight of the coverslip should flatten the abdomens of the embryos, preventing them from moving out of position (recall that beating cilia move embryos at this stage). 5. Wound healing occurs within 3 to 5 hours. After healing is accomplished, remove the piece of coverslip and gently lift the pair out of the depression. With a wide-mouth pipette, transfer the conjoined pair to a finger bowl containing spring water or 10% Amphibian Ringer’s solution. Make frequent observations on the developing embryos and keep the pair alive as long as you can. Fused individuals have actually been reared through metamorphosis to adulthood (fig. 6.5).

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Materials EQUIPMENT Glass microsurgical needles Watchmaker’s forceps Agar-bottom operating dishes (see Appendix B) with lids Ball-tipped glass needle (optional) Wide-mouth “frog pipette” Dissecting microscope Illuminator Pieces of thin-grade coverslips SOLUTIONS

AND

CHEMICALS

Barth and Barth’s operating medium Spring water or 10% Amphibian Ringer’s solution 90% ethyl alcohol LIVING MATERIAL Stage-14 Rana pipiens embryos (for extirpation and transplantation experiments) Stage-17 Rana pipiens embryos (for parabiosis experiment)

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7. Dissociation and Reaggregation of Amphibian Embryonic Cells


L A B O R A T O R Y

7 Dissociation and Reaggregation of Amphibian Embryonic Cells

Many developmental studies have concentrated on the behavior of individual cells that have been isolated from their normal tissue relationships through various dissociation techniques. An early report of such work was H.V.Wilson’s (1907 ) study of the reaggregation of dissociated sponge cells. He found that sponges could be cut up and squeezed through cheesecloth to separate individual cells from one another and from skeletal material.These isolated individual cells moved about actively on the floor of seawater-containing vessels and aggregated into clumps of cells. The clumps ultimately underwent internal reorganization and differentiation to form functional miniature sponges. Johannes Holtfreter and his coworkers conducted a series of experiments in the 1940s and 1950s on the behavior of amphibian cells dissociated from embryos at early developmental stages.They found that dissociated amphibian cells also moved about and reaggregated into clusters, and they observed further that there was continuing cell movement within these reaggregated clusters.This movement resulted in a sorting of cells that ended with specific cell types localized in specific areas of the aggregates.They concluded that this sorting behavior of various cell types was based upon the tissue type or embryonic area from which the cells originally came.They called this tissue-type sorting process “selective affinity.” In the 1950s, Aron Moscona and his colleagues extended research on dissociation and reaggregation of embryonic cells to embryos of homeothermic vertebrates.Their work on chick and mouse embryos contributed a great deal to our understanding of behavioral properties of embryonic cells. For instance, they reported that cells dissociated during early stages of organ formation sorted out in reaggregates and participated in further development with other cells on the basis of tissue type rather than on the basis of species.They found chick cartilage cells reaggregated readily with mouse cartilage cells and cooperated in the production of a cartilage with the two cell types randomly dispersed in a common, continuous matrix. On the other hand, a mixture of chick cartilage and chick liver cells resulted in aggregates in which there was sorting into discrete cartilage and liver areas without any apparent intermingling of cell types. Malcolm Steinberg and his colleagues explored the cellular basis of the sorting that occurs within reaggregates and proposed a quantitative, thermodynamic model for cell sorting involving differential cell surface binding strengths. More recent investigations on cell surface properties by Gerald Edelman and his colleagues have helped to refocus these questions of cell affinities on the development of the intact embryo. They have established the existence of cell adhesion molecules (CAMs) that are tissuetype specific and that appear to be important for the assembly of the precise cell-to-cell contacts that are necessary for normal morphogenesis in developing embryos. References to various studies of dissociated cells and to research on cell surface properties of developing cells are cited in “Suggestions for Further Investigation” on page 90.

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In this laboratory, we will examine basic aspects of reaggregation by dissociated cells of amphibian embryos. Amphibian cells are easier to work with than cells of homeothermic vertebrates ( birds and mammals). Amphibian cells have a much broader range of temperature tolerance, and because of their yolk content, they need not be provided with elaborate nutrient-containing media that also provide good potential growth conditions for a variety of bacteria and molds. You will employ a modified version of a culture technique developed by K.W. Jones and T. R. Elsdale in the 1960s. Jones and Elsdale obtained good reaggregation and, later, differentiation, using Steinberg’s physiological salt solution, supplemented with serum protein, to culture dissociated frog embryo cells. Egg albumin has been found to be an adequate substitute for the serum proteins and will be used in your medium. Techniques Dissociation of Embryonic Cells 1. Obtain a cluster of frog embryos in the late blastula, or very early gastrula, stage of development. With forceps, remove one embryo from the cluster and immerse it in 70% ethyl alcohol for 5 seconds only. This treatment will reduce the number of surface bacteria adhering to the jelly coats without adversely affecting the embryo. Immediately rinse off the alcohol by dipping the embryo in sterile Steinberg’s solution and then transfer the embryo to another container of Steinberg’s solution. 2. Before the cells of an embryo can be dissociated, it is necessary to remove the jelly coats around the embryo. You may remove the jelly coats and vitelline membrane with watchmaker’s forceps as described in Laboratory 6 (refer again to fig. 6.1). It is often preferable, however, to remove the jelly coats from the embryo using the “hand” method, rolling an egg around the palm of one hand. Carefully swab your hands with 70% alcohol and allow them to dry before handling eggs. Place an individual embryo cut loose from the egg mass on the palm of one hand.Then using either a forceps or the index finger of the other hand, gently roll the egg around in your palm.The jelly coats will stick to your hand, and you will be able to remove most of the jelly by continued careful rolling.You will get the knack and develop your own techniques by the second or third attempt. It is also possible to use embryos that have been chemically dejelled (see Laboratory 9). 3. Use a wide-mouth “frog pipette” to transfer the embryo to a small volume (2–3 ml) of dissociating (disaggregating) medium in a small container.This medium is Steinberg’s solution minus calcium and magnesium ions and contains 30 mg of EDTA (versene) per liter. If you use the hand dejelling technique or use embryos that have been chemically dejelled, you will need to break the vitelline membrane after this transfer. Sometimes it is helpful to hold the embryo steady by applying very gentle suction with a Pasteur pipette. With the embryo thus held in place, it is relatively easy to tear the vitelline membrane open with a watchmaker’s forceps.The cells of the embryo should begin to fall apart within a short time, but the process can be aided by gentle manipulation with watchmaker’s forceps or by gently squirting the cells and cell clumps in and out of a Pasteur pipette.

Preparation of Cultures 1. Dissociated cells can be collected from the bottom of the dissociating vessel with a clean Pasteur pipette. A washing step in which the cells are transferred to fresh Steinberg’s solution may be inserted at this point, or the cells may be transferred directly to reaggregating medium in a flat-bottom well slide or to reaggregating medium on a coverslip in a small Stender dish or a 60-mm petri dish.Transfer just enough cells to give a rather sparse scattering of cells on the culture floor. Set up as many cultures as you can with the cells that are available. 2. If you use well slides, set a coverslip over the top of each of them. Prewetting the coverslips by dipping them in Steinberg’s solution will help to prevent the fogging that interferes with later observations.Another method for fogging prevention is to fill each well completely so that the coverslip actually touches the medium. Drying will be prevented if you place a thin coat of vaseline around the rim of the well before putting the coverslip in place. Set the preparation on the stage of a compound microscope and focus the scanning objective on the cells. Try to disturb the cultures as little as possible during subsequent observations. If enough microscopes are available, set up several cultures on the stages of different compound microscopes. You can even observe additional cultures set on the stage of a dissecting microscope.This will be a less satisfactory way to observe cell behavior, but you may be able to detect reaggregation in these cultures also. In all cases, prevent overheating of the cultures by turning on microscope lamps or illuminators only when you are making observations.You may need to dissociate several embryos in order to set up the number of cultures that you want.

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Analysis of Results 1. Observe the process of reaggregation during the next few hours.Watch for several specific aspects of cell activity. Internal cytoplasmic streaming, cyclosis, is frequently observed in dissociated cells, as is the extension of cell surface protrusions. Watch for cell movement and the organization of cell clusters. Sometimes linear clumps, or chains of cells, form as a stage of reaggregation prior to the formation of spherical clusters.You may consider reaggregation to be complete when smooth, dark clusters have been formed. 2. Record the events of reaggregation, including various cellular activities. Note the stages and time progression of the process, as well as any other observations that you might make. One technique for recording results might be to sketch microscope fields of view at various time intervals. 3. You may elect to maintain your cultures for several days in an attempt to observe cell differentiation over a period of time. Maintain the cultures at room temperature and avoid contamination by covering the microscope with a shield of transparent plastic sheeting. After 5 or 6 days, you might be able to observe evidence of histological differentiation of some cells. However, such observations usually require that the reaggregates be fixed, sectioned, and stained for histological examination.

Materials EQUIPMENT Watchmaker’s forceps Wide-mouth “frog pipette” Clean, disposable Pasteur pipettes Glass well culture slides ( plastic culture slides or coverslips set in Stender dishes or small petri dishes may be substituted—see Techniques section) Coverslips Dissecting microscope Illuminator Compound microscopes SOLUTIONS

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CHEMICALS

70% ethyl alcohol Steinberg’s solution Disaggregating solution Reaggregating solution containing 1% egg white LIVING MATERIAL Late blastula or early gastrula frog embryos (see Techniques sections in Laboratory 4 and Laboratory 6)

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8. The Primordial Germ Cells of the Frog


L A B O R A T O R Y

8 The Primordial Germ Cells of the Frog

Late in the nineteenth century, August Weismann proposed that an organism consists of two discrete portions—a germ plasm that functions as a hereditary vehicle and is passed on from one generation to the next, and the remainder of the organism’s body that provides a supporting environment for the germ plasm during the individual’s lifetime. Weismann’s historically interesting concept focused biologists’ attention on the primordial germ cells, the forerunners of eggs and sperm, and provided an important stimulus for the initial attempts to elucidate their development before they can be detected in the developing gonads of vertebrate embryos. In the cases of several vertebrates (and some invertebrates as well), it was discovered that the antecedents of the primordial germ cells had their origins outside the developing gonads and that the cytoplasm later incorporated into primordial germ cells could be identified as early as the two-cell stage. In 1934, Bounoure traced the developmental history of the germ cells of the European frog, Rana temporaria. A specific portion of the cortical cytoplasm in the vegetal hemisphere that has an affinity for certain specific stains was designated the “germ plasm” because it subsequently becomes localized in the primordial germ cells. In the 1950s, Blackler exploited the unique staining response of this germ plasm to the stain Azure A to make more detailed observations of the origin and migration of primordial germ cells. As the cleavage period of frog development comes to an end, this specially staining cytoplasm, with its RNA-rich granules, is located in cells in the blastocoel floor.These cells later leave this endodermal location and migrate to the developing gonads. In the 1980s, additional data were obtained regarding both the very early developmental history of the germ plasm and the mechanisms by which the primordial germ cells make their way from the gut to the developing gonad (see the “Suggestions for Further Investigation” section on p. 91). A series of experiments involving surgical removal or ultraviolet irradiation established that removal, or destructive inactivation, of a specific portion of the vegetal hemisphere of the zygote results in the formation of gonads that lack primordial germ cells. The fundamental experiments that determined the dosages of U V radiation needed to produce sterile gonads and the action spectra of various wavelengths of U V light were reported by L. D. Smith in 1966. This laboratory is designed to repeat some of Smith’s U V irradiation experiments.

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Fortunately, experimental effects on primordial germ cell development can be determined by direct visual observation of the genital ridge of the stage-25 tadpole.The primordial germ cells appear as distinct, light-colored protuberances (fig. 8.1) that can be observed with a dissecting microscope. It is relatively easy to assess the results of experiments affecting the quantity of germ cells in the developing gonads simply by dissecting the tadpole at stage 25. Techniques Irradiation 1. Obtain frog gametes using techniques that you have used previously and proceed with fertilization.When the eggs have been flooded with fresh spring water or 10% Amphibian Ringer’s solution (15–20 minutes after fertilization), work quickly to remove the jelly coats that surround them.The jelly coats must be removed since they might absorb UV radiation strongly enough to prevent it from reaching the egg. It is possible to dissect the jelly coats from the egg by careful manipulation under the dissecting microscope with two pairs of watchmaker’s forceps. For an inexperienced worker, however, this is often a frustrating and tedious task and might take more time than you can afford in this experiment.An alternative technique is to place an individual egg, cut loose from the egg mass, on the palm of one hand.Then use either a forceps or the index finger of the other hand to roll the egg around your hand gently.The jelly coat will stick to your hand, and by careful movement of the egg, you will be able to get most of the jelly off in a short time. You will get the knack and develop your own techniques by the second or third attempt. Quickly place dejellied eggs in a separate container of spring water. Discard any eggs that appear to be ruptured or damaged in any other way. Dejelly as many eggs as you can during a period of 30 minutes, inspect them under the dissecting microscope, and discard, or repeat dejellying, those with significant amounts of jelly remaining. 2. With a wide-mouth “frog pipette,” transfer several dejellied eggs to a quartz microscope slide that you have prepared in advance. (Quartz permits passage of much more UV radiation than glass does.) The slide should have a very thin roll of modeling clay at each end. Permit the eggs to rotate so that the vegetal hemispheres are downward, and then remove most of the water from the slide with a Pasteur pipette or some absorbent material. Place a second slide on top of the first. Flatten the eggs between the two slides by exerting firm but gentle pressure on the upper slide over the rolls of modeling clay. Do not press so hard that the eggs rupture! This flattening will keep the eggs from rotating if you should need to invert the slides. Slight flattening also permits more even irradiation of one hemisphere and reduces the amount of water (which would also absorb UV radiation) intervening between the egg and the slide surface. 3. Make certain that proper safety precautions have been taken in arranging the UV source for your experiments.The UV source should be set up behind a glass barrier. It is convenient, for example, to place the lamp inside a fume hood. Make certain that your skin and eyes are protected at all times, even if the lamp is well shielded, and be careful in positioning slides for irradiation or in removing them after exposure. 4. All experiments on irradiation of the vegetal pole should be accompanied by appropriate control experiments in which the animal hemispheres of other eggs receive an equal dose of radiation. Slides should be placed horizontally below (or above) the UV source, depending upon its positioning. Irradiate either the vegetal hemisphere (experimental group) or the animal hemisphere (control group). If the supply of quartz slides is limited, the second slide in each preparation may be an ordinary glass slide. However, always make certain, both for experimentals and controls, that the portion of the eggs to be irradiated is adjacent to a quartz slide. 5. The dosage of UV radiation received by the eggs will be a function of exposure time and distance from the UV source. Smith (1966) reported that dosages of about 15,000 erg /mm2 of UV irradiation caused depletion of germ cells without causing obviously abnormal development or significant mortality.As there are various types of UV sources, conduct several experimental trials with your particular UV source varying the times and distances widely.This is especially important to do if equipment for measuring UV output is not available.With a Mineralite lamp or a 15-watt germicidal lamp mounted in a fluorescent fixture, for example, a good initial trial dose of irradiation would be a 15-minute exposure at a distance of 45 cm.

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(a)



(b)

FIGURE 8.1 Tadpoles fused in parabiotic union in experiments of E. P. Volpe and S. Curtis designed to test the possibility that amphibian primordial germ cells migrate via the circulation as they do in chick embryos. (a) Unirradiated parabionts at Shumway stage 25; both embryos possess two conspicuous rows of whitish, large primordial germ cells (indicated by arrows). (b) Irradiated embryo (right ), devoid of primordial germ cells, joined to an unirradiated embryo (left ), having two prominent rows of germ cells.


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6. After irradiation, place the eggs in spring water (or 10% Amphibian Ringer’s solution) and allow them to develop to stage 25 (see fig. 4.5). At this stage, the primordial germ cells, if present, can be observed in dissected tadpoles as light-colored bodies clustered on the genital ridges, which are a pair of elevated longitudinal bands of tissue along the dorsal wall of the body cavity (fig. 8.1).The genital ridges are easily observed after the viscera of an anesthetized tadpole have been removed.

Analysis of Results 1. Record the protocol of your experiment. Include data on time from fertilization to irradiation for each group of eggs. It is also very useful to record data on time from fertilization to first cleavage for each group, because such data permit you to describe the time of irradiation as a function of the total fertilization-to-cleavage period.This makes it easier to compare the outcomes of different experiments in which temperature differences or other unavoidable variations in experimental conditions might otherwise complicate comparisons and conclusions. 2. Record data on stages of developmental arrest in those embryos that die. Are there differences between control and experimental embryos in the total numbers of developmental arrests or the stages at which they occur? Are there differences from the other groups of embryos that you have observed earlier? 3. Prepare a table that includes results in terms of germ cell quantities related to UV dosages (either measured and calculated dosages or your distance and exposure time data). Designations such as “many” (or “normal?”), “some,”“few,” and “none” (or “sterile?”) probably will be necessary because exact numbers of germ cells are often difficult to determine.

Materials EQUIPMENT Wooden-handled probe (dissecting needle) Scissors and forceps for dissection Disposable Pasteur pipettes, other clean pipettes, or medicine droppers Petri dishes Clean scalpel or section lifter Watchmaker’s forceps 4-inch finger bowls or other containers for developing eggs Wide-mouth “frog pipettes” Quartz microscope slides and clean glass microscope slides Dissecting microscope Illuminator UV radiation source SOLUTIONS

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CHEMICALS

Spring water or 10% Amphibian Ringer’s solution (see Appendix B) Modeling clay Ethyl m-aminobenzoate methanesulfonate solution (1⬊3000 in spring water or in 10% Amphibian Ringer’s solution) LIVING MATERIAL Pituitary-injected female Rana pipiens Male Rana pipiens

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9. Experimenting with Xenopus Development


L A B O R A T O R Y

9 Experimenting with Xenopus Development

The African clawed frog, Xenopus laevis, has been the subject of a great deal of research on vertebrate development, especially in investigations of cellular interactions in cell differentiation specification during establishment of basic body patterns. For many reasons, Xenopus is also being used increasingly in developmental biology teaching laboratories. Xenopus adults are hardy and easy to maintain, and they are responsive at all times of the year to hormonal stimulation of reproductive activity. Xenopus can be reared in captivity in great numbers, and its use can relieve demand on wild frogs whose populations appear to be in decline in some areas (see p. 238). Development of Xenopus embryos proceeds more rapidly than that of Rana pipiens, so more aspects of development can be observed in a shorter time, although Xenopus’ rapid development can result in some early developmental stages being missed. This laboratory includes basic techniques for working with Xenopus embryos and several experiments on developmental processes in Xenopus. Cleavage and gastrulation stages of Xenopus superficially resemble those of Rana pipiens (shown on p. 47), but Xenopus development proceeds more rapidly (compare table 9.1 with table 4.1, p. 39), and there are developmental differences between the species in some processes such as specific cell movements during gastrulation. As development continues, Xenopus embryos and tadpoles come to differ more noticeably from those of Rana pipiens. Selected normal stages of Xenopus development are illustrated in figure 9.1. Xenopus tadpoles hover steadily in the water and filter out small food particles. This feeding behavior is in marked contrast to Rana pipiens tadpoles, which attach and feed by rasping material off plant surfaces. If you wish to feed Xenopus tadpoles and observe continuing development, consult Appendix C for sources of more information on methods.

IN VITRO FERTILIZATION Some observations of Xenopus embryos and experiments of Xenopus development can be conducted with embryos obtained from matings of hormonally stimulated Xenopus pairs, but others require access to zygotes shortly after fertilization or the availability of a number of embryos at the same stage of development.These latter needs can be met by in vitro fertilization techniques similar to those described for use with Rana pipiens in Laboratory 4.

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(a) Stage 5 1.5 mm



(b) Stage 7 1.5 mm

(c) Stage 111//2 1.5 mm

(d) Stage 17 1.6 mm

(e) Stage 26 3.0–3.3 mm

(f) Stage 35/36 5.3–6.0 mm

(g) Stage 45 8–10 mm

(h) Stage 51 28–36 mm

(i)

St. 48

St. 49

St. 50

St. 51

St. 52

St. 53

St. 54

FIGURE 9.1 Selected Nieuwkoop and Faber stages in the development of the African clawed frog, Xenopus laevis. Ages given are approximate times required to reach the various stages at room temperature (22–24° C). Measurement of greatest dimension is given for each stage because the embryos are drawn at several different scales. (a) Stage 5: Age 2 3/4 hr. Advanced 16-cell stage. Vegetal pole cells (macromeres) are completely separated by cleavage grooves. (b) Stage 7: Age 4 hr. Early (large cell) blastula. (c) Stage 11 1/2: Age 11 3/4 hr. The large yolk plug is almost completely circular. Its diameter is about one-third the diameter of the entire embryo. (d ) Stage 17: Age 18 3/4 hr. Late neural fold stage. Anterior, lateral enlargements of the neural folds are eye rudiments. (e) Stage 26: Age 1 day, 5 1/2 hr. First muscular contractions produce movements. Eyes and auditory vesicles protrude, gill area is distinctly grooved, and pronephric kidney rudiment is visible. Myotomes are evident, indicating segmental muscle development. (f ) Stage 35/36: Age 2 days, 2 hr. Beginning of hatching. Pigment cells (melanophores) are visible on the back, the heart has been beating for several hours, and gill rudiments are visible. (g) Stage 45: Age 4 days, 2 hr. Feeding begins. Tentacle rudiments are present and the operculum partly covers the gills. The intestine is coiled with one and one-half revolutions. (h) Stage 51: Age about 17 days. Swimming, feeding tadpole. Hind-limb bud is conical in shape; its length is about one and one-half times its breadth. Forelimb bud appears oval in lateral views. (i ) Sketches of the structure of the hind limbs at several stages up to stage 54 (about 26 days after fertilization).

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TABLE 9.1 Xenopus Development at 22 to 24° C Stage*

Description

Approximate Age in Hours

1 2 3 5 7 9 11 13 15 17 19 22 24 28 35/36 45 53 58

Fertilization 2-cell 4-cell 16-cell Large cell blastula Small cell blastula Horseshoe-shaped blastopore Slit blastopore Early neural fold Late neural fold Neural tube 1-day Muscular response Transparent outer fin Hatching Feeding Paddle limbs Forelimb eruption

0 1.5 2 2.75 4 7 11.75 14.75 17.5 18.75 20.75 24 26.25 32.5 50 98 576 (24 Days) 1056 (44 Days)

*Stages after Nieuwkoop and Faber (1956).

Techniques 1. Obtain a female Xenopus that has been injected with human chorionic gonadotropin. Use the stripping technique described on page 37 to see if she is releasing eggs. 2. If you have a female that is releasing eggs, dissect one testis from a humanely sacrificed male Xenopus (see p. 35). ( Wrap the male frog in wet paper towel and place it in the refrigerator so that the second testis will be available for subsequent use.) Clean the testis by rolling it on a piece of paper towel.Then place it in a watch glass or a very small finger bowl with about 10 ml of diluted Amphibian Ringer’s solution. Macerate the testis with clean forceps or dissecting needles. 3. Wait for about 5 minutes to allow sperm to become active. Strip eggs from the female into a petri dish that has previously been rinsed with dilute Ringer’s solution. Pipette several ml of the sperm suspension over the eggs.To assure complete exposure to sperm, draw up the sperm suspension in the pipette and spread it over them again. Repeat this procedure two or three times during the next 5 minutes. Record the time of initial insemination as the time of fertilization. 4. Fifteen minutes after insemination, flood the petri dish with diluted Amphibian Ringer’s solution. Zygotes can now be transferred to other containers for observation or experimental manipulation.

Materials EQUIPMENT Dissecting microscope and illuminator Compound microscope Wooden-handled probe (dissecting needle) for pithing Scissors and forceps for dissection Watchmaker’s forceps Watch glasses or small finger bowls Petri dishes “Frog pipette” Pasteur pipettes Finger bowls or other rearing dishes

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SOLUTIONS

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CHEMICALS

Diluted (10%) Amphibian Ringer’s solution LIVING MATERIAL Hormonally stimulated female and male Xenopus

EXPERIMENTING WITH PRECLEAVAGE PROCESSES Subcortical cytoplasmic rotation, which occurs after sperm entry, is a key event in embryonic axis specification. Inductive interactions critical for axis determination do not occur normally if this rotation is disrupted by treatments such as altered gravity relationships, chilling, increased atmospheric pressure, or UV radiation. Subcortical rotation is accomplished by the action of microtubules. Normal microtubule assembly and microtubule-powered cytoplasmic motility can be disrupted by chilling because these processes are very temperature sensitive. In this experiment, eggs are chilled during the critical period after insemination when these cytoplasmic movements are occurring. For more information, consult Scharf and Gerhart (1983) cited on page 93 in “Suggestions for Further Investigation of Amphibian Development.” Techniques 1. Before fertilization, prepare 100-ml beakers containing about 40 ml of chilled (1–2° C) diluted Amphibian Ringer’s solution. These beakers may have been in a controlled temperature chamber long enough to come to temperature, or they may have been in water in a larger container set in an ice-water bath. Check the temperature of the chilled Ringer’s solution before proceeding with fertilization. 2. Proceed with in vitro fertilization and maintain the eggs in diluted Amphibian Ringer’s solution at room temperature until 50 to 60 minutes after insemination. 3. Pipette a few eggs into chilled, diluted Ringer’s solution and leave them in place for 4 minutes. 4. Carefully remove the beakers from the chamber or bath. If you have two beakers with chilled eggs, allow one to warm gradually to room temperature.Warm the eggs in the other beaker quickly by carefully pouring off the chilled dilute Ringer’s solution and replacing it with room temperature dilute Ringer’s solution. If you have only one experimental beaker, choose one of these methods of warming. 5. When the Ringer’s solution has come to room temperature, cover the beakers to prevent excessive evaporation. Your instructor may suggest that you carefully transfer the eggs to appropriate rearing dishes for observations of further development. Place unchilled control embryos in similar dishes adjacent to the dishes containing the experimentally chilled embryos.

Observations 1. Examine both the experimental and control embryos frequently over the next 2 or 3 days. 2. Pay particular attention to development of axial structures, such as neural tube and somites. Compare heads of experimental and control embryos. Are there tail abnormalities in the experimental embryos? Are there any embryos that develop as largely disorganized masses of cells? Is there evidence of swelling that might indicate fluid accumulation (edema)? 3. Continue to make comparisons between experimental and control embryos, at least until there are hatched, swimming tadpoles among the controls.

Materials EQUIPMENT Dissecting microscope and illuminator “Frog pipette” 100-ml beakers

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Finger bowls or other rearing containers Ice bucket or other container for chilling diluted saline solution or temperature-controlled chamber set at 1–2° C SOLUTIONS

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CHEMICALS

Diluted (10%) Amphibian Ringer’s solution LIVING MATERIAL Xenopus zygotes just after fertilization (see p. 81 for materials needed)

CHEMICAL METHOD FOR EGG JELLY REMOVAL A number of experimental treatments require removal of the jelly coats around eggs at the time of ovulation.A mechanical technique for jelly removal was described on page 61.There are several chemical treatments for jelly removal, one of which is described in the following. Techniques 1. Fifteen minutes after insemination, transfer the eggs to be dejellied to a 100-ml beaker containing about 40 ml of a 2% solution of cysteine HCl in diluted Amphibian Ringer’s solution. 2. At half-minute intervals, gently swirl the eggs and observe them with a dissecting microscope.As the jelly coats are removed, the eggs come to lie closer together.The time required to remove jelly coats varies among egg batches, but usually is between 2 and 7 or 8 minutes. Begin rinsing the eggs just before the jelly is completely removed because the cysteine HCl will continue to act during the first rinse, and excessive treatment, which will damage the eggs, must be avoided. Gently pour off the cysteine HCl solution and add diluted Ringer’s solution. Gently swirl the beaker, pour off this solution, and add fresh diluted Ringer’s. Repeat this washing procedure at least once more, but be gentle in handling the eggs, as they are no longer protected from damage by their jelly coats. 3. The dejellied eggs are now ready for observation and experimentation. 4. This technique can also be applied to developing embryos through cleavage and early gastrulation stages if done with great care.

Materials EQUIPMENT 100-ml beakers “Frog pipette” Finger bowls or other rearing containers Dissecting microscope and illuminator SOLUTIONS

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CHEMICALS

Diluted (10%) Amphibian Ringer’s solution 2% Cysteine HCl in diluted Amphibian Ringer’s solution (pH adjusted to 7.8 with NaOH ) LIVING MATERIAL Xenopus zygotes just after fertilization (see p. 81 for materials needed)

LITHIUM AND PATTERN FORMATION It has been known for many years that early developmental processes in a number of animals are disrupted by exposure to lithium ions.The nature of cellular responses to lithium has been investigated only more recently, and it now seems clear that lithium interferes with some cell signaling processes. In amphibian embryos, it appears that lithium ions alter the specificity of the cells’ responses to inductive signals, particularly in the specification of various mesodermal regions. Dorsal, axial structures

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develop from cells that would otherwise contribute to ventral structures.These alterations can have dramatic effects on the establishment of body patterns. In this experiment, early Xenopus embryos will be exposed to lithium ions. Techniques 1. Embryos used in this experiment should have their jelly removed but remain inside their vitelline membranes.The embryos may be ones that have been chemically dejellied or ones that have been dejellied mechanically (see p. 61). 2. Pipette dejellied embryos into a 0.3 M LiCl solution in a 100-ml beaker. The embryos should have between 32 and 128 cells (embryos between stage 5 and stage 7, p. 80) if possible. ( Exposure of later blastula-stage embryos gives interesting, but quite different, results if the only embryos you have available are more advanced.) 3. Leave the embryos in the LiCl solution for 6 to 8 minutes if they have been chemically dejellied or for 12 to 15 minutes if they were mechanically dejellied. 4. Wash the embryos by gently pouring off the LiCl solution and replacing it with diluted Amphibian Ringer’s solution. Repeat the washing twice more by pouring off the medium and replacing it with fresh diluted Ringer’s solution. 5. Gently transfer the treated embryos and appropriate control embryos to rearing dishes where they can be maintained for further observation.

Observations 1. Examine both the experimental and control embryos frequently over the next 2 or 3 days. For information and guidance, consult Kao et al. (1986), Kao and Elinson (1988), and Kao and Danilchik (1991), cited on page 92 in “Suggestions for Further Investigation of Amphibian Development.” 2. Pay particular attention to any tendency for dorsal and anterior structures to be more prominent than usual, relative to posterior and ventral structures. In other words, watch for exaggerated head development and stunted tail development. Watch also for swelling indicative of fluid accumulation (edema). 3. Continue your comparisons between experimental and control embryos until there are hatched, swimming tadpoles among the controls.

Materials EQUIPMENT Dissecting microscope and illuminator 100-ml beakers Finger bowls or other rearing containers SOLUTIONS

AND

CHEMICALS

Diluted (10%) Amphibian Ringer’s solution 0.3 M LiCl in diluted Amphibian Ringer’s solution LIVING MATERIAL Dejellied Xenopus embryos, preferably having between 32 and 128 cells (embryos between stage 5 and stage 7). If embryos are obtained from a mated pair, sort embryos to find some at appropriate stages.

INHIBITING GASTRULATION WITH TRYPAN BLUE The effects of the polysulfonated dye Trypan Blue on vertebrate development were first discovered in investigations of teratological agents, that is, substances that cause grossly abnormal development. Trypan Blue was found to interfere with the cell movements of gastrulation. In several more

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recent studies on the effects of inhibited gastrulation, small quantities of Trypan Blue have been injected into the blastocoel (see Kao and Danilchick 1991, cited in “Suggestions for Further Investigation of Amphibian Development,” p. 92). In this experiment, we will briefly immerse embryos in a Trypan Blue solution.This is a technically easier, though somewhat less precise, method of introducing Trypan Blue into embryos, but it was the method used when the effect was first discovered (Waddington and Perry 1956, cited in “Suggestions for Further Investigation of Amphibian Development,” p. 92). Techniques 1. Embryos should have their jelly removed but remain inside their vitelline membranes.The embryos may be either chemically or mechanically dejellied (see p. 61). 2. Transfer some dejellied embryos having between 8 and 32 cells (about stage 5 embryos) to a 0.025% Trypan Blue solution in a 100-ml beaker. If you do not have embryos at these stages available, substitute slightly more advanced embryos. In any case, make certain that you record the developmental stages of the embryos transferred. 3. Put dejellied control embryos into diluted Amphibian Ringer’s solution in an identical beaker and place it near the experimental beaker. Loosely cover all the beakers to retard evaporation. 4. After 24 hours, gently wash the experimental embryos by carefully pouring off the Trypan Blue solution and adding diluted Ringer’s solution. Repeat this gentle washing procedure several times until the solution around the embryos is clear. 5. Gently transfer the treated embryos and appropriate control embryos to rearing dishes where they can be maintained for further observation.

Observations 1. Observe and compare Trypan Blue-treated and control embryos at the time that the experimental embryos are washed. Pay particular attention to the development of the head region of the embryos. Are there differences in neural tube development, eye development, or olfactory placode development? Are there any embryos in which gastrulation appears to have been completely inhibited, that is, the embryo is an unorganized ball of cells? Are there tail abnormalities in the treated embryos? Is there any evidence of swelling that might indicate fluid accumulation (edema)? 2. Compare control and treated embryos at intervals over the next several days, at least until there are hatched, swimming tadpoles among the controls. 3. Compare results with those of your classmates. Are there any evident correlations between severity or types of abnormality and the stage of development at which Trypan Blue treatment was begun?

Materials EQUIPMENT Dissecting microscope and illuminator “Frog pipette” 100-ml beakers Finger bowls or other rearing containers SOLUTIONS

AND

CHEMICALS

Diluted (10%) Amphibian Ringer’s solution 0.025% Trypan Blue solution in diluted Amphibian Ringer’s solution LIVING MATERIAL 8- to 32-cell (about stage 5) Xenopus embryos, or slightly more advanced embryos—if embryos are obtained from a mated pair, sort embryos to find some at appropriate stages

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Some Useful Information Sources WEBSITES www-cbd.ups-tlse.fr/organismes/xenopus.html This website, assembled at the Centre de Biologie du Developpement in Toulouse, France, contains illustrations of the Xenopus developmental stages defined by P. D. Nieuwkoop and J. Faber, as well as an atlas of Xenopus development and links to sources of related information, including care and breeding of Xenopus. http://sdb.bio.purdue.edu/ This is the very useful website of the Society for Developmental Biology. It includes several sources of information about amphibian development, which can be found by following links to “Virtual LibraryDevelopmental Biology” and “Education.” http://worms.zoology.wisc.edu/frogs/welcome.html This website, prepared by Jeff Hardin, University of Wisconsin, is a good source of information on several aspects of Xenopus development. www.luc.edu/depts/biology/dev.htm This is Bill Wasserman’s Developmental Biology Page from Loyola University of Chicago. It contains a listing of Web resources, several of which provide information about development of amphibians, including Xenopus. www.utexas.edu/courses/zoo321/ This is a website prepared by Klaus Kalthoff, University of Texas, for his developmental biology course. Information about amphibian development can be found by following the links to “Movies” and “Related Web Sites.” http://gto.ncsa.uiuc.edu/pingLeto/xenopus.html This website, prepared by Alan Beck, San Diego, California, contains information about care and maintenance of Xenopus adults and provides some interesting background information about the animal. VIDEO—A DOZEN EGGS

This video includes a video sequence of frog gastrulation, photographed by Ray Keller and John Shih, University of California, Berkeley. The video was produced under the auspices of the Society for Developmental Biology and is available from Sinauer Associates, Inc., P. O. Box 407, 23 Plumtree Road, Sunderland, MA 01375-0407.

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Suggestions for Further Investigation of Amphibian Development


SUGGESTIONS FOR FURTHER INVESTIGATION OF AMPHIBIAN DEVELOPMENT There are many good sources of additional information about amphibian development, including your textbook and the general sources given in the reference list. Amphibian embryos have been studied in thousands of descriptive and experimental studies that have elucidated many principles, not only of amphibian development, but of vertebrate development in general. Some important classical historical papers on amphibian development are available in Willier and Oppenheimer (1964). Viktor Hamburger’s book on experimental embryology (Hamburger 1988) emphasizes work in Hans Spemann’s laboratory, work in which Hamburger himself participated. Fässler and Sander (1996) describe the discoveries of the very gifted Hilde Mangold who suffered a tragic death in an accident at the age of twenty-six. The suggestions that follow are limited to topics introduced in the laboratories included in this manual, but other specific aspects of amphibian development can readily be pursued through various search programs. Frog Development Patterns Developmental patterns of the leopard frog, Rana pipiens (Shumway 1940), are familiar to biologists and biology students in North America, and many important investigations of amphibian development have been done on the embryos of this frog. But for a number of reasons, the African clawed frog, Xenopus laevis, has largely supplanted the leopard frog in research and is being used increasingly in teaching laboratories (Deuchar 1972; Dawid and Sargent 1988; Kay and Peng 1991). Details of cell movement and interaction during gastrulation have been investigated in Xenopus (Keller 1986; Keller et al 1991), and the cortical rotation in the egg that follows sperm contact has been thoroughly studied in Xenopus (Gerhart et al 1989; and see Experiments with Precleavage Processes, in a following section.). Investigators using improved cell marking techniques have clarified details of several basic developmental processes (Saint-Jeannet and Dawid 1994; Davis and Kirschner 2000; Chalmers and Slack 2000). Possibly the most exciting recent advances attained studying Xenopus have had to do with inductive interactions in the early embryo (Gerhart et al 1986; Gerhart 1987). Growth factors similar to, or identical with, ones that act in regulating growth in cell cultures and in vivo in adult organisms appear to act during inductive events in amphibian embryonic pattern determination (Wylie 1990;Thomsen et al 1991; Jessel and Melton 1992). It is not yet fully clear what exact role these diffusible control factors might have in establishing specific regional differentiation patterns (Smith et al 1989; Marx 1991; Green and Smith 1991; Dawid 1994; Green 1994; Slack 1994). Some very old problems in amphibian development are being revisited in light of this research on diffusible factors. For example, the importance of regional specificity of the archenteron roof in inducing the anteroposterior differentiation of the nervous system has been assumed since Otto Mangold’s classic experiments. And yet, normal anteroposterior expression of four genes, including three homeobox genes, was obtained in a system where the mesoderm and the responding ectoderm were kept end to end, without any possibility of specific vertical signals passing between them (Doniach et al 1992). Thus, planar signals passing through the ectoderm from end to end can induce the anteroposterior pattern. If such signaling is a part of normal development, additional explanations for anteroposterior regional specification in nervous system differentiation must be sought. It is interesting that such planar induction has not been demonstrated in Rana pipiens embryos (SaintJeannet and Dawid 1994). Even the history of some famous discoveries has been reexamined. For example, it has been suggested (Lenhoff 1991) that Spemann’s development of the concept of embryonic induction (Hamburger 1988) may have been anticipated and inspired by work that Ethel Browne Harvey did on Hydra regeneration in the very early 1900s.

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Surgery A number of aspects of amphibian development have been investigated through the use of microsurgical techniques. For example, developmental biologists analyzing pigment pattern development in frogs employed some very bold microsurgical techniques pioneered by the great American experimental embryologist Ross Harrison (1908) in his studies of nervous system development. Harrison used tissue culture (a technique then in its infancy), surgical separation, and recombination of large pieces of developing embryos to elucidate principles of nervous system development. He cut embryos transversely and swapped anterior halves between embryos. Such chimeric embryos actually developed into normally formed tadpoles and frogs. Following DuShane’s (1945) and Twitty’s (1945) work on pigment pattern development in frogs, Volpe (1963, 1964) reemployed Ross Harrison’s embryo halving and fusing technique to investigate pigment patterns in chimeric frogs. Microsurgery was the key technique in studies of embryonic induction by Hans Spemann and his student and colleague Hilde Mangold (Hamburger 1988). Mangold transplanted blastopore dorsal lip tissue and her results led to development of the “primary organizer” concept. Another Spemann student, Otto Mangold, transplanted pieces of archenteron roof tissue in his study of regional inductive specificity. Nieuwkoop (1973, 1977) did microsurgery on blastulae to combine animal cap ectoderm with vegetal hemisphere cells after the removal of marginal (equatorial) cells. He discovered that mesodermal cell type differentiation was induced in the animal cap tissue. His results helped to open the way to a modern understanding of inductive relations among cells during early development in amphibian embryos. Gurdon and his colleagues (1985) used the same techniques and tissue combinations in their study of specific genetic activation in response to induction by vegetal cells. By the 1980s, it had become very clear that induction by Spemann and Hilde Mangold’s “primary organizer” was by no means the first demonstrable set of cellular interactions in amphibian development (see earlier “Frog Patterns” section). Again, microsurgical techniques played a key role in the pivotal discoveries. For example, Dale and Slack (1987) did an elegant series of experiments in which they made various combinations of inducing and responding cells of the 32-cell stage Xenopus embryo. If you would like more information on applications of microsurgical techniques, consult your text, the papers just cited, and papers cited in the “Frog Patterns” section. Dissociation Study of the behavior and reaggregation of dissociated cells began with the experiments of H. V. Wilson (1907). His original sponge reaggregation experiments are interesting to repeat and can be spectacularly extended by mixing cells from two species of sponges that differ in pigmentation. Tom Humphreys also used sponge cells to examine the mechanism of cell aggregation. His work is of interest because he used techniques developed in Aron Moscona’s laboratory, such as rotating cultures that permit greater repeatability and quantification of the results of reaggregation experiments (see Humphreys 1963). Jones and Elsdale (1963) found that the ultimate course of the differentiation of reaggregated frog embryo cells was altered if the protein component of the medium was subjected to heating prior to use. Comparison of differentiation following reaggregation in media containing slightly heated or unheated egg albumen could be made.The original report, by an undergraduate student, on the use of egg albumen in reaggregation medium (Bradfield 1967) indicated that there was a concentration effect.The relationship of concentration of albumen to the rate of reaggregation could be studied further.You may also wish to compare the techniques of Barth and Barth (1959) with those of Jones and Elsdale (1963). Giudice (1962) dissociated the cells of sea urchin embryos and found that they would reaggregate, and reorganize, the general framework of the embryos. Other workers have extended this research on dissociated echinoderm embryos. (See “Suggestions for Further Investigation,” p. 27). Aron Moscona’s papers on his work with dissociated and reaggregated chick embryos trace the early development of a fascinating area of research. Malcolm Steinberg’s interesting papers present his quantitative, thermodynamic model of cell sorting.

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Another historically interesting extension of the dissociation studies was the work of Weiss and Andres (1952).They dissociated (by grinding up!) tissues from young chick embryos of pigmented breeds and injected the cells into blood vessels of older embryos. They interpreted their results as indicating that cells disseminated by the vascular route selectively lodged on the basis of tissue type in the organs of the host.Their interpretation was based largely on the fate of pigment cells. Malcolm Steinberg’s quantitative models focused attention on cell surface interactions such as those involving cell adhesion molecules (CAMs). Studies on CAMs by Gerald Edelman and others helped to relate the cell surface properties studied in dissociated cells to cell activities in embryonic morphogenesis (Steinberg 1996). Primordial Germ Cells The primordial germ cells of vertebrate embryos are a particularly interesting group of cells, both because they are ancestors of the gametes that the individual will eventually produce and because of their intriguing migration during development. In the developing frog embryo, the origin of primordial germ cells can be traced back to a specific area of cytoplasm, known as the germinal plasm, located in the vegetal area of the egg. During cleavage, this cytoplasm becomes incorporated into the cells that give rise to primordial germ cells (Bounoure 1934; Blackler 1958; Wiley 1980). Experiments by Blackler on Xenopus laevis, the South African clawed frog, have provided evidence that primordial germ cells give rise to all gametes that ultimately develop in the gonad (Blackler 1962). This countered the view that some gametes may not be descendants of the primordial germ cells but were induced to become gametes by the presence of primordial germ cells. Blackler (1965) also demonstrated that primordial germ cells develop in harmony with the gonad that they colonize and differentiate into the type of gamete characteristic of that gonad (that is, egg or sperm), regardless of their own genetic constitution.You may find the radical transplantation techniques that Blackler employed in these experiments interesting. Volpe and Curtis (1968) joined tail-bud stage (stage 17) frog embryos at the gill region in parabiotic unions to examine the possibility of vascular transfer of germ cells between parabionts. In unions of vegetal pole-irradiated embryos with unirradiated embryos, the irradiated member of the pair invariably lacked germ cells.These results provided evidence against the migration of frog primordial germ cells via the vascular route. These experiments might be extended by examining parabiotic unions involving fusions of other areas of the body surface (see Rugh 1962). As with other aspects of frog development, much of the more recent research on primordial germ cells has been done on Xenopus. Some experiments have focused on the reorganization of germinal plasm after fertilization (Ressom and Dixon 1988) and on the factors determining the number of primordial germ cells formed (Akita and Wakahara 1985).Research on effects of ultraviolet irradiation on primordial germ cell development in Xenopus included investigation of UV irradiation of early embryos (Thomas et al 1983) and of oocytes (Holwill et al 1987). If you wish to do additional experiments on primordial germ cell development, you might begin by consulting the papers cited, especially L. D. Smith’s (1966) paper on his fundamental UV irradiation experiments. Metamorphosis and Regeneration You have had opportunities to investigate early development in amphibians, and most of the topics in these “Suggestions for Further Investigation” are related to those early developmental processes. In this section, however, there are some suggestions about several aspects of later stages of development that you might find interesting. Metamorphosis and its hormonal control have been investigated focusing on Xenopus tail resorption (Brown et al 1996; Elinson et al 1999). Several studies have employed isolated, cultured tails of Xenopus larvae (Weber 1961; Schaffer 1963; Nishikawa 1989). It is relatively easy to set up experiments with isolated Xenopus larvae tails, and if you want to repeat or extend these experiments, good basic information on techniques is available in Scadding (1984) as well as in the papers just cited.

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Amphibians have been favorite subjects for research on regeneration for many years. Some of the most extensive studies have involved the axolotl,Ambystoma mexicanum (Schreckenberg and Jacobson 1975; Stocum 1979; Armstrong and Malacinski 1989), but limb regeneration in Rana pipiens (for example, Fry 1966) and Xenopus larvae has also been investigated. In 1962, Dent explored the developmental loss of limb regenerating capacity that occurs in Xenopus, and work on Xenopus tail (Maden and Corcoran 1996) and limb regeneration is continuing (Korneluk and Liversage 1984; Sessions and Bryant 1988; Filoni and Paglialunga 1990; Endo et al 2000). Wound healing (Radice 1980) and eye regeneration (Underwood and Ide 1992) have also been studied in Xenopus tadpoles. E. M. Deuchar (1975) worked with much earlier developmental stages when she examined tail-bud regeneration between stages 26 and 39. If you want to study regeneration in Xenopus, you might begin by repeating or extending Deuchar’s experiments or Hauser and Lehmann’s (1962) very interesting experiments on regeneration in isolated, cultured Xenopus larvae tails. Vitamin A and similar compounds, such as retinoic acid, have striking effects on the regeneration process. These effects have been studied in axolotls (for example, Scadding 1989; Ludolph et al 1990) and in Xenopus (Scadding and Maden 1986 a & b; Scadding 1989).Although there are unanswered questions about what was once thought to be an established role for retinoic acid as a morphogen during normal limb pattern formation, the effects of retinoic acid on limb regeneration certainly are fascinating. If you wish to investigate these effects further, consult the papers cited or one of several reviews of retinoic acid effects and other molecular aspects of limb regeneration (Tsonis 1990; Bryant and Gardiner 1992; Muneoka and Sasson 1992). You will find other general and theoretical information about limb development in your textbook and in papers such as those by Bryant et al (1981) and Muneoka et al (1986). Lithium and Other Chemical Treatments The study of effects of lithium ions on developmental processes has a long history, and lithium effects on amphibian development and differentiation have been under investigation for some time (for example, Barth and Barth 1967). Lithium ions disrupt normal anteroposterior pattern development in Xenopus embryos in a dose- and timing-dependent maneuver (Kao et al 1986; Kao and Elinson 1987; Klein 1991; Drysdale and Elinson 1993). Cellular mechanisms of lithium effects are being elucidated (Berridge et al 1989; Klein and Melton 1996), and it appears that a lithium effect on a cellular signal transduction system could indeed be involved in lithium’s pattern-disrupting effect on amphibian development (Maslanski et al 1992). For many years, biologists tested the effects of literally hundreds of substances on developmental processes. Many of these studies could be classified as part of teratology, the study of grossly abnormal development.Teratology provided clues regarding birth defects and other unexplained instances of abnormal development. But more recently, results of earlier teratology studies have suggested approaches to be employed in modern investigations of developmental mechanisms. For example,Waddington and Perry’s (1956) study of the teratogenic effect of the dye Trypan Blue suggested a tool that could be used in investigation of the effects of disrupted gastrulation (Gerhart et al 1989). Other similar applications are currently being made (Kao and Danilchick 1991). Experimenting with Precleavage Processes Several microtubule-dependent events in early development can be disrupted by chemical or cold treatments that interfere with microtubule assembly. One of these is resumed meiosis, which normally leads to completion of the second meiotic division and establishment of the haploid egg pronucleus. Cold treatment within a short period of time after insemination interferes with the process so that two sets of maternal chromosomes are retained. Subsequent fertilization events establish a triploid genome in the zygote. If you wish to experiment with this response, you might try an initial treatment of 15 minutes of chilling at 2–3° C, starting between 10 and 15 minutes after insemination (Smith 1958; Kawahara 1978). Consult Laboratory 4 for methods of preparing cells for chromosome observation.Artificially produced

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triploidy has been used as a cell marker in a number of developmental studies (for example, Smith et al 1988; Rollins-Smith and Blair 1990).You may wish to consult these papers regarding methods and applications. Cortical cytoplasmic rotation, another event that occurs shortly after insemination, is also subject to disruption by several experimental treatments. Interference with cortical rotation disrupts subsequent development so that there is deficient development of anterodorsal parts of the embryo (Gerhart et al 1989). Inductive interactions critical for axis determination do not occur normally if cortical rotation is disrupted.This has been demonstrated in experiments where normal rotation was impeded by altering gravity relationships, chilling, applying increased atmospheric pressure, or treating with UV irradiation (Scharf and Gerhart 1983; Vincent et al 1986; Vincent and Gerhart 1987). These experiments showed clearly that there are important determinative events that precede even the beginning of cleavage.Although there is still far to go, our understanding of the cell biology of early development is growing. Developmental Diversity in Amphibians The basic pattern of frog development presented in textbooks is based on the development of several North American and European species as well as Xenopus.This pattern includes embryonic development leading to a tadpole that feeds for some time before undergoing metamorphosis. There are other patterns. In some frog species, the free-swimming tadpole phase is very much shortened or even absent.There are frogs that produce large eggs that develop quickly into small froglets, bypassing the free-swimming tadpole phase. Some even have internal fertilization.These patterns allow for completion of development without access to water, or in some cases, with unpredictable timing of access to water. The familiar pattern of reproduction in North American frogs involves no parental care of offspring, but some Central and South American species brood their developing young in various ways. Some species actually have brood pouches within which development proceeds. You can learn more about these fascinating frogs by consulting papers by Del Pino and Elinson and their colleagues. An almost bizarre form of parental care occurs in an Australian frog, Rheobatrachus silus (Corben et al 1974;Tyler and Carter 1981).A female of this species swallows her young and broods them in her stomach until they undergo metamorphosis and pop out of her mouth as small frogs! During the time that she is brooding young, the female frog refrains from eating and suspends normal digestive functioning, including all gastric glandular secretion. Sadly, this interesting gastric-brooding species seems to have disappeared from its limited range in southeastern Queensland, Australia. Between the early 1970s, when its reproduction was first studied, and the early 1980s, numbers declined dramatically. By the early 1990s, it appeared that this amazing frog might even be extinct. Unfortunately, this pattern of decline toward extinction is all too common and seems to be the condition of numerous amphibian species worldwide (see Appendix G,Amphibian Conservation section, p. 238). Amphibian Development References General Sources Deuchar, E. Xenopus laevis and developmental biology. Biol. Rev. 47:37–112; 1972. Deuchar, E. Xenopus: The South African Clawed Frog. London: Wiley; 1975. DiBernardino, M.A. Frogs. In:Wilt, F. H.;Wessells, N. K., eds. Methods in Developmental Biology. New York:Thomas Y. Crowell; 1967: 53–74. Elinson, R. P.Amphibians. In: Gilbert, S. F.; Raunio,A. M., eds. Embryology: Constructing the Organism. Sunderland, MA: Sinauer; 1997: 309–329. Fässler, P. E.; Sander, K. Hilde Mangold (1898–1924) and Spemann’s organizer: achievement and tragedy. Roux’s Arch. Dev. Biol. 205:323–332; 1996. Guille, M. Molecular Methods in Developmental Biology; Xenopus and Zebrafish.Totowa, NJ: Humana Press; 1999: Methods in Molecular Biology, vol. 127.

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Gurdon, J. B.African Clawed Frogs. In:Wilt, F. H.;Wessells, N. K., eds. Methods in Developmental Biology. New York: Thomas Y. Crowell; 1991: 75–84. Hamburger, V. A Manual of Experimental Embryology. Chicago: University of Chicago Press; 1960. Hamburger,V. The Heritage of Experimental Embryology: Hans Spemann and the Organizer. New York: Oxford University Press; 1988. Hausen, P.; Riebesell, M. The Early Development of Xenopus laevis: An Atlas of the Histology. New York: SpringerVerlag; 1991. Jones, C.M. et al.Amphibian Embryo. In: Sharpe, P.T.; Mason, I., eds. Molecular Embryology Methods and Protocols. Totowa, NJ: Humana Press; 1999: Methods in Molecular Biology, vol. 97. Kay, B. K.; Peng, H. B., eds. Xenopus laevis: Practical Uses in Cell and Molecular Biology. (Methods in Cell Biology; vol. 36) San Diego: Academic Press; 1991. New, D. A.T. The Culture of Vertebrate Embryos. New York: Logos/Academic; 1966. Nieuwkoop, P. D.; Faber, J. Normal Table of Xenopus laevis. Hamden, CT: Garland Publishing; 1994 (reprint of 1956 edition with new forward by J. Gerhart and M. Kirschner). Rugh, R. Experimental Embryology. 3d ed. Minneapolis: Burgess; 1962. Slack, J. M. W. From Egg to Embryo. 2d ed. Cambridge: Cambridge University Press; 1991. Slack, J. M.W. Xenopus and other amphibians. In: Bard, J. B. L. Embryos: Color Atlas of Development. London:Wolfe; 1994: 37–53. Willier, B. H.; Oppenheimer, J. M. Foundations of Experimental Embryology. Englewood Cliffs, NJ: Prentice-Hall; 1964 (contains reprinted original papers of Roux and Spemann). Wolpert, L. The Triumph of the Embryo. Oxford: Oxford University Press; 1991.

Reviews and Research Papers FROG DEVELOPMENT PATTERNS Chalmers, A. D.; Slack, J. M. W.The Xenopus tadpole gut: fate maps and morphogenetic movements. Development 127:381–392; 2000. Davis, R. L.; Kirschner, M. W. The fate of cells in the tailbud of Xenopus laevis. Development 127:255–267; 2000. Dawid, I. B.; Sargent, T. D. Xenopus laevis in developmental and molecular biology. Science 240:1143–1447; 1988. Dawid, I. B. Intercellular signaling and gene regulation during early embryogenesis of Xenopus laevis. Jour. Biol. Chem. 269:6259–6262; 1994. Deuchar, E. M. Xenopus laevis and developmental biology. Biol. Reviews 47:37–112; 1972. Doniach,T. C.; Phillips, R.; Gerhart, J. C. Planar induction of anteroposterior pattern in the developing nervous system of Xenopus laevis. Science. 257:542–545; 1992. Gerhart, J. Determinants of early amphibian development. Am. Zool. 27:593–605; 1987. Gerhart, J.; Black, S.; Scharf, S.; Gimlich, R.; Vincent, J. P.; Danilchik, M.; Rowning, B.; Roberts, J. Amphibian early development. BioScience 36(8):541–549; 1986. Gerhart, J.; Danilchik, M.; Doniach,T.; Roberts, S.; Rowning, B.; Stewart, R. Cortical rotation of the Xenopus egg: consequences for the anteroposterior pattern of embryonic dorsal development. Development 1989 Supplement 37–51; 1989. Green, J. B.A.; Smith, J. C. Growth factors as morphogens—Do gradients and thresholds establish body plan? Trends in Genetics 7:245–249; 1991. Green, J. B.A. Roads to neuralness: embryonic neural induction as derepression of a default state. Cell 77:317–320; 1994. Jessel,T. M.; Melton, D. A. Diffusible factors in vertebrate embryonic induction. Cell 68:257–270; 1992. Keller,R.E.The cellular basis of amphibian gastrulation.In:Browder,L.,ed.Developmental Biology:A Comprehensive Synthesis; Vol. 2. New York: Plenum, 1986: 241–327. Keller, R.; Shih, J.;Wilson, P.; Sater,A. Pattern and function of cell mobility and cell interactions during convergence and extension in Xenopus. In: Gerhart, J., ed. Cell-Cell Interactions in Early Development. New York: WileyLiss, 1991: 31–62. Lenhoff, H. M. Ethel Browne, Hans Spemann, and the discovery of the organizer phenomenon. Biol. Bull. 180:72–80; 1991. Marx, J. How embryos tell heads from tails. Science 254:1586–1588; 1991. Pogany, G. C. Effects of sperm ultraviolet irradiation on the embryonic development of Rana pipiens. Dev. Biol. 26:336–345; 1971. Saint-Jeannet, J.-P.; Dawid, I. B. A fate map for the 32-cell stage of Rana pipiens. Dev. Biol. 166:755–762; 1994.

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Saint-Jeannet, J.-P.; Dawid, I. B. Vertical versus planar neural induction in Rana pipiens embryos. Proc. Natl. Acad. Sci. 91:3049–3353; 1994. Shumway, W. Stages in the normal development of Rana pipiens: I. External form. Anat. Rec. 78:139–147; 1940. Slack, J. M. W.The inducer that never was. Nature 369:279–280; 1994. Smith, J. C.; Cooke, J.; Green, J. B.A.; Howes, G.; Symes, K. Inducing factors and the control of mesodermial patterns in Xenopus laevis. Development 1989 Supplement 149–159; 1989. Thomsen, G.; Woolf,T.; Whitman, M.; Sokol, S.; Melton, S. A. Activins and the induction of axial mesoderm and dorsanterior structures in Xenopus. In: Gerhart, J., ed. Cell-Cell Interactions in Early Development. New York: Wiley-Liss, 1991: 79–91. Winkelbauer, R. Mesodermal cell migration during Xenopus gastrulation. Dev. Biol. 142:155–168; 1990. Wylie, C. Activins and induction. Nature 347:337–338; 1990. SURGERY Dale, L.; Slack, J. M.W. Regional specificity within the mesoderm of early embryos of Xenopus laevis. Development 100:279–295; 1987. DuShane, G. P. The embryology of vertebrate pigment cells. Part I. Amphibia. Quart. Rev. Biol. 18:109–127; 1945. Gimlich, R. L. Acquisition of developmental autonomy in the equatorial region of the Xenopus embryo. Dev. Biol. 116:340–352; 1986. Gimlich, R. L.; Gerhart, J. C. Early cellular interactions promote embryonic axis formation in Xenopus laevis. Dev. Biol. 104:117–130; 1984. Gurdon, J. B.; Fairman, S.; Mohun,T. J.; Brennan, S.The activation of muscle-specific actin genes in Xenopus development by an induction between animal and vegetal cells of a blastula. Cell 41:913–922; 1985. Harrison, R. G. Embryonic transplantation and development of the nervous system. Anat. Rec. 2:385–437; 1908. Nieuwkoop, P. D.The “organization center” of the amphibian embryo: its origin, spatial organization and morphogenetic action. Adv. Morphog. 10:1–39; 1973. Nieuwkoop, P. D. Origin and establishment of embryonic polar axes in amphibian development. Curr. Top. Dev. Biol. 11:115–132; 1977. Twitty, V. C.The developmental analysis of specific pigment patterns. Jour. Exp. Zool. 100:141–178; 1945. Volpe, E. P. Interplay of mutant and wild-type pigment cells in chimeric leopard frogs. Dev. Biol. 8:205–221; 1963. Volpe, E. P. Fate of neural crest homotransplants in pattern mutants of the leopard frog. Jour. Exp. Zool. 157:179–196; 1964. DISSOCIATION Barth, L. G.; Barth, L. J. Differentiation of cells of the Rana pipiens gastrula in unconditioned medium. Jour. Embryol. Exp. Morph. 7:210–222; 1959. Bradfield, B. The use of egg white as a protein additive in medium for embryonic frog cell reaggregation. Proc. S. D. Acad. Sci. 46:259–260; 1967. Edelman, G. M. Expression of cell adhesion molecules during embryogenesis and regeneration. Exp. Cell Res. 161:1–16; 1985. Edelman, G. M. Cell adhesion molecules in the regulation of animal form and pattern. Annu. Rev. Cell Biol. 2:81–116; 1986. Edelman, G. M.Topobiology. Sci. Amer. May: 7–88; 1989. Giudice, G. Restitution of whole larvae from disaggregated cells of sea urchin embryos. Dev. Biol. 5:402–411; 1962. Giudice, G.; Mutolo, V. Reaggregation of dissociated cells of sea urchin embryos. Advances in Morphogenesis 8:115–158; 1970. Humphreys,T. Chemical dissolution and in vitro reconstruction of sponge cell adhesions: isolation and functional demonstration of the components involved. Dev. Biol. 8:27–47; 1963. Jones, K.W.; Elsdale,T. R.The culture of small aggregates of amphibian embryonic cells in vitro. Jour. Embryol. Exp. Morph. 11:135–154; 1963. Moscona, A. Cell suspensions from organ rudiments of chick embryos. Exp. Cell Res. 3:535–539; 1952. Moscona,A.The dissociation and aggregation of cells from organ rudiments of the early chick embryo. Jour. Anat. 86:287–301; 1952. Moscona, A. Development of heterotypic combinations of dissociated embryonic chick cells. Proc. Soc. Exp. Biol. Med. 92:410–416; 1956. Moscona, A. The development in vitro of chimeric aggregates of dissociated chick and mouse cells. Proc. Natl. Acad. Sci. 43:184–194; 1957. Moscona, A. How cells associate. Sci. Amer. May: 132–144; 1961.

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Spiegel, M.; Spiegel, E. S. Reaggregation of dissociated embryonic sea urchin cells. Am. Zool. 15:583–606; 1975. Steinberg, M. S. Reconstruction of tissues by dissociated cells. Science 141:401–408; 1963. Steinberg, M. S. Does differential adhesion govern self-assembly processes in histogenesis? Equilibrium configurations and the emergence of a hierarchy among populations of embryonic cells. Jour. Exp. Zool. 173:395–434; 1970. Steinberg, M. S. Adhesion in development: an historical overview. Dev. Biol. 180:377–388; 1996. Steinberg, M. S.; Poole,T. J. Cellular adhesive differentials as determinants of morphogenetic movements and organ segregation. In: Subtelny, S.; Green, P. B., eds. Developmental Order: Its Origin and Regulation. New York: Alan R. Liss; 1982: 351–378. Steinberg, M. S.; Takeichi, M. Experimental specification of cell sorting, tissue spreading, and specific spatial patterning by quantitative differences in cadherin expression. Proc. Natl. Acad. Sci. 91:206–209; 1994. Townes, P. L.; Holtfreter, J. Directed movements and selective adhesion of embryonic amphibian cells. Jour. Exp. Zool. 128:53–120; 1955. Weiss, P.; Andres, G. Experiments on the fate of embryonic cells (chick) disseminated by the vascular route. Jour. Exp. Zool. 121:449–488; 1952. Wilson, H. V. On some phenomena of coalescence and regeneration in sponges. Jour. Exp. Zool. 5:245–258; 1907. PRIMORDIAL GERM CELLS Akita,Y.;Wakahara, M. Cytological analyses of factors which determine the number of primordial germ cells (PGCs) in Xenopus laevis. Jour. Embrol. Exp. Morph. 90:251–265; 1985. Blackler, A. W. Contribution to the study of the germ-cells in Anura. Jour. Embryol. Exp. Morph. 6:491–503; 1958. Blackler, A. W. Transfer of primordial germ-cells between two subspecies of Xenopus laevis. Jour. Embryol. Exp. Morph. 10:641–651; 1962. Blackler, A. W. Germ-cell transfer and sex ratio in Xenopus laevis. Jour. Embryol. Exp. Morph. 13:51–61; 1965. Bounoure, L. Recherches sur la lignee germinale chez la grenouille rousse aux premiers stades du developpement. Ann. Sci. Nat. 10e Ser. 17:67–248; 1934. Holwill, S.; Heasman, J.; Crawley, C. R.; Wylie, C. C. Axis and germ line deficiencies caused by u.v. irradiation of Xenopus oocytes in vitro. Development 100:735–743; 1987. Ressom, R. E.; Dixon, K. E. Relocation and reorganization of germ plasm in Xenopus embryos after fertilization. Development 103:507–518; 1988. Smith, L. D. The role of germinal plasm in the formation of primordial germ cells in Rana pipiens. Dev. Biol. 14:330–347; 1966. Thomas,V.; Heasman, J.; Ford, C.; Nagajski, D.;Wylie, C. C. Further analysis of the effect of ultra-violet irradiation on the formation of the germ line in Xenopus laevis. Jour. Embryol. Exp. Morph. 76:67–81; 1983. Volpe, E. P.; Curtis, S. Germ cell chimerism: absence in parabiotic frogs. Science 160:328–329; 1989. Wylie, C. C. Primordial germ cells in anuran embryos:Their movement and its guidance. BioScience 30(1):27–31; 1980. METAMORPHOSIS

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REGENERATION

Armstrong, J. B.; Malacinski, G. M.; eds. Developmental Biology of the Axolotl. New York: Oxford University Press; 1989. Brown, D. D.; Wand, Z.; Furlow, J. D.; Kanamori, A.; Schwartzman, R. A.; Remo, B. F.; Pinder, A.The thyroid hormoneinduced tail resorption program during Xenopus laevis metamorphosis. Proc. Natl. Acad. Sci. 93:1924–1929; 1996. Bryant, S. V.; French, V.; Bryant, P. J. Distal Regeneration and Symmetry. Science 212:993–1002; 1981. Bryant, S.V.; Gardiner, D. M. Retinoic acid, local cell-cell interactions, and pattern formation in vertebrate limbs. Dev. Biol. 152:1–25; 1992. Dent, J. N. Regeneration in larvae and metamorphosing individuals of the South African clawed toad. Jour. Morphol. 110:61–77; 1962. Deuchar, E. M. Regeneration of the tail bud in Xenopus embryos. Jour. Exp. Zool. 192:381–390; 1975. Elinson, R. P.; Remo, B.; Brown, D. D. Novel structural elements identified during tail resorption in Xenopus laevis metamorphosis: lessons from tailed frogs. Dev. Biol. 215:243–252; 1999. Endo, T.; Tamura, K.; Ide, H. Analysis of gene expression during Xenopus forelimb regeneration. Dev. Biol. 220:296–306; 2000. Filoni, S.; Paglialunga, L. Effect of denervation on hind limb regeneration in Xenopus laevis larvae. Differentiation 43:10–19; 1990. Fry, A. E. Hind limb regeneration in Rana pipiens larvae. Copeia (3):530–534; 1966.

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Hauser, R.; Lehmann, F. E. Regeneration in isolated tails of Xenopus larvae. Experientia 18:83–84; 1962. Korneluk, R. G.; Liversage, R.A.Tissue regeneration in the amputated forelimb of Xenopus laevis froglets. Can. Jour. Zool. 62:2383–2391; 1984. Ludolph, D. C.; Cameron, J. -A.; Stocum, D. L.The effect of retinoic acid on positional memory in the dorsoventral axis of regenerating axolotl limbs. Dev. Biol. 140:41–52; 1990. Maden, M.; Corcoran, J. Role of thyroid and retinoid receptors in the homeotic transformation of tails into limbs in frogs. Dev. Genetics 19:85–93; 1996. Muneoka, K.; Holler-Dinsmore, G.; Bryant, S. V. Intrinsic control of regenerative loss in Xenopus laevis limbs. Jour. Exp. Zool. 240:47–54; 1986. Muneoka, K.; Sassoon, D. Molecular aspects of regeneration in developing vertebrate limbs. Dev. Biol. 152:37–49; 1992. Nishikawa, A.; Kaiho, M.; Yoshizato, K. Cell death in the anuran tadpole tail: Thyroid hormone induces keratinization and tail-specific growth inhibition of epidermal cells. Dev. Biol. 131:337–344; 1989. Radice, G. P.The spreading of epithelial cells during wound closure in Xenopus laevis. Dev. Biol. 76:26–46; 1980. Scadding, S. R.Thyroxine induced resorption of Xenopus laevis tail tissue in vitro. Jour. Biol. Ed. 18:82–84; 1984. Scadding, S. R.; Maden, M. Comparison of the effects of vitamin A on limb development and regeneration in Xenopus laevis tadpoles. Jour. Embryol. Exp. Morph. 91:35–53; 1986a. Scadding, S. R.; Maden, M.The effects of local application of retinoic acid on limb development and regeneration in tadpoles of Xenopus laevis. Jour. Embryol. Exp. Morph. 91:55–63; 1986b. Scadding, S. R. Vitamin A inhibits amphibian tail regeneration. Can. Jour. Zool. 65(2):457; 1987. Scadding, S. R. Histological effects of vitamin A on limb regeneration in the larval axolotl, Ambystoma mexicanum. Can J. Zool. 68:159–167; 1989. Schaffer, B. M. The isolated Xenopus laevis tail: a preparation for studying the central nervous system and metamorphosis in culture. Jour. Embryol. Exp. Morph. 2:77–90; 1963. Schreckenberg, G. M.; Jacobson,A. G. Normal stages of development of the axolotl, Ambystoma mexicanum. Dev. Biol. 42:391–400; 1975. Sessions, S. K.; Bryant, S.V. Evidence that regenerative ability is an intrinsic property of limb cells in Xenopus. Jour. Exp. Zool. 247:39–44; 1988. Stocum, D. L. Stages of forelimb regeneration in Ambystoma maculatum. Jour. Exp. Zool. 209:395–416; 1979. Tsonis, P. A. Molecular approaches in limb development and regeneration. Trends in Biochemical Sciences. 15:82–83; 1990. Underwood, L. W.; Ide, C. F. An autoradiographic time study during regeneration in fully differentiated Xenopus eyes. Jour. Exp. Zool. 262:193–201; 1992. Weber, R. Induced metamorphosis in isolated tails of Xenopus larvae. Experientia 18:84–85; 1961. LITHIUM

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OTHER CHEMICAL TREATMENTS

Barth, L. G.; Barth, L. J. Competence and sequential induction in presumptive epidermis of normal and hybrid frog gastrulae. Phys. Zool. 40:97–103; 1967. Berridge, M. J.; Downes, C. P.; Hanley, M. R. Neural and developmental actions of lithium: a unifying hypothesis. Cell 59:411–419; 1989. Drysdale,T. A.; Elinson, R. P. Head ectodermal patterning and axial development in frogs. Amer. Zool. 33:417–423; 1993. Gerhart, J.; Danilchik, M.; Doniach,T.; Roberts, S.; Rowning, B.; Stewart, R. Cortical rotation of the Xenopus egg: consequences for the anteroposterior pattern of embryonic dorsal development. Development 1989 Supplement 37–51; 1989. Kao, K.; Danilchik, M. Generation of body plan phenotypes in early embryogenesis. In: Kay and Peng (see General Sources section earlier); 1991: 271–284. Kao, K. R.; Elinson, R. P. Dorsalization of mesoderm induction by lithium. Dev. Biol. 132:81–90; 1988. Kao, K. R.; Masui, V.; Elinson, R. P. Lithium-induced respecification of pattern in Xenopus laevis embryos. Nature 322:371–373; 1986. Klein, P. S.; Melton, D. A. A molecular mechanism for the effect of lithium on development. Proc. Natl. Acad. Sci. USA 93:8455–8459; 1996. Klein, S. L. Xenopus dorsal pattern formation is lithium-sensitive. Roux’s Arch. Dev. Biol. 199:427–436; 1991. Maslanski, J. A.; Leshko, L.; Busa, S. B. Lithium-sensitive production of inositol phosphates during amphibian embryonic mesoderm induction. Science 256:243–245; 1992. Waddington, C. H.; Perry, M. M.Teratogenic effects of Trypan Blue on amphibian embryos. Jour. Embryol. Exp. Morph. 4:110–119; 1956.

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CORTICAL ROTATION

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COLD-INDUCED TRIPLOIDY

Gerhart, J.; Danilchik, M.; Doniach,T.; Roberts, S.; Rowning, B.; Stewart, R. Cortical rotation of the Xenopus egg: consequences for the anteroposterior pattern of embryonic dorsal development. Development 1989 Supplement 37–51; 1989. Kawahara, H. Production of triploid and gynogenetic diploid Xenopus by cold treatment. Dev. Growth & Differ. 20:227–236; 1978. Rollins-Smith, L. A.; Blair, P. Contribution of ventral blood island mesoderm to hematopoiesis in postmetamorphic and metamorphosis-inhibited Xenopus laevis. Dev. Biol. 142:178–183; 1990. (This research included experimental production of triploidy.) Scharf, S. R.; Gerhart, J. C.Axis determination in eggs of Xenopus laevis: a critical period before first cleavage, identified by the common effects of cold, pressure and ultraviolet irradiation. Dev. Bio. 99:75–87; 1983. Smith, P. B.; Flajnik, M. F.;Turpen, J. B. Experimental analysis of ventral blood island hematopoiesis in Xenopus: embryonic chimeras.Dev.Biol.131:302–312;1988.(This research included experimental production of triploidy.) Smith, S. Induction of triploidy in the South African clawed frog, Xenopus laevis (Daudin). Nature 181:290; 1958. Vincent, J.-P.; Gerhart, J. C. Subcortical rotation in Xenopus eggs: an early step in embryonic axis specification. Dev. Biol. 123:526–539; 1987. Vincent, J.-P.; Oster, G. F.; Gerhart, J. C. Kinematics of gray crescent formation in Xenopus eggs: the displacement of subcortical cytoplasm relative to the egg surface. Dev. Biol. 113:484–500; 1986. AMPHIBIAN DEVELOPMENTAL DIVERSITY Corben, C. J.; Ingram, G. J.;Tyler, M. J. Gastric brooding: unique form of parental care in an Australian frog. Science 186:946–947; 1974. Del Pino, E. M. Marsupial Frogs. Sci. Amer. May: 110–118; 1989. Del Pino, E. M. Modifications of oogenesis and development in marsupial frogs. Development 107:169–187; 1989. Del Pino, E. M.; Elinson, R. P. A novel development pattern for frogs: gastrulation produces an embryonic disk. Nature 306:589–591; 1983. Del Pino, E. M.; Loor-Vela, S. The pattern of early cleavage of the marsupial frog Gastrotheca riobambae. Development 110:781–789; 1990. Elinson, R. P.; Del Pino, E. M.;Townsend, D. S.; Cuesta, F. C.; Eichhorn, P. A practical guide to the developmental biology of terrestrial-breeding frogs. Biol. Bull. 179:163–177; 1990. Elinson, R. P. Change in developmental patterns: embryos of amphibians with large eggs. In: Raff, R. A.; Raff, E. C., eds. Development as an Evolutionary Process. New York: Alan R. Liss; 1987: 1–21. Elinson, R. P. Fertilization and aqueous development of Puerto Rican terrestrial-breeding frog, Eleutherodactylus coqui. Jour. Morph. 193:217–224; 1987. Elinson, R. P.; Del Pino, E. M. Cleavage and gastrulation in the egg-brooding marsupial frog, Gastrotheca riobambae. Jour. Embryol. Exp. Morph. 90:223–232; 1985. Tyler, M. J.; Carter, D. B. Oral birth of the young of the gastric brooding frog Rheobatrachus silus. Anim. Behav. 29:280–282; 1981.

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III. Chick Embryo Development

10. Patterns of Chick Development


L A B O R A T O R Y

10 Patterns of Chick Development

The study of the chick embryo has a long history that reaches back to the time of Aristotle and includes such great names in the history of biology as William Harvey, C. F.Wolff, and many others. In modern times, the chick embryo has been exploited in so many developmental studies that it can safely be said that the developing chick is one of the most intensively studied of all organisms. Interest in patterns of chick development goes beyond the basic description of avian ontogeny because there are marked parallels between early chick development and the development of mammalian embryos, which are more difficult to obtain and study.This laboratory is intended to introduce you to the general processes of chick development and to some of the techniques used in the study of chick embryos. Many of the early events in chick ontogeny are not as readily observed as are similar processes in the frog. One reason for this difficulty is the organization of the chicken’s “egg.” The egg proper (the “yolk” in everyday terminology) is surrounded by secretory products of the female reproductive tract that include the white (or albumen), the shell membranes, and the shell (fig. 10.1). All of these accessory layers are added after the time of fertilization.The egg itself is a large, extremely yolky cell with a restricted less yolky cytoplasmic area on one side that contains the nucleus. As the large mass of yolk is not cleaved, the mitotic divisions of cleavage are restricted to this small active area, the blastodisc.As cleavage proceeds, the blastodisc is converted into a multicellular structure that is several cells thick. Further development proceeds within this active area, which is called the blastoderm after cleavage divisions have made it multicellular. You will begin your study of chick development by removing blastoderms from the yolk and examining living chick embryos at several stages of development. First, you will briefly examine an embryo that has been incubated for 2 or 3 days.After this initial exposure to embryo handling techniques, you will make more detailed observations of an embryo at the stage of development commonly called the “33-hour chick embryo.”This embryo clearly displays many results of the completion of earliest developmental processes. Following this introduction to the living 12- to 15-somite chick embryo, you will examine embryos at several other stages of development and trace some general patterns of chick development.

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vitelline membrane

blastoderm dense albumen less dense albumen

outer shell membrane air space chalaza inner shell membrane

shell

yolk

FIGURE 10.1 The organization of a chicken’s egg and accessory material surrounding it.

FIGURE 10.2 Opening the incubated egg into saline solution.

FIGURE 10.3 Cutting the blastoderm free from the yolk.

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Observations Learning to Handle Chick Embryos You can make only limited observations of living chick embryos while they remain in place on the yolk surface, so you need to learn techniques for removing the embryo and surrounding blastoderm tissue from the yolk and for transferring the embryo to a dish for further observations.You can begin your study of chick development by removing a 2- or 3-day blastoderm from the yolk.You will study details of embryos at this developmental stage later, so use this particular embryo only to practice the removal technique and to become familiar with the relationships of the embryo and blastoderm to the yolk and the vitelline membrane that encloses them. 1. Pour sterile saline solution into a finger bowl to a depth of about 1.5 cm. Crack the incubated egg out into the saline (fig. 10.2). It is sometimes helpful to crack the shell gently around at least half of its circumference before attempting to open it. Hold the egg very near the saline as you apply pressure to separate the two halves. If the yolk breaks in such a way that the blastoderm is badly torn, discard the egg and saline and start over with a new egg. The blastoderm usually will be in view when the egg has been broken into the bowl, but it may be necessary to gently turn the yolk, using a section lifter, spatula, forceps handle, or your finger.The embryo and extraembryonic blood vessels around it should be plainly evident at 2 or 3 days of incubation. If you see only a small, dense, white spot, the egg is probably sterile and ought to be discarded. 2. Firmly grasp the blastoderm near the edge of the area that contains blood vessels with watchmaker’s forceps. Penetrate the yolk about 0.5 cm before closing the forceps. Hold the forceps closed as you begin to cut around the outside of the blastoderm with a sharp scissors (fig. 10.3). This cutting of the blastoderm, and the vitelline membrane that covers it, should be done with a fairly rapid snipping action.You will encounter problems caused by the yolk rushing out at the first cuts unless the blastoderm is held in place at the very top of the yolk. After you have cut around the entire blastoderm, the vitelline membrane and blastoderm area can be gently floated off the yolk into the saline. When the blastoderm is floating free, gently slide it onto a submerged plastic spoon. Then, use your watchmaker’s forceps to hold the blastoderm in place as you slowly lift the spoon out of the saline and transfer it to a Syracuse dish or small petri dish containing saline. 3. Briefly examine the embryo and blastoderm under a dissecting microscope. If the vitelline membrane has also been transferred, use your watchmaker’s forceps to separate it from the blastoderm and examine it.Your instructor also may ask you to make other general observations or to practice turning over the blastoderm. When you have finished, discard the embryo and blastoderm, as well as the yolk, and proceed to detailed observations of stages of chick development.

The Living 12- to 15-Somite (33-Hour) Chick Embryo The length of incubation time required to provide embryos of this stage is somewhat variable and depends upon such factors as the previous handling of the eggs, incubation temperature, and possibly even upon the season of the year. In modern incubators, it usually takes somewhat more than 33 hours of incubation to carry development to this stage. It is, therefore, more accurate to describe the developmental stage of this, or any other, embryo in terms of either a recognized standard such as the Hamburger-Hamilton stages or of a clear morphological criterion such as somite number. 1. Crack an incubated egg and empty its contents into saline as done previously. The yolk is less likely to break at this earlier stage than at more advanced stages of incubation, but if the yolk breaks, discard the egg and saline and start over with another egg. You may need to rotate the yolk to bring the blastoderm into view. At this stage of development, the blastoderm area appears to consist of a series of irregular concentric rings. If there is a single, small, dense, white spot, the egg is probably sterile and ought to be discarded, but be cautious about discarding eggs until you are relatively certain about them. Even in the case of a normal blastoderm, the immediate response of students frequently is,“I can’t see a thing.” If the blastoderm area of yolk protrudes above the level of saline, add enough saline to submerge the blastoderm.

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proamnion

prosencephalon

optic vesicle mesencephalon rhombencephalon

border of foregut

sinus terminalis ventricle vitelline (omphalomesenteric) vein somite area opaca

closed neural tube

open neural folds notochord

primitive streak

area pellucida blood island

FIGURE 10.4 Dorsal view of a 14- or 15-somite chick embryo. prosencephalon

optic vesicle aortic arch I ventral aorta dorsal aorta bulbus cordis

border of foregut

ventricle sinoatrial region vitelline (omphalomesenteric) vein

anterior intestinal portal dorsal aorta somite notochord

FIGURE 10.5 Diagrammatic ventral view of a 14- or 15-somite chick embryo.

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2. Firmly grasp the area just outside the blastoderm with your watchmaker’s forceps. Continue to hold the forceps closed as you cut around the outside of the blastoderm. It is important to maintain your hold on the blastoderm as you complete your circular cut just outside the position of your forceps. If you relax your hold on the blastoderm at this time, it almost certainly will be lost as yolk rushes out through the cuts. After you have cut around the entire blastoderm, the vitelline membrane and blastoderm area can be gently slid off the yolk into the saline. Some yolk will adhere to the blastoderm, and it may be necessary to break the connection of this yolk with that being left behind by cutting into the yolk repeatedly with your scissors. When the blastoderm, with a small amount of adherent yolk, has been floated free, it can be transferred in a wide-mouth “chick pipette” to a Syracuse dish or small petri dish that is about 2/3 full of saline. Several difficulties can be encountered in this step. Since the blastoderm and vitelline membrane tend to stick to the inside of the pipette unless it is completely wetted, it is necessary to flush the pipette several times by repeatedly drawing saline in and out. Another problem is the tendency of stringy albumen to remain attached to the vitelline membrane and to pull the membrane and the blastoderm back out of the pipette during the transfer. This can cause damage to the blastoderm and is a source of considerable frustration. This stringy albumen either can be cut with a scissors or pinched off with a forceps. Sometimes, however, the only effective technique is to hold the flat handle of the forceps over the mouth of the pipette during the transfer process. It is also helpful to hold the pipette as close to horizontal as possible and to make the transfer quickly because there is a tendency for everything to run out of such a wide opening. Gently expel the contents into saline in a Syracuse dish. 3. Further work should be done under a dissecting microscope, and it is frequently helpful to switch from a light to a dark background and back again. Most students find it easier to see details of an embryo if it is illuminated from below or from the side rather than from above. Experiment with various patterns of illumination to find one that gives you a good view of the embryo.The blastoderm may still be covered by the vitelline membrane. If it is lying with the vitelline membrane on top, you will need to turn it over gently. When you have the ventral side exposed, remove the largest yolk granules by picking them off with your forceps or by squirting a gentle stream of saline from a Pasteur pipette. If the vitelline membrane still is attached, free the blastoderm by teasing it loose around the edges, using a forceps and a wooden-handled probe or a microsurgical needle. If this is done carefully, the blastoderm should survive the procedure without injury. It probably will be necessary to transfer the blastoderm to a second Syracuse dish with clean saline before further observations can be made. Be very careful during this final transfer of the fragile, unprotected blastoderm and make certain that the inside of your “chick pipette” is thoroughly pre-wetted. 4. Examine, sketch, and/or describe dorsal and ventral views of the 12- to 15-somite chick embryo using figures 10.4 and 10.5 for reference. It is difficult to locate some of the structures represented in figure 10.4 with certainty in the living embryo, but there are a number of general structural features that can be observed. Identify the somites, which are paired, compact chunks of mesodermal tissue that occur lateral to the neural tube. With your embryo against a dark background, count the number of somite pairs. If your embryo has fewer than 10 pairs, or more than 17 pairs, of somites, it may be difficult to use the descriptions that follow, and you should ask your instructor whether you ought to try another embryo. 5. You will note that there are differences in the apparent consistency of the blastoderm parts surrounding the embryo body proper. The relatively clear area adjacent to the embryo (area pellucida), along with the embryo itself, is separated from the yolk by a fluid-filled space when it is in place in the egg. The remainder of the blastoderm (area opaca) is situated in closer contact with the yolk, and its cells have attached yolk granules.This gives it a more opaque appearance. 6. During your examination of the dorsal side of the blastoderm, gently poke the embryo with a microsurgical needle, probe, or forceps tip in order to clarify its structural dimensions. Note that the embryo has a definite head region that is a clearly delineated tubular structure. Gently lift the head a little to demonstrate this for yourself.The head end of the embryo is, therefore, segregated from the remainder of the blastoderm. In the more posterior areas, you will note continuity of the still-flat posterior part of the embryo with the remainder of the flat blastoderm. The progressive segregation of the originally flat and continuous blastoderm into a rounded embryonic body area and an extraembryonic area is accomplished by a set of processes known as body folding. The necessity for this body folding is a function of the pattern of chick embryo cleavage, which is restricted to the flat disc of active cytoplasm on one side of the egg.

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first cleavage furrow

blastoderm (cellular blastodisc)

discrete cells cleavage furrow

FIGURE 10.6 Surface views of chick embryo cleavage.

blastoderm

yolk subgerminal space

epiblast hypoblast

FIGURE 10.7 Diagrammatic representation of the separation of the hypoblast and epiblast (delamination) in the chick embryo blastoderm. (These sections of blastoderms are more advanced than the blastoderms shown in fig. 10.6.)

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7. Examine the embryo against a light background and look for scattered reddish or orangish spots, the blood islands.These blood islands are early sites of blood cell production and are in the process of fusing to produce blood vessels.You may also be able to see some color in the sinus terminalis, which is a ringlike vessel encircling the margin of the blastoderm area that contains developing blood vessels. The cellular blastoderm does extend beyond the sinus terminalis, so the sinus does not mark the blastoderm margin. 8. While examining the ventral surface of the embryo, observe the heart and the vitelline (omphalomesenteric) veins, the large vessels that enter it posteriorly.You may observe some heart beating, but it probably will be just irregular pulsations. Note that there is an enclosed hollow gut ( foregut) in the anterior part of the embryo that is open posteriorly (fig. 10.5).This posterior opening to the outside is known as the intestinal portal.The foregut at this stage has been described as resembling the finger of a glove. Probe the gut to demonstrate its dimensions to your own satisfaction. 9. You may supplement your study of the 12- to 15-somite embryo by examining a commercially prepared stained slide of an embryo at this stage of development (that is, a “33-hour chick” slide).

Cleavage in the Chick Embryo Now that you have had opportunity to examine an embryo when some of the early phases of development have been completed, we will return to the initial stages of chick development and retrace some of the processes that produced the embryo that you have studied. Fertilization occurs in the upper portion of the hen’s oviduct, and cleavage begins there.The first developmental processes following fertilization occur as the egg passes through the hen’s oviduct and shell gland (uterus), and it is quite difficult to obtain embryos at these stages and prepare them for study.Therefore, we will simply describe some of the highlights of development before egg laying without asking you to make direct observations. Cleavage proceeds as the egg moves through the oviduct. Cleavage in the chick embryo is a form of incomplete (meroblastic) cleavage since the huge mass of yolk is not divided into separate cells. It is called discoidal cleavage because cleavage divisions are restricted to the active cytoplasmic area, the blastodisc. Early in cleavage, mitosis is not followed immediately by complete cytokinesis; that is, the nuclei are segregated by furrows in the center of the blastodisc (fig. 10.6). As additional mitotic divisions occur, the furrowing process spreads peripherally from the center. Soon, however, some discrete cells are produced as cytokinesis is completed in the center of the blastodisc (properly called the blastoderm from this point). Eventually, the blastoderm becomes several cells thick, and a fluid-filled subgerminal space develops underneath it. Gastrulation and Neurulation The beginning of gastrulation, the process that ultimately produces the major body layers (germ layers), also occurs during passage through the hen’s reproductive tract. Some of the cells of the blastoderm separate themselves from the remainder of the blastoderm and move down toward the subgerminal space to form small islands of cells (fig. 10.7).This initial segregation process occurs through the movements of individual cells or small groups of cells.Then a sheet of cells moves forward from a sickle-shaped area at the posterior margin of the blastoderm into the subgerminal space.This sheet incorporates the islands of cells that entered earlier. As a result of these processes, the embryo consists of two layers separated by a space.These layers are the newly formed lower layer, or hypoblast, which is rather loosely organized, and the upper layer, or epiblast, within which the cells are arranged in an orderly epithelial type of organization. This process of separation into two layers is known as delamination. The developmental fate of the hypoblast will be the production of the extraembryonic endoderm of the yolk sac and the yolk stalk, which connects the yolk sac to the embryonic gut. Following delamination, cells of the epiblast move to new positions in the embryo where they will participate in

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prospective extraembryonic ectoderm prospective epidermal ectoderm prospective neural ectoderm prospective notochord

prospective endoderm

prospective mesoderm

FIGURE 10.8 Fate map of the epiblast of the chick blastoderm before migration through the primitive streak commences. Cells of the future notochord, mesoderm, and endoderm areas will leave the surface and pass to the interior through the primitive streak.

Anterior Stage 1

area opaca

Stage 2

area pellucida

thickened area

Posterior

Anterior Stage 3

Stage 3⫹

developing primitive streak

Posterior

FIGURE 10.9 Stages in the formation of the primitive streak (Hamburger-Hamilton stages 1, 2, 3, 3⫹).

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the development of all parts of the embryonic body and several extraembryonic membranes. The developmental roles of the cells in various areas of the original epiblast can be summarized in the form of a fate map, such as the one shown in figure 10.8. At the time that the egg is laid, delamination is arrested and is resumed only if the egg is brought to incubation temperatures. When incubation is initiated, delamination continues, and other organizational changes occur in the blastoderm.A markedly thickened area develops in the epiblast on the posterior side of the blastoderm (fig. 10.9).This thickening is the beginning of the formation of the site of cellular migration that will occur during later stages of gastrulation.This area, the primitive streak, elongates as incubation proceeds. When active cell migration begins in the primitive streak, cells of the epiblast layer move across the surface to congregate at the primitive streak, leave the surface to move down in the direction of the subgerminal space, and then change direction to move out away from the streak to take their new places (fig. 10.10). Some of them spread laterally to establish the mesodermal layer, which is a new, third layer situated between the two previously existing layers. Other cells migrating through the primitive streak move downward where they displace hypoblast cells to establish the midline tissue that becomes the embryonic endoderm that produces the gut. 1. As migration through the streak continues, several structural features of the fully developed, or definitive primitive streak, can be identified (fig. 10.10). The midline of the streak becomes depressed, forming a shallow trough, the primitive groove, that lies between paired primitive ridges. At the anterior end of the streak, there is a particularly marked accumulation of migrating cells, Hensen’s node. Just posterior to this enlargement, the midline portion of the streak is deeply depressed to form the primitive pit.

Henson’s node (primitive knot) primitive pit primitive ridge primitive groove mesoderm cells epiblast

(a)

hypoblast

migrating cells

Henson’s node

primitive pit ectoderm

primitive streak

notochord

mesoderm endoderm

(b)

Anterior

Posterior

FIGURE 10.10 The mechanics of gastrulation in the chick embryo. (a) Diagrammatic cross section of the blastoderm showing the migration of epiblast cells through the primitive streak to produce mesoderm and embryonic endoderm. (b) Diagrammatic longitudinal section of the primitive streak showing the migration of cells that produce the notochord.

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proamnion Hensen’s node notochord

primitive groove primitive ridge

(b)

(a)

head fold

blood islands

(c) FIGURE 10.11 Early stages in the development of the chick embryo. (a) Definitive streak stage (stage 4). (b) Head-process stage (stage 5). (c) Head-fold stage (stage 6). (The stages are defined in V. Hamburger and H. L. Hamilton, 1951, J. Morphol. 88:49–92.) Photographs provided by Dr. H. L. Hamilton with the permission of the Wistar Press.

proamnion head fold anterior intestinal portal neural fold notochord neural plate hensen’s node

FIGURE 10.12 Dorsal view of “24-hour” chick embryo (stage 8, four pairs of somites). (The stage is defined by V. Hamburger and H. L. Hamilton, 1951, J. Morphol. 88:49–92.) Photograph provided by Dr. H. L. Hamilton with permission of the Wistar Press.

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Examine a commercially prepared whole-mount slide of a definitive primitive streak stage ( H.-H. stage 4, 12to 16-hours incubation time) chick blastoderm and identify the areas of the streak and other features of the embryo (see fig. 10.11). 2. During early stages of migration through the primitive streak, the cells of the forming mesodermal layer spread laterally and posteriorly from the streak in the form of cell sheets. Migration of mesoderm in an anterior direction from the streak begins with a cord of cells that has moved through the primitive pit region. This cell cord produces the middorsal supporting element known as the notochord (figs. 10.10 and 10.11). As the notochord develops, the primitive streak begins to regress. The anterior end of the streak retreats, causing a shortening in the length of the streak.The shortening streak leaves behind the lengthening notochord that almost appears to be reeled out by the retreating tip of the primitive streak (fig. 10.11). Examine a commercial whole-mount slide of an 18-hour chick blastoderm. It may be necessary to examine several slides of this stage of development and to focus up and down on the slides in order to satisfy yourself that you have actually seen the notochord and other features of the blastoderm at this stage. 3. Before the completion of gastrulation, other developmental events are already under way.The formation of the central nervous system, neurulation, begins with the formation of a thickened area of ectoderm lying over the region of the notochord. This thickened neural plate rolls up into the cylindrical neural tube by processes that are roughly similar to those seen in the frog embryo. We will examine details of neural tube formation later. At the same time that the neural tube is forming, mesoderm paralleling the midline of the embryo becomes organized into the paired, segmentally arranged somites. As mentioned earlier, somite number is a much more reliable standard for communication of developmental stage than is incubation time because development of additional pairs of somites is closely related to the general level of ontogeny of the whole embryo.The first evidence of the body folding process that segregates the embryo proper from the remainder of the blastoderm can also be seen at about the same time.The head fold has undercut an embryo of this incubation stage to establish the embryo’s head (fig. 10.12). Examine a commercial “24-hour” embryo slide and identify as many of these features as possible. Prepare a living blastoderm of 28 to 30 hours incubation for study, using the technique employed in sections A and B. Identify the major landmarks of the embryo once again and then proceed to investigate the three-dimensional structure of the embryo. Probe the head region from the dorsal side and the foregut from the ventral side in order to clarify the nature and results of the head-folding process. Your instructor may request that you examine sections of the 24-hour embryo or other slide material to amplify the general outlines of early development that we have considered. Whether or not you make further observations on very early stages of chick development, we urge you to expand your understanding of early chick development by filling in details from your textbook and from the references listed in the “Suggestions for Further Investigation” section.

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wall of neural tube

skin ectoderm of head

optic vesicle somatic mesoderm splanchnic mesoderm subcephalic pocket

(a)

dorsal aorta

notochord aortic arch I head mesenchyme

(b)

endoderm

foregut

ventral aorta ectoderm dorsal aorta foregut

wall of neural tube notochord

(c)

coelomic space ventral aorta

dorsal aorta foregut

endocardium of ventricle epimyocardium of ventricle

thyroid rudiment

(d) dorsal aorta

vitelline vein (e)

anterior intestinal portal

FIGURE 10.13 Representative cross sections of 12- to 15-somite chick embryo.

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Internal Structural Details of the 12- to 15-Somite Stage Chick Embryo Now that you have surveyed some basic events in early development and have a general grasp of some of the processes that lead to the formation of the “33-hour” chick, we will return to that stage of development for a somewhat more detailed study. The relationships of the various body layers to one another and their differentiation into specific structural elements of the embryo can be reconstructed through the study of serial sections of the embryo. 1. In order to put your study of serial sections in perspective, reexamine a commercially prepared “33-hour” chick whole-mount slide before you begin to examine serial sections. Note again such features as the differentiation of various areas of the neural tube, the structure and location of the heart, and the apparent extent of progress of body folding in freeing the head from the flat blastoderm (consult figs. 10.4 and 10.5). 2. Begin your study by skimming rather quickly through all of the sections from the anterior tip of the embryo through the last section. You will notice that the embryo apparently is unattached in the first sections that you encounter and that it is separated from the remainder of the blastoderm by a definite space, the subcephalic pocket (fig. 10.13a). As you proceed through the sections, you will reach those where the embryo is no longer physically separated from the remainder of the flat blastoderm.The point where this continuity is first observed is at the end of the free, tubular portion of the embryo, produced by body folding up to this time. Move on through the remainder of the slides noting the continuity of the embryo with the rest of the flat blastoderm and the general shape changes seen in embryonic structures at various levels of the body as you move posteriorly. 3. Return to the first sections of the head and trace the neural tube posteriorly through the body. Within a few sections, the neural tube will broaden laterally (fig. 10.13a).These lateral extensions are the optic vesicles that will produce major portions of the eyes. As you move further posteriorly, the neural tube changes in shape and size as the optic vesicles fade out and various other regions of the brain are sectioned. Eventually, the future spinal cord level of the neural tube is reached. Sections of the neural tube in the spinal cord region are characterized by a narrow, tall, sometimes slitlike, central cavity and a laterally compressed shape. Note also that the floor and roof of the spinal cord are relatively thin compared to the lateral walls.As you continue tracing sections, you will move into the region where the neural tube has not yet closed. In this region, you can identify the open neural groove and the paired neural folds (fig. 10.13g). By moving back and forth through the region of transition from closed to open neural tube, you should be able to mentally reconstruct the process of neural tube closure.You may wish to make sketches of your concept of neural tube closure and later compare them with those in textbooks. 4. During your examination of the neural tube, you undoubtedly will have noticed the circular notochord that lies just ventral to the neural tube. Quickly trace the notochord through its entire extent. Note that it does not extend all the way to the tip of the head, but rather is first encountered at a level of the body just posterior to the point where the optic vesicles fade out. Near its posterior end, sections of the notochord enlarge, and eventually, the notochord fuses with the floor of the open neural plate.As you progress beyond this point, you are entering the area of the primitive streak. It is somewhat difficult to give an exact description of guidelines that indicate that you have reached sections of the streak, but figure 10.13h is a representative section of the primitive streak level of the embryo.As you examine sections that include the primitive streak, you will note that streak tissue, the developing mesoderm, and probably the endoderm as well, are connected.This tissue continuity is characteristic of the levels of the body where the primitive streak is functioning. 5. The foregut (fig. 10.13b) appears in serial sections just posterior to the level at which the optic vesicles fade out.Trace the foregut posteriorly.You will note that it becomes laterally expanded and that it appears to be quite flattened dorsoventrally. However, remember that during your study of the living embryo at this stage, it was possible to insert a substantial object such as the tip of a microsurgical needle through the intestinal portal into the foregut. Continue tracing the foregut until the floor of the gut appears to pull apart within a span of three or four sections.You have now reached the level of the anterior intestinal portal (fig. 10.13e).You should now be able to reconstruct the significance of these sections mentally if you think back to the organization of the living embryo, the structure of the foregut, and the process of body folding that is progressively defining the embryonic body at this stage of development. Posterior to the level of the intestinal portal, the prospective endodermal lining of the remainder of the digestive tract is continuous with the more lateral future yolk sac endoderm.This region of flat endoderm is called the open gut.

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ectoderm somatic mesoderm splanchnic mesoderm endoderm


neural canal somite pronephric cord dorsal aorta notochord

(f) neural groove neural fold

segmental plate notochord (g) primitive groove of primitive streak primitive ridge of primitive streak mesoderm (h)

FIGURE 10.13 Continued.


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A few hours beyond the time when this stage of development is observed, body folding will begin at the posterior end of the embryo and will produce a hindgut and a posterior intestinal portal.Then lateral body folding will progress from both directions, adding to the extent of the foregut and hindgut, thus shaping more of the embryonic body. Ultimately this process reduces the area of the embryo in continuity with the remainder of the blastoderm to the very small umbilical region that includes a small area of open gut.This small persisting area of continuity provides the embryo’s contact with the extraembryonic membranes and a pathway for blood vessels to and from the membranes, and it will be maintained until just before the time of hatching. 6. Figures 10.4 and 10.5 illustrate the general organization of the major portions of the circulatory system of the 12- to 15-somite embryo. Return to the level at which the foregut first appears in your sections, and in that section, or one or two sections anterior to it, note a vertical space on either side of the region of the tip of the gut.These are sections through the first pair of aortic arches (fig. 10.13b, see also fig. 10.5) that loop up around the front of the foregut. Posterior to this point, the laterally expanded gut lies between the paired ventral aortae carrying blood ahead from the heart and the paired dorsal aorta carrying blood back toward the posterior areas of the embryo. Later in development, posterior portions of the dorsal aortae will fuse into a single, large arterial trunk, the descending aorta. Trace sections posteriorly and watch both ventral and dorsal aortae. The ventral aortae appear to fuse into a single vessel.This is actually a section through the anterior end of the heart, the bulbus cordis.As you move posteriorly through the sections, the heart will appear to move sideways from its original position beneath the midline of the embryo.This laterally deflected portion of the heart is the developing ventricle. In subsequent sections, as you continue posteriorly, the heart appears to return to the midline and then to split into two portions. This single midline region and the first portions of the paired vessels represent the sinoatrial portion of the heart that will produce the atrium and sinus venosus regions. These paired vessels continue posteriorly, without any marked boundary, as the vitelline or omphalomesentric veins (fig. 10.13e).These veins drain blood from the developing yolk sac region and return it to the heart. Note the relationship of the vitelline veins to the intestinal portal in your sections (see fig. 10.5).As you continue tracing the vitelline veins posteriorly, they appear to become laterally expanded.This is actually an indication that you have reached the point where the vitelline veins approach the embryo at nearly right angles. All trace of the vitelline veins will soon be lost in succeeding sections. Continue following the dorsal aortae through the sections. Note that there are apparent lateral expansions of the aorta in the posterior region of the body and that, ultimately, the aortae disappear from the sections.These lateral expansions represent the origins of the vitelline arteries that supply the developing yolk sac region. It might be helpful for your understanding of the circulatory pattern to sketch a complete circulatory system with arrows indicating the directions of blood flow. Your instructor may also ask you to read about earlier stages of heart development during which the single tubular heart develops from paired rudiments. 7. Choose a section in the spinal cord region of the body that has well-developed somites (fig. 10.13f ) and study the general relationships of the various body structures and layers to one another.This type of section displays a number of characteristic features of a general body organization pattern that is part of a recognizable stage in the development of all vertebrate embryos. However, remember that at this stage of chick development, the body layers (germ layers) still are stacked on top of one another on a flat plane in this area of the embryo. It might be helpful for you to sketch this section and add details as your study proceeds.The presence of a hollow neural tube and a rodlike notochord just ventral to it are obvious features that need little further mention, but we will examine some other features of this characteristic vertebrate embryonic body plan in more detail.The upper ectodermal and lower ectodermal layers are relatively simply organized, but there are regional specializations in the mesoderm that are important characteristics of this basic body plan.We will concentrate on lateral aspects of the mesoderm after simply asking you to recall the method of formation and structural relationships of the mesodermal notochord. Focus your attention on the somites and scan back and forth through eight or ten sections in this part of the body. Note that the somites appear to be dense and compact in some sections and rather diffuse and loosely organized in others.The sections showing apparently diffuse organization are actually cut through the spaces between the somites, while the dense, compact areas are sections through the somites themselves. These observations should serve as a reinforcement of your understanding of the somites as paired, segmentally arranged blocks of mesodermal tissue.

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metencephalon mesencephalon first (hyomandibular) branchial groove optic cup myelencephalon

auditory vesicle

lens vesicle telencephalon

aortic arch III

bulbus cordis ventricle anterior intestinal portal

margin of amniotic fold vitelline artery

tail bud FIGURE 10.14 Outline drawing of a 29-somite chick embryo (after slightly more than 2 days of incubation).

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Shift your attention laterally from the somite. Just lateral to the somite there is a solid strip of tissue that is known as the nephrotome. This is the portion of the mesoderm from which excretory and reproductive structures will develop. In fact, you may be able to distinguish the very first elements of the developing excretory system, the pronephric cords, in some sections in the area of the embryo under examination (see fig. 10.13f ). The portion of the mesoderm beyond the nephrotome is known as the lateral plate.The lateral plate mesoderm is split into two layers: the lower of these, the splanchnic mesodermal layer, comes to lie adjacent to the endoderm, while the other, the somatic mesodermal layer, is adjacent to the ectoderm.The space between these layers is the coelomic space. 8. One further exercise may aid your understanding of the organization of the basic body plan of this stage of development. Figure 5.8e is a cross section of the trunk of a frog embryo that possesses the same basic structural features that you have observed in the chick embryo. Diagrammatically alter a sketch of this chick embryo section to make it resemble the tubular body of the 3- or 4-mm frog embryo.The significance of this exercise in diagrammatic modification of a chick embryo section goes beyond simply facilitating your comparison of body organizations in the tubular frog embryo and the still-flat chick blastoderm.The structural modifications that you have performed on the chick cross section in this sketch are very similar to the changes brought about by the process of body folding that segregates the embryonic portion of the blastoderm and produces the tubular body form of the advanced chick embryo.

The Chick Embryo After Two Days of Incubation You should now have a good understanding of the internal organization of the “33-hour” chick embryo.We will now turn our attention to later stages of development, which we will consider in a more general way. Our goal is to gain an overview of successive stages in chick development, and we will emphasize those points that will be helpful in your later experimental work.Your instructor may elect to ask you to go into more detail than is offered in the descriptions that follow. If this is the case, consult references cited in the “Suggestions for Further Investigation” section. 1. Figure 10.14 will provide you with guidance for a preliminary examination of a prepared slide of a “48-hour” chick embryo.This embryo is at a stage of development roughly comparable to that of the living 2-day embryo that you will examine. Your instructor will specify the amount of detail required in your examination of brain, circulatory system, pharynx, and other parts of the embryo on the prepared slide, but there are some important general points. Note the orientation of the embryo’s head, which is no longer a simple, tubular structure as it was in the 12- to 15-somite chick.The head has been tipped forward in a process known as flexion.At the same time, the body has begun to twist on its axis in such a way that the anterior portion of the head has come to lie on its left side.This twisting of the body axis is called torsion. The primitive streak has regressed into a restricted area known as the tail bud.The tail bud continues to play a role in organizing the posterior area of the embryo. Note the rather prominent vitelline arteries that extend outward at right angles from the posterior part of the embryo. In a prepared slide, it is more difficult to see the vitelline veins approaching the embryo, but the complex nature of the yolk sac circulation will be obvious in the living embryo. Finally, note the margin of the amniotic fold, which is in the process of covering the embryo.The amniotic fold produces the amnion that will come to surround and enclose the embryo and the chorion that will spread peripherally around the whole complex of embryo and yolk. We will examine some details of amnion production later. 2. The living 2-day embryo can be initially observed on the surface of the yolk. Break an incubated egg into saline in a finger bowl. Crack the egg gently for quite a distance around its circumference and partially submerge it in the saline as you pull the halves apart. Even if you observe these precautions carefully, it is quite possible that you will break the yolk because its consistency is beginning to change at this stage, and yolk breakage is a greater hazard now than at earlier stages. If the yolk breaks very near the body of the embryo, try again. If the blastoderm is not submerged, add enough saline to cover it completely. When the yolk is in the saline, the embryo and the yolk sac vessels should be seen readily because the sinus terminalis now completely encircles an area that is several centimeters in diameter. The blastoderm area should rotate upward, but you may have to gently assist the yolk’s rotation.

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myelencephalon auditory vesicle

acoustico-facialis ganglion aortic arch II first branchial groove gasserian (semilunar) ganglion

glosspharyngeal ganglion

metencephalon

third branchial groove aortic arch IV

mesencephalon lens optic cup

wing bud diencephalon epiphysis olfactory pit telencephalon ventricle leg bud

vitelline vein vitelline artery

tail

FIGURE 10.15 Outline of a 36-somite chick embryo (3 days of incubation), dorsal view.

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Make observations of the circulation pattern with the embryo still in place on the yolk. Attempt to determine which major extraembryonic vessels are arteries and which are veins. Focus on the vessels carefully with your dissecting microscope and check the direction and rate of blood flow in vessels, if possible. Blood will be flowing away from the embryo in arteries and toward it in veins, and flow in arteries shows strong surging pulses, while the venous flow is smoother and more regular. If you are unable to make all of these observations with the embryo in place on the yolk, do so after you have transferred it to a dish. Examine the heart, which has folded on itself to produce a coiled tube, and trace the flow of blood through the beating heart. 3. Further observations can be made more easily after the blastoderm has been removed from the yolk. Grasp the blastoderm with your forceps and cut a circle just outside the sinus terminalis. Pull the blastoderm off and transfer it to a dish of saline for observation, using techniques that you have previously used. You can make this transfer using either a plastic spoon or your chick pipette. If you use the chick pipette, very careful pre-wetting and cautious bulb manipulation are essential for a successful transfer. You probably will find that the vitelline membrane floats off more easily than it did at earlier stages of development. 4. Examine the dorsal side of the embryo and blastoderm.There are general features that should be examined, in addition to structural details of the embryonic body itself. Note that the amniotic folds form a hoodlike covering that has moved back over part of the embryo’s body. Ultimately, the amniotic folds will cover the embryo completely. Use a probe or microsurgical needle to investigate the nature of this relationship. Other features that should be noted are the increased somite number and any aspects of the circulatory pattern that you were unable to observe with the embryo in place on the yolk.All of the extraembryonic vessels observed here are part of the blood supply of the yolk-sac membrane that is spreading peripherally to cover an increasing area of the yolk. Turn the blastoderm over and examine its ventral surface. Note the restricted area of blood vessel connection between the embryo and the yolk sac.The anterior intestinal portal can still be probed, but the structure of the foregut has been altered considerably by head flexion. Body folding in the posterior part of the embryo may be apparent. If so, you should be able to insert an instrument through a posterior intestinal portal into the hindgut. 5. In order to examine structural details of the embryo, it will be necessary to dissect away the amniotic folds. Note the increased extent of the free head. Use figure 10.14 and reference sources for information for study of the embryo itself.

The Chick Embryo After Three Days of Incubation Our study of the 3-day chick embryo and its relationship to extraembryonic structures will emphasize changing general structural patterns, but your instructor may direct you to do more detailed study in some places. 1. Examine a prepared “72-hour” chick slide, using figure 10.15 as a guide. 2. Carefully open an incubated egg in a dish of saline. It is hard to avoid breaking the yolk because of the fluid consistency of the yolk at this stage, but small breaks can usually be tolerated. It is much easier to observe the embryonic and the extraembryonic circulation with the embryo in place on the yolk at 3 days’ incubation than it was in the case of the 2-day embryo. Reaffirm your understanding of the vitelline circulatory pattern. Note that the area enclosed within the sinus terminalis has increased markedly during an additional day of incubation. 3. It usually is not practical to attempt to remove the whole vascular area when transferring the embryo for further study; therefore, make your encircling cut well inside the sinus terminalis. Transfer of the 3-day embryo and blastoderm with a “chick pipette” should not be attempted. It is probably best to use a plastic spoon for this transfer, but if you hold your Syracuse or petri dish close enough to the finger bowl to avoid a long transfer distance, it is possible to pick up the blastoderm with watchmaker’s forceps. Yet another alternative method is to float the blastoderm onto a watch glass and hold it in place as you lift it out of the saline for transfer to your Syracuse or petri dish.The vitelline membrane may be lost in the transfer, but if it is still attached, remove it carefully. 4. You will find that the embryo’s body is largely or, perhaps completely, covered by the amniotic folds by this stage of development. If a small circular uncovered area remains over your embryo, note that in addition to the anterior amniotic fold that you observed in the 2-day chick, there is amniotic folding moving in from the rear

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amniotic fold

ectoderm somatic mesoderm

fusion of amniotic folds

extraembryonic coelom

yolk sac

chorion amnion

amniotic cavity extraembryonic coelom

FIGURE 10.16 Stages in the establishment of the amnion and the chorion by fusion of the amniotic folds.

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and the sides of the embryo.These folds come together in a process that some observers have likened to the closing of a drawstring purse. The amniotic folds are produced by a buckling of the extraembryonic ectoderm and the somatic mesoderm adjacent to it. These folds meet over the body of the embryo and fuse (fig. 10.16). At the point of contact, a separation occurs that leaves two distinct membranes. One is the amnion enclosing the amniotic cavity within which the embryo continues its development.The other is the chorion (serosa) that spreads peripherally to enclose the embryo and yolk (see fig. 10.17 also). 5. Before making further observations of the dorsal view of the embryo, turn the blastoderm over and examine the ventral surface. Examine the relationship of the large yolk sac vessels to the embryo.You will see that the area of entry and exit for large veins and arteries is restricted to a small area, the developing umbilical cord region. Note that just anterior to the tail region, a round vesicle has grown out ventrally from the body.This hollow sac is the developing allantois.The allantois plays several roles in later development. It will serve as a storage depot for excretory wastes produced during the course of development.Also, it will expand distally (see fig. 10.17), spread, and fuse with the chorion to produce a complex, highly vascular membrane known as the chorioallantoic membrane.This membrane plays a critical role as an area of gas exchange, where oxygen diffusing in through the porous shell is absorbed for transport back to the embryo and carbon dioxide is released. It also functions in absorbing calcium from the shell. This absorption supplies necessary calcium for bone development and helps later in development to thin and weaken the shell, which facilitates hatching. The chorioallantoic membrane remains attached by a narrow stalk running through the umbilical cord. 6. It will be necessary to dissect away the chorion and amnion in order to examine details of the structure of the embryo itself (consult figure 10.15). You will observe marked changes in the flexion of the head and the differentiation of various regions of the brain. The clear establishment of a tail region and the development of prominent limb buds also mark this stage of development. Increased complexity in heart structure, in the pharynx, and in the aortic arch region can be seen as well.

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chorion amniotic cavity

amnion albumen (egg white)

embryo allantois

extraembryonic coelom

air space

eggshell yolk sac yolk

FIGURE 10.17 Membrane relationships in the chick embryo. This drawing is highly diagrammatic. It is designed to show structural relationships, not proportional sizes of the various structures.

(a)

(b)

(c)

FIGURE 10.18 Later stages of development of the chick embryo. (a) Stage 36 (10 days). (b) Stage 38 (12 days). (c) Stage 43 (17 days). (The stages are defined by V. Hamilton and H. L. Hamilton, 1951, J. Morphol. 88:49–92.) Photographs provided by Dr. H. L. Hamilton with the permission of the Wistar Press.

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Later Stages of Chick Development 1. Your instructor may elect to provide a series of chick embryos for study that have been incubated for longer periods of time. If so, we would suggest a series that includes embryos of 5, 7 (or 8), 10, and 17 (or 18) days’ incubation.Your instructor may elect to eliminate some stages or to substitute others.The best technique for examination of older stages probably is to locate the embryo by candling and then to tap the egg repeatedly in the area over the embryo until the shell is extensively cracked, but not broken through. Work carefully, using a dissecting forceps (but not your watchmaker’s forceps) to remove the shell piece by piece to expose the membranes and the embryo. Refer to figure 10.17 for a general impression of the relationships of the embryo and extraembryonic membranes. As you examine the various stages, attempt to sort out the relationships of the membranes in each case. For instance, the spreading of the highly vascular allantois and its fusion with the chorion to produce the complex chorioallantoic membrane can be followed by examining embryos between 5 and 10 days of incubation. Note also the extent to which the yolk sac has enveloped the yolk in later stages. Relatively sparse vascularization is a very obvious characteristic of the amnion. 2. Following your examination of membrane relationships in each case, dissect the embryo free from the membranes and study external characteristics such as the development of limbs, feather formation, and changing proportional relationships of the body parts to one another (fig. 10.18). Dissection of the embryos is also informative, if time permits. If you decide to dissect an embryo that has been incubated for more than 7 or 8 days, the embryo should be decapitated before dissection is begun (your instructor will advise you concerning humane sacrifice of chick embryos). Record notes and sketches of these older embryos for later reference in experimental work.

Materials EQUIPMENT Finger bowls Syracuse dishes or small petri dishes Watchmaker’s forceps Dissecting forceps Fine scissors with sharp points “Chick pipette” Plastic spoon Dissecting microscope Illuminator Compound microscope Fine-mouth pipettes (for example, disposable Pasteur pipettes) Tungsten or cactus spine microsurgical needle Wooden-handled probe (dissecting needle) SOLUTIONS

AND

CHEMICALS

Howard Ringer’s solution (or other chick physiological saline solution) LIVING MATERIAL 12- to 15-somite embryos (eggs incubated 38–42 hours) Head-fold embryos (eggs incubated 28–30 hours) “2-day” embryos (eggs incubated 55–60 hours) “3-day” embryos (eggs incubated 75–82 hours) Older embryos—5, 7 (or 8), 10, and 17 (or 18) days of incubation (if assigned) COMMERCIALLY PREPARED SLIDES “33-hour” chick “16-hour” chick “18-hour” chick “24-hour” chick “33-hour” chick “48-hour” chick “72-hour” chick

embryo, whole mount embryo, whole mount (“primitive streak” embryo) embryo, whole mount embryo, whole mount embryo, serial cross sections embryo, whole mount embryo, whole mount

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Some Useful Information Sources WEBSITES http://sdb.bio.purdue.edu/SDBEduca/qt_embryos1.html This site, asembled by Laurie Iten, Purdue University, contains movies of serial sections and discussion of the value of serial section study. http://zygote.swarthmore.edu/index.html This comprehensive site, assembled by Scott Gilbert, Swarthmore College, includes a good deal of information about developmental processes in chick embryos and has links to other sites. www.uoguelph.ca/zoology/devobio/dbindex.htm This site contains Stephen Scadding’s supplemental materials for his developmental biology class at the University of Guelph, Ontario. It includes a nice set of labeled cross sections of chick embryos plus other information about chick development. http://sdb.bio.purdue.edu/ This is the very useful website of the Society for Developmental Biology. It includes several sources of information about chick embryo development. www.ucalgary.ca/UofC/eduweb/virtualembryo/ This is a comprehensive website prepared by Leon Browder, University of Calgary, that has links to sources of information on chick embryo development. www.luc.edu/depts/biology/dev.htm This is Bill Wasserman’s Developmental Biology Page from Loyola University of Chicago. It contains a listing of Web resources, several of which provide information about chick embryo development. www.utexas.edu/courses/zoo321/ This is a website prepared by Klaus Kalthoff, University of Texas, for his developmental biology course. It includes information about chick embryo development and some useful links. VIDEO—A DOZEN EGGS

This video includes a very good video sequence of early chick embryo development photographed by Hilde Bortier, University of Ghent, Belgium.The video was produced under the auspices of the Society for Developmental Biology and is available from Sinauer Associates, Inc., P. O. Box 407, 23 Plumtree Road, Sunderland, MA 01375-0407.

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11. In Vitro Culture of Chick Embryos and Embryonic Heart Duplication


L A B O R A T O R Y

11 In Vitro Culture of Chick Embryos and Embryonic Heart Duplication

The term in vitro fertilization has become part of our everyday conversation because it is used in connection with several treatments of human infertility. The phrase literally means “fertilization in glass.” However, in the context of human medicine, it more generally describes procedures in which both fertilization and early embryonic development occur in culture vessels, away from the environment in which they normally take place. Techniques used in in vitro fertilization for culturing cells and embryos have been derived from culture techniques developed over many years in cell biology and developmental biology research.There are many questions in developmental biology that can be answered only when cells, tissues, or organs can be removed from their normal environment within the developing organism. Such problems often are investigated by maintaining isolated parts (explants) in precisely defined media in culture vessels made of glass (literally in vitro) or plastic. An important common factor in these techniques is that developmental responses are tested in isolation from other parts of the developing system. Unfortunately, the culture media used in these techniques provide favorable environments for the growth of bacteria and molds as well. Most in vitro culture work must be done under rigorously controlled conditions because of this persistent threat of contamination by microorganisms. However, the embryo culture technique devised by Nelson Spratt in the 1940s circumvents some of the difficulties met in other types of culture work because the Spratt culture medium contains egg white, which has antibacterial activity.Although embryos can be maintained in Spratt cultures for only a limited time and the medium’s contents are not defined, Spratt cultures of chick embryos provide a good introduction to in vitro culture techniques. It is interesting to note that during the 1980s, techniques were finally worked out that make it possible to culture chick embryos continuously from the beginning of development to hatching (see “Suggestions for Further Investigation” section on p. 140). In vitro culture techniques allow experiments on embryos at early stages of development that could not be done with embryos in place inside the eggshell. For example, surgical manipulation of the developing heart requires access to the ventral side of the embryo.This lab introduces such an experiment, surgically induced heart tube duplication. Body folding in the chick embryo brings two separate heart rudiments together on the ventral midline. The rudiments fuse and blend together to form a single ventral tubular heart. However, if something interferes with body folding so as to prevent fusion of these two lateral rudiments, each has the potential to form a separate beating heart tube. It is sometimes possible to cause this to happen surgically by making a tiny cut in the region where the heart rudiments are being brought together by the lateral body folds.

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culture medium petri dish cover water

inverted cover of 60 mm petri dish

FIGURE 11.1 Cross section of culture vessel.


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IN VITRO CULTURE OF CHICK EMBRYOS Techniques Preparation of Culture Medium Before you begin, wash the table surface and arrange your equipment and instruments for easy access during your work. Because preparation of materials is such a critically important part of any culture study, your instructor may wish to have you prepare your own culture medium and culture dishes. Preparation techniques are described in Steps 1–3 below. If your instructor provides culture dishes made up in advance, skip to Section B. 1. You can make a simple culture chamber by placing the inverted cover of a 60-mm petri dish in a standard petri dish (fig. 11.1).Add a small amount of water to the outer dish so that the small inverted cover has water around it but is not floating freely on the water’s surface. If you are using semisterile techniques, these vessels, the saline solution to be used in medium preparation, and all glassware should be sterilized in an autoclave, but sterilization is not absolutely essential, as reasonably good results can be obtained by maintaining scrupulous cleanliness. 2. The two major components of the medium (solution A and solution B) are prepared in separate steps and mixed just before the medium is poured into the culture dishes. a. Preparation of solution A. Combine 50 ml of Howard Ringer’s solution (see Appendix D) with the white of one egg in a capped container. Shake the mixture vigorously and set it aside in a cool place until solution B, the saline-agar-glucose component, has been prepared. b. Preparation of solution B. Add 0.4 gm of plain agar (not nutrient agar) to 70 ml of Howard Ringer’s solution in a flask and heat the mixture to boiling, stirring continuously. Cool this mixture after it has come to a boil, and as it cools, stir in 1 gm of glucose. When this mixture has cooled to about 45° C, you are ready to combine the two components and pour the medium into the culture vessels. 3. Add 30 ml of solution A (saline-albumin mixture) to solution B (saline-agar-glucose mixture). Pour solution A carefully so that the frothy material on the surface is excluded from the final mixture. Swirl the combined medium to mix the components thoroughly. 4. Proceed at once to pouring the medium into the culture dishes because the agar will begin to solidify rather quickly with further cooling. Pour enough medium to cover the bottom of each inverted cover. Check with your instructor if you need help with proper medium pouring technique. Culture dishes can be stored in a refrigerator if necessary. If culture dishes have been refrigerated, however, they should be allowed to warm to room temperature before use.

Explantation of Embryos 1. When you have culture vessels ready, proceed with embryo explantation. Remove a blastoderm from a fertile egg incubated for 24 to 30 hours, using the same technique that you used earlier in the examination of living embryos (see Laboratory 10, “Patterns of Chick Development”), and place it in a Syracuse dish. Use warm Howard Ringer’s solution for all steps requiring saline in the procedure described in Laboratory 10. After you have transferred the blastoderm to a Syracuse dish, you may have to spend some time carefully removing yolk granules and freeing the blastoderm from the vitelline membrane. Try to work efficiently, however, because it is best to avoid extensive handling of the blastoderm. Some people who culture embryos report that they obtain better growth if they trim the blastoderm and discard the portion anterior to the embryo’s head. If you decide to do so, this trimming can be done with a pair of steel needles (clean dissecting needles are adequate for this). Cross two needles over the blastoderm with the tips kept on the bottom of the dish and draw the needles across one another to produce a scissors action that should cut the blastoderm cleanly. Do not trim other sides of the blastoderm too closely, however, because you might obtain poor results if you do not include part of the area opaca in the explant.

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surgical cut

fusion area



(a)



(b)

FIGURE 11.2 Heart duplication surgery. (a) View of the ventral side of a chick embryo showing the fusion area where the two heart rudiments are coming together on the midline as body folding proceeds. A small cut needs to be made in this “shelf” of tissue. (b) An enlargement showing tissue separation in the fusion area after it’s been surgically cut. A small nick in the fusion area usually is adequate to prevent further rudiment fusion.


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2. Make certain that you keep each petri dish open only as long as is absolutely essential for transfer and positioning of the blastoderm. Flush your “chick pipette” with clean saline and use it to transfer the blastoderm onto the culture medium. Gently spread the blastoderm with your forceps. It is important to leave a little saline around the blastoderm, but you may need to remove some excess saline in order to flatten the blastoderm. Carefully draw off this excess saline with a Pasteur pipette. 3. Mark the culture vessels so that you can identify each of your cultures. Make a rough sketch of the embryo in each culture. Record notes on such features as somite number, head folding, neural development, and any other features that you observe. Note whether the dorsal or ventral surface of the blastoderm is in contact with the surface of the medium in each case. If you have several culture dishes available and find that you are becoming adept at handling blastoderms, you might wish to cut up at least one blastoderm and culture the fragments. 4. Place the culture vessels in the incubator.

Analysis of Results 1. Observe your cultures after 24 hours in the incubator. Make rough sketches of each embryo and determine developmental stages as accurately as you can. Report results using the Hamburger-Hamilton stages, or morphological criteria including somite number, along with information about hours or days of incubation. Compare the rate of development in vitro with the rate in the egg (in ovo). Your instructor may be able to provide some eggs that were not used for cultures and have been incubated since the time when the cultures were prepared. You might find it helpful in assessing relative developmental rates to open some of these eggs and compare the embryos they contain with your cultured embryos. If it is possible for you to continue your cultures for another day, return the cultured embryos to the incubator as quickly as possible for further incubation. Discard cultures in which the embryos obviously have failed to develop or that show signs of gross contamination (milky, white fluid is evidence of bacterial contamination). However, be cautious about discarding any uncontaminated cultures even though they do not appear to contain well-developed embryos because even embryo fragments or grossly abnormal development can produce interesting results. For example, look carefully for any sign of heart development such as a misshapen, but nevertheless pulsating, heart tube. 2. If possible observe your cultures at the end of a second day of incubation and record observations in the same way that you did at the end of the first day. By this time, you may expect contamination of most cultures, particularly if only minimal precautions were taken to prevent infection.

HEART DUPLICATION EXPERIMENTS Techniques Surgical Procedure By standard techniques (see Laboratory 10), isolate an embryo that has been incubated 22 to 25 or 26 hours. Remove the vitelline membrane if necessary and turn the embryo ventral side up. Locate the anterior intestinal portal and “shelf” of tissue where heart tube rudiments are fusing (fig. 11.2a).You probably will not be able to see the heart rudiments themselves, but you can do the necessary microsurgery without actually seeing the rudiments. Cross a pair of microsurgical needles and use them in a scissorslike motion to make a small cut in the fusion area. If you succeed in making the cut, you will be able to see a small notch in the fusion area (fig. 11.2b). Once you have made the cut, transfer the embryo to a culture vessel as described previously and spread the blastoderm out on the culture medium surface. Results will be about the same whether the embryo lies dorsal or ventral side up on the culture medium surface so there is no need to attempt to turn the blastoderm over.Your instructor will let you know how many embryos and culture dishes are available for your use. If necessary, you can place two blastoderms on the culture medium surface in a single petri dish.

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Analysis of Results Observe the operated embryos after the cultures have been in the incubator for 24 hours. Look for evidence of beating hearts.You will probably need to examine each embryo very carefully as heart tubes may lie in unexpected orientations because of distortions caused by the sometime contorted positioning of the blastoderm on the culture medium surface. Since the embryonic heart’s functioning is very temperature dependent, it is important to observe each culture very soon after it has been removed from the incubator and before the embryo has cooled appreciably. If you think that your embryo has one or two heart tubes that are not beating, it is sometimes possible to restart beating for a short time by touching the heart with a metal probe or forceps tip. Apparently, the metal surfaces sometimes provide enough electrical charge to restart the heart. If there are two beating hearts in an embryo, do they initiate their beats in unison? What happens to the beats when the contractile waves reach the anterior, undivided portion of the heart? Can you think of an experiment to investigate the temperature dependence of the embryonic heartbeat? Materials EQUIPMENT Erlenmeyer flasks (for medium preparation) Standard 100-mm (or 90-mm) petri dishes 60-mm petri plates Finger bowls Syracuse dishes Scissors Watchmaker’s forceps “Chick pipette” Pasteur pipettes Wooden-handled probes (dissecting needles) Tungsten or cactus spine microsurgical needles Instrument jar Dissecting microscope Illuminator SOLUTIONS

AND

CHEMICALS

Howard Ringer’s solution Unincubated eggs Agar (plain agar, not nutrient agar) Glucose LIVING MATERIAL Eggs incubated for 24 to 30 hours (for basic in vitro culture technique) Eggs incubated for 22 to 25 or 26 hours (for heart rudiment experiments)

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12. Surgery on the Chick Embryo


L A B O R A T O R Y

12 Surgery on the Chick Embryo

Surgical techniques have been used to examine many different developmental processes in the chick embryo. Numerous experiments done at various stages of development have tested the effects of altering normal structural relationships among parts of developing embryos. In other experiments, development in the absence of various parts has been studied following their surgical removal (extirpation). Surgical procedures used on chick embryos are initially somewhat difficult for inexperienced workers, but most students enjoy attempting chick embryo surgery because of its fundamental and historical importance as an experimental method in developmental biology. This laboratory provides an introduction to chick embryo surgery, and the two surgical procedures suggested here are ones that can give inexperienced workers a modest return in positive results. The first of these procedures is the removal of part of the developing limb bud of the 3-day embryo. Students generally find it difficult to precisely control the portion of the bud removed, but it is possible to produce a morphologically deficient, or abnormal, limb by removing a substantial part of the limb bud. Some of your results will be similar to those obtained in the classical surgical experiments done on the developing limb (see “Suggestions for Further Investigations” on p. 140). It is also possible that some of the abnormalities produced may resemble certain genetically caused limb abnormalities. Modern experiments involving surgical manipulations of developing chick limbs continue to yield information about details of vertebrate limb development.They also contribute to understanding of developmental pattern formation in which adjacent groups of cells differentiate in strikingly divergent ways.For example,data from experiments on chick limbs have contributed substantially to understanding the roles of localized specific gene expression and diffusible morphogens in determining differentiation of complex patterns in many developing systems (see “Suggestions for Further Investigation” section). The second, and more difficult, of the surgical manipulations suggested in this laboratory is the disruption of the normally closed neural tube of the 3-day chick embryo in the posterior spinal cord region. Because the normal development of the vertebral column is dependent upon the structural integrity of the spinal cord, surgical reopening of the neural tube at this stage of development can lead to the condition known as spina bifida. This syndrome ranges in severity from minor malformations of the vertebrae to such severe abnormalities as the protuberance of the virtually unprotected spinal cord at the surface of the back. Spina bifida occasionally occurs as a human birth defect.

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(a) Cutting window in egg

Cuts in shell

(b) Removal of limb bud

Removal of flap

Exposure of embryo

(c) Spina bifida production

FIGURE 12.1 (a) Steps in opening a “window” in the eggshell. (b) Removal of a portion of the limb bud of a 3-day chick embryo. (c) Disruption of a posterior portion of the neural tube of a 3-day chick embryo.


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Techniques General Surgical Procedures Steps 1–6 below are general ones that apply to either surgical technique. Sections B and C describe the two surgical procedures. Before you start, a brief review of the structural relationships of the embryo, the extraembryonic membranes, and the egg would help you in doing the work in this laboratory. 1. Wash the table surface in your work area. Organize your equipment and instruments around your dissecting microscope. Be careful to keep your instrument jar away from your alcohol lamp in order to avoid ignition of the alcohol in the jar. Whenever you remove instruments from the jar, flame off the alcohol by igniting it over the alcohol lamp to complete sterilization. Avoid tipping instruments up in such a way that flaming alcohol runs back onto your fingers. It is difficult to see the clear blue flame of burning alcohol, and thus, it is possible to receive a painful burn. Be very careful not to transfer a flaming instrument to the instrument jar by accident! 2. You will need to prepare an approximately 1-cm2 “window” in the egg to gain access to the embryo. Set the egg in a styrofoam egg nest and select an area of the shell that is slightly nearer the blunt end of the egg than to the other end. Swab the shell area and your fingers with a piece of cotton dampened (not soaked!) with alcohol from your instrument jar. After the shell has dried, begin to cut into it with a sharpened hacksaw blade that has been flame sterilized. During this sawing, hold the fingernail of your index finger against the edge of the blade as a guide to keep the cutting edge from sliding around on the surface of the shell.The cut is deep enough when the blade feels as if it is “catching” rather than sawing smoothly. This process should be repeated twice more in order to leave cuts on three sides of a square “window” (fig. 12.1a). 3. Flame sterilize a single-edged razor blade and slip it under the cut opposite the uncut side of the square. A gentle lifting motion should flip back a small piece of shell that can be discarded. If you experience difficulty in lifting the shell, your hacksaw cuts may have to be deepened. Note: Many people find it more efficient to cut the eggshell using a powered cutter such as a Dremel Tool with a flat cutting wheel. If you choose to use this method, sterilize the shell surface as previously described and cut all four sides of the window.Then remove the square of shell with a sterilized forceps and proceed with any necessary shell membrane removal. 4. When you remove this piece of shell, the shell membranes will be revealed. Do not be misled by the use of the plural “membranes,” as the shell membranes are closely applied to one another and can be removed together in a single step. Squirt a small volume of sterile saline solution on the membranes to wash away small shell remnants that may be left on their surface. It is also helpful to rock the egg back and forth to free any minor adhesions of embryonic membranes to the shell membranes. Peel the shell membranes off with your watchmaker’s forceps. In order to prevent damage to underlying structures, puncture the shell membranes with forceps held at an oblique angle and make certain that penetration is only to the minimal depth necessary to grasp the membranes. The membranes usually can be removed as a series of small strips. If you lose hold of a strip and drop it on the underlying embryo or membranes, retrieve it carefully, avoiding injury to the delicate structures below.You may need to add a drop of saline if this happens, because strips of shell membranes tend to stick to embryonic membranes and sometimes must be floated free. 5. Adjust your light source to give the best possible illumination of the embryo.You may detect a reflection of light off the vitelline membrane over the body of the embryo. This membrane is easily punctured with a microsurgical needle, so you needn’t worry if you are not certain that you actually see it. The surgical procedures that follow require the use of a microsurgical needle. If you are using a tungsten needle, do not risk damage to the point of the needle by dipping it in the instrument jar.A brief excursion through the flame of an alcohol lamp is adequate to sterilize it. If you are using a cactus spine needle, briefly dip it in alcohol and let it dry in the air. Do not flame a cactus spine needle; it will burn. Contact with the shell edges around your “window” also may break off the tip of your needle. 6. When you have completed your surgery, cover the hole with one or two strips of cellophane tape and return the egg in its nest to the incubator. Make certain that the egg is well sealed with tape because drying can cause problems. It sometimes takes several pieces of tape to seal an opening.

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Extirpation of the Limb Bud Cut away a portion of the right leg bud with either your microsurgical needle or your watchmaker’s forceps (fig. 12.1b). You may find it necessary to use both instruments to hold the bud steady and cut it.There is a tendency to be too conservative in an operation such as this, so try to take out a substantial piece of the bud.The piece must be cut away cleanly because if it is left partially attached, the great healing powers of the embryo can come into play and produce an essentially normal limb. Close the egg as quickly as you can after finishing your work. Record the results of the operation in a simple sketch of the limb bud made after surgery. You should make notes on the operation so that you have a reference later when you examine the results or in case of embryo death. Each egg should be marked or numbered so that you can identify it specifically. Carefully avoid puncturing the yolk sac during the limb-bud operation because yolk-sac puncture usually causes death of the embryo. You may wish to use some of your embryos for wing-bud, rather than leg-bud, operations. Disruption of the Neural Tube Spina bifida can be induced experimentally by cutting open a posterior section of the neural tube of the 3-day chick embryo.When the tube is cut open at this stage, reclosing does not occur, and there is a permanent disruption of normal structural relationships.You must use great care in this operation because the dorsal aorta lies only a short distance beneath the neural tube and puncture of the aorta will kill the embryo.You may expect some bleeding from smaller vessels, but if you see large spurts of blood in the area of the needle tip, you should discard the egg and begin again with another one. Use a very fine microsurgical needle that has its tip bent, or is attached at an angle to its handle, to cut open a small section of the neural tube at about the level of the leg buds (fig. 12.1c).Attempt to enter the tube from a posterior direction. If your needle tip actually enters the neural canal, it is easy to lay open the tube with a lifting movement. At first, however, you may have to do some rather extensive probing of the neural tube in this area, and you may cut the tube open largely by chance. If the tube has been cut successfully, the lateral walls of the tube flatten and its outlines in that area become indistinct. You may notice only the seeming disappearance of the lateral walls of the tube. Continue work on the tube until you think that you have seen some change. Record notes on each operation that can be compared with the results observed when the egg is opened. Include observations on bleeding and other problems that you encounter. Carefully mark eggs so that you will be able to relate survival and results with observations made at the time of the operation. Analysis of Results 1. During a period of further incubation, your instructor may ask you to candle your eggs and make decisions about removing eggs from the incubator and discarding them.You may expect a number of embryos to die following operations, but be conservative in discarding eggs so that you avoid throwing away a good specimen. Open the eggs you discard, and if possible, determine and record the approximate stage of development at which they died.Your instructor may elect to candle the eggs with you or for you. 2. Results of limb-bud operations may be taken after either 1 or 2 weeks of further incubation. Check the operated limbs for any growth deficiencies, structural anomalies, or other evidence of abnormal development.The limb on the opposite side of the body is a good reference for comparison. Record your observations and, in your notes or discussion, attempt to correlate your results with the types of operations that you performed. It is sometimes possible to relate a specific deficiency in the limb to removal of a specific part of the bud. Compare results with those obtained by your classmates. Your instructor may direct you to preserve the embryos and make dissections or other types of observations on the operated limbs. 3. The embryos with disrupted neural tubes also may be opened after either 1 or 2 weeks of further incubation.After 2 weeks of further incubation, it is usually necessary to pluck some feathers from the back to detect a spina bifida.You may wish to preserve your specimens in alcohol or formalin for later observation. If time permits, a dissection of preserved specimens may help to clarify the full extent of any deformity of the spinal cord and surrounding tissues. In your notes or discussion, relate results obtained to observations made during the operations.

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Materials EQUIPMENT Styrofoam “egg nests” Illuminator Dissecting microscope Sharpened hacksaw blade Single-edged razor blade (or Dremel Tool with flat cutting wheel) Clean Pasteur pipettes Instrument jar Alcohol lamp Tungsten (or cactus spine or glass) microsurgical needles Cellophane tape SOLUTIONS

AND

CHEMICALS

Howard Ringer’s solution LIVING MATERIAL Eggs incubated 75–82 hours (or slightly longer, see Appendix D)

Useful Information Source http://sdb.bio.purdue.edu/dbcinema/ledouarin/ledouarin07qt.html This is a video sequence from the laboratory of Nicole LeDourain showing the steps in a delicate microsurgical procedure in which part of a quail neural tube is substituted for a region of a chick embryo’s neural tube.

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13. Chorioallantoic Membrane Grafting


L A B O R A T O R Y

13 Chorioallantoic Membrane Grafting

Transplantation to the chorioallantoic membrane (CAM) of the developing chick embryo provides a means of isolating embryonic, or other, tissues from their normal environment while still permitting further growth and differentiation.This extraembryonic site provides a very favorable environment for development of the tissue being studied, and the complex preparations required for in vitro culture techniques are unnecessary.The extraembryonic fluids make a suitable “medium,” and if the graft “takes” effectively, blood vessels of the host’s extraembryonic circulation grow into the graft and vascularize it. These vessels then transport nutrients and oxygen to the graft and carry metabolic wastes away from it. This technique has somewhat different applications than in vitro culture methods because, in CAM grafting, the materials and environment provided for the isolated tissue cannot be precisely defined and controlled. For instance, increasing quantities of hormones in the embryonic circulation reach a graft during the period that it is in place on the CAM. The CAM grafting technique, however, is a relatively simple way to study tissues in isolation from their normal environment. Although it is possible to grow tissues taken from many different animals on the chorioallantoic membrane, the most extensive use of this technique has been in analysis of chick embryo development itself. For example, throughout a long and very productive career, Mary Rawles employed CAM grafting to investigate various aspects of chick development, including determination, in early embryos (see “Suggestions for Further Investigation” section on p. 142). In one extensive set of experiments, she divided blastoderms into carefully defined parts and grew the pieces as CAM grafts. She then sectioned the grafts and examined them histologically in order to determine what tissue types could be detected in grafts derived from each part of the blastoderm. By careful and repeated replications of these experiments, she was able to catalogue determination of the various parts of the blastoderm in terms of the developmental potentials of these isolated pieces. Later, Rawles took skin from several different regions, separated mesodermal (dermis) and ectodermal (epidermis) components, recombined them in artificial relationships, and transplanted these skin fragments to the CAM where she could follow their subsequent development.Through these experiments, she contributed to understanding of the control mechanisms involved in skin differentiation. This laboratory will give you an opportunity to work with this interesting and historically important technique.

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(a) External view of egg using candler



air space

embryo “Y” region with rich blood supply

(b) Placing of graft through window

FIGURE 13.1 (a) Marking a Y-shaped junction of CAM blood vessels located by candling. (b) Transferring a graft to the CAM.

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Techniques Preparation of Hosts 1. Eggs that have been incubated for 9 or 10 days will serve as hosts. Candle the eggs to locate and mark an area where you can see a Y-shaped junction of blood vessels in the CAM (see fig. 13.1a and “Candling” in Appendix D). Various investigators who used the CAM grafting technique have suggested that such junctions are the best sites for grafts. 2. After marking the eggs, place a “window” in the shells of several host embryos. The technique for opening the shell is basically the same as that described on page 131 in Laboratory 12 (“Surgery on the Chick Embryo”). However, the technique should be slightly modified for CAM grafting. Before you begin sawing the shell, puncture the air space at the egg’s blunt end.This causes the CAM to drop away (“lower”) beneath the window when it is opened, and this lowering helps to prevent membrane damage during subsequent manipulations. After you have swabbed the blunt end of the egg with alcohol, stick the needle of a wooden-handled probe about 1/2 cm into the egg’s blunt end. Carefully control the probe during this step because it takes some pressure to puncture the shell, but be cautious, since it is easy to drive the needle right on through the air space into the interior of the egg. Proceed with window cutting after you have punctured the air space.When the window is complete, loosely cover it and the hole you have punched in the egg’s blunt end with pieces of cellophane tape until the donor material has been prepared. Do not press the tape down around the window yet. Return the host eggs to the incubator while you prepare donor material.

Preparation of Donor Tissue 1. Donor tissue can be taken from chick embryos of a variety of ages. Parts of early embryos sometimes give more spectacular results because of the marked degree of change that occurs during the culture period. On the other hand, tissues from older embryos are much easier to dissect and transfer to the CAM. Some tissues that give good results are eyes or posterior portions of “48-hour” embryos and limb buds of “72-hour” embryos. Back tissue from older embryos (5 or 6 days of incubation) will also develop well and should produce considerable skin growth and numerous feathers. 2. Donor embryos can be cut up in a Syracuse dish containing Howard Ringer’s solution. Cross dissecting needles over the body of the donor. Keep the tips against the bottom of the dish and draw the needles across one another in a scissorlike fashion. After one or two trials, you should become adept enough at slicing up the body of the donor to prepare the specific pieces that you wish to use. Sometimes cautious use of a microsurgical needle makes precise separation of embryonic parts easier. One donor can be used to provide tissue for several grafts.

Transplantation 1. Reopen a host egg when you are ready to transplant a piece of tissue. Focus your light source on the window so that you can see the blood vessels of the CAM clearly.Transfer the tissue with a watchmaker’s forceps and put it in place over the blood vessel junction that you previously located (fig. 13.1b). You may use a fine-mouth pipette to transfer the tissue, but the saline carried along in the pipette tends to float the tissue off the selected position. If the graft slips out of sight to the side or simply seems to disappear, transplant a second piece of tissue to the site. Such lost tissue pieces rarely show up later and should be replaced. Once the graft is in place, retape the shell as quickly as possible to avoid further desiccation, pressing the tape down around the window, and return the egg to the incubator. 2. Number your host eggs and record the size and original source of the graft transplanted to each one. Make note of any unusual observations or any problems such as bleeding that you encountered during the procedure. Also record the position of the graft relative to blood vessels.

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Analysis of Results 1. After 7 days of further incubation, open the eggs and recover the grafts. It is best to remove the cellophane tape and enlarge the window by picking away the shell piece by piece.As you remove small pieces of shell, watch carefully for evidence of the growing graft on the CAM. Successful grafts usually are easy to detect as masses of tissue on the membrane. If the graft has not “taken” and has degenerated, you usually will see a small reddish brown or black spot. A few grafts will be lost, and you will find no trace of them, but you should not discard the host until you have examined all parts of the CAM carefully. Remove each growing graft by cutting it out of the CAM. Include a large enough circle of CAM tissue to permit handling without touching the graft itself. Get directions from your instructor regarding disposition of host embryos. 2. Carefully examine the grafts under the dissecting microscope. Sketch each graft, measure its size, and identify as many structures as possible. In your notes or discussion, compare development of the tissue in the graft with the development of the comparable part of an intact embryo over the same period of time. Some extra donor embryos may have been left in the incubator for purposes of comparison. 3. If your instructor so directs, preserve each graft and put it in a small, carefully labeled container.

Materials EQUIPMENT Egg candler Styrofoam “egg nests” Wooden-handled probes (dissecting needles) Microsurgical needle Sharpened hacksaw blade (or Dremel Tool with flat cutting wheel) Single-edged razor blade Watchmaker’s forceps Dissecting forceps (for shell removal) Scissors Instrument jar Alcohol lamp Absorbent cotton Cellophane tape Finger bowls Syracuse dish Dissecting microscope Illuminator SOLUTIONS

AND

CHEMICALS

Howard Ringer’s solution LIVING MATERIAL Eggs incubated 9 or 10 days (hosts) Donor embryos (see page 137 for suggested incubation stages)

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Suggestions for Further Investigation of Chick Embryo Development


SUGGESTIONS FOR FURTHER INVESTIGATION OF CHICK EMBRYO DEVELOPMENT The history of chick embryo development study stretches back to Aristotle, making the chick embryo one of the longest studied examples of developing animals. This research on the chick embryo has contributed a great deal to our understanding of vertebrate development.This is especially true of those vertebrates whose development goes through a flat, several-layered disk phase (reptiles, birds, and mammals). There is much more yet to be learned through study of the chick embryo, and a few of the currently active areas of investigation of chick development are discussed in the following. References are provided so that if you wish, you can further pursue some of the aspects of chick development that you have studied in the labs in this course. Chick Development Patterns The status of investigations of chick embryo development has been summarized periodically over the past several decades (for example, Romanoff 1960; Freeman and Vince 1974; Metcalfe et al 1987; Stern 1994; Schoenwolf 1997; Bellairs and Osmond 1998). Each of those sources contains a treasure of information about chick development, and they are good starting points if you want more information about some aspect of chick development.There also have been several useful compilations of methods ( Bellairs 1993a; Bronner-Fraser 1996; Mason 1999). However, it would not be good to give the impression that our understanding of chick embryo development is now static or complete or that there is universal agreement concerning descriptions and interpretations of the processes involved. Even very general matters such as incubation conditions (Deeming 1989;Turner 1990) and the nature of the hatching process (Bond et al 1988; Aldhous 1995) are still topics for active and interesting research. Like all vigorous fields of science, the study of chick embryo development has progressed through a number of periods of controversy,and interpretations of certain developmental processes have changed substantially over time. For example, in the 1960s, Spratt and Haas (1965) challenged the long-standing, traditional view that the primitive streak functions mainly as a route for cell migration.They contended that the primitive streak is a proliferation center, functioning much like a regeneration blastema in contributing numerous newly divided cells. They argued that the streak is a mitotically active center that generates the cells that then migrate away to take their places, for example, as embryonic mesoderm. Spratt and Haas downplayed the importance of migration of epiblast cells toward, down into, and away from the primitive streak. Their contention inspired other research, and Rosenquist (1966), using tritiated thymidine labeling, thoroughly reinvestigated the role of the primitive streak. His results strongly supported the view that the primitive streak is indeed an active migration center.You might find these historical papers interesting to read as an example of the role of scientific controversy in clarifying understanding of even familiar and frequently observed biological processes, such as gastrulation in the chick embryo. Rosenquist’s work actually went well beyond response to the points raised by Spratt and Haas because it helped to establish the contribution of the epiblast to development of the embryonic endoderm, which had previously been thought to be a derivative of the hypoblast.The fate map of the early chick embryo was reshaped and descriptions of gastrulation processes were modified to incorporate this information ( Nicolet 1971; Eyal-Giladi and Kochav 1976).These were, by no means, the final words on early chick development. Study of very early cell interactions and cell movements continues as an active research area in developmental biology (Mitrani and Eyal-Giladi 1981; Khaner and Eyal-Giladi 1989; Schoenwolf and Sheard 1990; Stern 1990; Stern and Canning 1990; Selleck and Stern 1991; Eyal-Gilad et al 1992; Bellairs 1993b), as does research on gastrulation processes (Bellairs 1986; Stern and Canning 1988; Sanders 1991; Nieto et al 1994; Hatada and Stern 1994; Schoenwolf and Yuan 1995; Lemaire and Kessel 1997).The powerful tools of modern cell biology research can be applied to investigation of developmental processes such as details of neural tube induction (Dodd et al 1998;Vogel 1998) and rightleft asymmetry in the embryo ( Robertson 1997; Isaac et al 1997; Capedevila 2000; Garcia-Castro et al 2000).

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Research on events occurring as development proceeds,such as neurulation (Schoenwolf and Smith 1990; Alvarez and Schoenwolf 1991) and skeletal tissue development (Hall 1988) continues to be focused on chick embryos. Clearly, the chick embryo has both a substantial history and a promising future as a subject for investigations of developmental patterns and processes ( Landmesser 1992). In Vitro Culture Further information on embryo culturing by the Spratt technique can be obtained in Rugh (1962). DeHaan (1967) describes avian embryo culture techniques and provides references to some studies that have used culture techniques to approach various developmental problems. New (1966) compiled a broader treatment of the culture of various vertebrate embryos. Another interesting and highly useful technique for embryo culture was developed by New ( New 1955, 1966; DeHaan 1967; Hornbruch 1999).This technique involves transfer of the vitelline membrane along with the blastoderm to a liquid culture. With the New technique, it is possible to avoid some of the distortion of embryos that frequently is caused by semisolid culture media. You may be interested in comparing results obtained using the Spratt and New techniques. New’s technique has been simplified (Downie 1979) and modified to produce better, more normal development for a longer period of time (Flamme et al 1991). Each of these embryo culture technique modifications offers interesting possibilities that you might want to explore. Rosenquist (1966) used in vitro culture techniques to study cell migration through the primitive streak and autoradiographic labeling of embryos cultured by the New technique to determine origins of the cell populations that eventually produce various organs in the embryo (for example, Rosenquist 1971). The New culture technique has been employed in investigating various aspects of early development ( Lash et al 1990; England and Lawson 1993). In some investigations of specific cellular interactions involving growth factors, use of a chemically defined culture medium is essential, and Mitrani and Shimoni (1990) devised such a medium. One of the most exciting advances in the area of culture of chick embryos has been development of techniques for continuous culture of the embryo outside the normally enclosing eggshell.The chick embryo can be cultured from fertilization to hatching ( Rowlett and Simkiss 1985, 1987; Perry 1988; Naito et al 1990).This achievement is particularly important for possible applications in biotechnology. Transgenic chicks can be produced only if there is access to embryos at very early stages of development and an available means of supporting subsequent development to the hatching stage ( Naito et al 1995; Simkiss 1993, 1997). In vitro embryo culturing holds a certain fascination, and you might like to try your hand at chick embryo culturing using techniques that are simpler than any of those mentioned so far. It is quite easy to culture embryos for at least part of their development in petri dishes (Auerbach et al 1974; Scadding 1981) or even in plastic coffee cups ( Jakobson et al 1989)! The roots of embryo culture techniques can be traced to techniques developed earlier in tissue and organ culture research (Wessels 1967).You might wish to undertake an organ culture experiment, and Powell and Pitkin’s (1981) methods for organ culture of chick embryo hearts might give you a good starting point for an investigation that you could extend in several ways. In considering culture techniques of various types, it is important to remember that each is built on foundations of earlier research. For example, the physiological saline solution used in many applications with chick embryos, Howard Ringer’s solution, was developed by Evelyn Howard (1953) through a long, painstaking series of experiments. Virtually all experiments, in all areas of science, utilize information gained from previous investigations, and it is important when doing chick embryo culture experiments to recognize and appreciate the contributions of earlier workers such as Evelyn Howard, Nelson Spratt, and Dennis New. Surgery The chick embryo will tolerate and survive an impressive variety of surgical procedures (Hamburger 1960; Rugh 1962). It probably would be impossible to compile even a representative list of experiments involving chick embryo surgery that have been done, so only a few will be considered here.

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In the 1960s, James Weston combined surgical and autoradiographic techniques to study the migration of neural crest cells, which, among other developmental roles, become ancestors of the pigment cells. Weston (1963, 1970) substituted radioactively labeled sections of neural tube for pieces of neural tube that he had surgically removed from embryos in early stages of development. These substituted pieces healed into place and developed in harmony with the host tissue adjacent to them. Weston sacrificed these host embryos at intervals after the operation and used autoradiographic analysis to trace migration of labeled neural crest cells away from the grafted section of the neural tube.You can obtain further details of these technically difficult and cleverly executed experiments in Weston’s papers. Nicole LeDourain has done remarkable surgical transfers of large parts of the nervous system from quail to chick embryos (Barnes 1986, 1988;Teillet et al 1999). Some of the greatest triumphs of chick embryo surgery have come in studies of limb morphogenesis. Limb-bud transplantation has a long history (for example, Hamburger 1938; Murray and Wilson 1994), and Mary Rawles used limb-bud transplants in her classic studies of color pattern development. Studies of the proximodistal developmental sequence of establishment of limb elements (Saunders 1948, 1980; Zwilling 1961) included such delicate surgical manipulations as removal and reattachment of the apical ectodermal ridge (sometimes called Saunder’s ridge). Later studies led to the concept of a progress zone of mesoderm underlying ectoderm at the limb-bud tip that is responsible for originating the proximodistal pattern of the developing limb (Summerbell and Lewis 1975;Wolpert 1991). Surgical techniques have also been used to determine that experimentally applied sources of peptide growth factors, fibroblast growth factors (FGFs), can substitute for the apical ectodermal ridge in maintaining the proximal-distal progression in establishment of limb elements after the ridge has been surgically removed (Fallon et al 1994). FGFs can also induce regeneration of surgically removed portions of chick embryo limb buds (Taylor et al 1994; Kostakopoulou 1997).This discovery may have exciting implications for future research on vertebrate limb regeneration. FGF-soaked beads applied to the embryo’s flank can cause the growth of additional limbs between the normally developing ones (Kohn et al 1995). But a limb has other axes of polarity in addition to the proximodistal axis, and surgical methods were used in elucidating the location and role of the zone of polarizing activity, or ZPA (for example, Honig and Summerbell 1985; Hinchliffe and Samson 1985; Wolpert 1998), in establishing those axes. Surgical techniques were also used to demonstrate that retinoic acid-impregnated pellets would mimic the action of the ZPA and artificially induce anterioposterior duplication of digits (Tickle et al 1982; Summerbell 1983; Eichele 1986; Smith et al 1989). Despite this evidence, questions remain about the possible roles of endogenous retinoic acid in limb pattern development (Tickle 1991; Brockes 1991; Tabin 1991).The Sonic hedgehog gene appears to be involved in ZPA activity (Riddle et al 1993; Riddle and Tabin 1997), though it is not yet clear whether the product of Sonic hedgehog expression acts as a morphogen or whether expression of this gene might be a sequential step leading to morphogen production (Rennie 1994; O’Farrell 1994). Morphogenetic cell death is another interesting aspect of limb development that has been explored through surgical experiments. There are populations of embryonic cells whose normal developmental fate is to die and degenerate. Such large-scale cell death occurs in the shaping of limbs and their terminal digits (Saunders et al 1962; Saunders 1966; Saunders and Fallon 1967; Vaux et al 1994). If you should wish to experiment further with surgery on chick embryos, some technique modifications have been suggested by Bagnall (1988). Experiments on wound healing in the embryo (see Thevenet and Sengel 1986) might be somewhat less traumatic to embryos than more drastic experiments that you might attempt. If you wish to attempt a major surgical manipulation that yields very interesting results, you might undertake experiments on endocrine relationships during chick development. It is possible to remove the rudiments of the developing pituitary gland by cutting off the forepart of the head after about 1 1Ⲑ2 days of incubation.This pituitary removal (hypophysectomy) by partial decapitation has drastic effects on a variety of structural and physiological aspects of development and has been done in a number of studies on endocrine regulation of developmental processes (Hinni and Watterson 1963;Thommes and Jameson 1980;Thommes 1987).You may wish to attempt this surgical procedure. It is not as difficult as you might think, and you may be able to do the operation if you have had some success with other

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types of surgery.The rather spectacular results of successful operations (stunted embryos lacking eyes and upper beaks) are very interesting because the embryos show multiple symptoms of massive disruption of endocrine gland functioning. Such results underscore the importance of normal hormonal balances in the developing chick embryo (see also Murphy and Clark 1990 under “Chick Development Patterns” in the reference section following). If you do experiments on limb buds or on other aspects of development in which the skeleton might be affected, it is interesting to prepare embryos with the skeleton selectively stained and the rest of the body tissues cleared to make the skeleton visible (Guyer 1953; Jensh and Brent 1966; Hamburger 1960; and see also Carlson et al 1986). In examining results of surgical experiments on the developing nervous system, there is a method for dissecting embryos following pretreatment in nitric acid that makes possible the isolation of a nearly intact central nervous system. These dissections add a great deal to the study of results of operations on the developing nervous system such as your spina bifida operation. For description and application of the technique, consult Huber (1936) and Watterson (1949). Kim et al (1995) did surgical manipulation of the developing chick embryo nervous system at later stages of development. There are still more developmental problems that can be approached surgically (Degennaro 1991), and though much is known about developmental patterns in vertebrate embryos (Wolpert 1992), there is still much more to be learned by further experimental study of chick embryo development. Chorioallantoic Membrane Grafting The richly vascularized chorioallantoic membrane (CAM) serves as the gas exchange surface for the developing embryo. It also mobilizes and transports eggshell calcium, thereby making it available to the developing embryo (Tuan and Ono 1986;Tuan 1987). The CAM has been of interest to developmental biologists, not only for the exchange and transport functions that it conducts for the embryo, but because it provides an excellent site for grafted tissues.When a tissue is placed on the CAM, blood vessels grow into it, and this vascular supply provides nutrients, waste removal, and gas transport for the grafted tissue. Although it has been demonstrated that diverse tissues from various donors—ranging from frogs to humans—can be grown on the chick CAM, the most extensive use of this technique has been in experimental analysis of chick development.The work of Mary Rawles (1936, 1943, 1952) is an excellent example of the kind of results that can be obtained using this technique. She divided early chick embryos into carefully defined parts and grew these pieces as CAM grafts. She sectioned the grafts and examined them histologically in order to determine what tissue types could be detected in grafts derived from each part of the blastoderm. By careful replications of these experiments, she was able to catalogue the developmental potentials of various parts of the blastoderm.You may wish to attempt further experimentation on the ability of parts of very young embryos to carry on independent development and differentiation on the CAM. In experiments on interaction of ectodermal and mesodermal components from various skin areas, Rawles (1963) found that drying caused some abnormal developmental patterns. She modified conventional CAM techniques by a daily addition of fluid (albumen and saline in a 15⬊1 mixture) that kept the graft submerged.This treatment caused the membrane to rise and brought the graft nearer the shell opening. In order to accommodate this rising membrane, she constructed a “chimney” device over the original shell opening. You might find her paper interesting because of the technical modification of CAM grafting reported in it and because it is another excellent example of application of CAM grafting in the experimental analysis of development. Weiss and Taylor (1960) applied CAM grafting to problems of behavior and interaction of individual embryonic cells.They dissociated cells from organs of relatively advanced chick embryos (8 to 14 days of incubation) and centrifuged the cell suspensions into lumps that were transplanted to the CAM. Sectioning of the aggregates after further development on the CAM revealed that there had been reorganization into tissues characteristic of the organs from which the cells had originated. For example, mesonephros aggregates developed apparently normal kidney tubules and skin aggregates produced

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feathers. Thus, CAM grafting was teamed with techniques of cell dissociation to demonstrate that dissociated cells from organs in advanced stages of differentiation still possessed the ability to reaggregate and reorganize into histologically normal tissues. You may wish to compare this work with that of Moscona (see “Suggestions for Further Investigation of Amphibian Development”, p. 95) that concentrated on very early organ rudiments. The work of Weiss and Taylor is of particular interest because it introduced the possibility of using the CAM to replace elaborate and expensive culture arrangements in dissociation experiments. This could make experiments on dissociated cells of embryonic homeotherms possible for interested developmental biology students. The CAM continues to be used as a culture site in experiments involving embryonic chick tissues (Wanek et al 1991; Bertossi et al 1998), as well as tissues from other organisms (Hardy et al 1990; Fuenzalida et al 1990; Gajovic and Gruss 1998). The CAM also serves as a model system for the study of factors controlling angiogenesis (West et al 1985; Burton and Palmer 1992; Flamme et al 1991; Ribatti et al 1996; Sikorski and Peters 1998).This research is of interest in relation to tumor biology because one of the most striking characteristics of neoplastic growths is their ability to stimulate development of a rich blood supply that allows them to thrive at the expense of other body tissues.You might wish to design some experiments on factors affecting CAM angiogenesis responses. Edgar Zwilling (1958) developed a modified CAM grafting technique that can improve results in CAM grafting experiments. You might find it interesting to consult Zwilling’s paper and compare his CAM grafting with the standard technique described in Laboratory 13. If you want to try a different kind of tissue grafting, you may want to experiment with the intracoelomic grafting technique developed by Dossel (1954) and described as well by Coulombre (1967), who also describes several other techniques for grafting embryonic rudiments (also consult Hamburger 1960 and Rugh 1962). Chick Embryo Development References General Sources Bellairs, R. Endpoints in the development of chick embryos. Toxic. in Vitro 7:701–706; 1993a. Bellairs, R.; Osmond, M. Atlas of Chick Development. San Diego: Academic Press; 1998. Bronner-Fraser, M. Methods in Avian Embryology. Methods in Cell Biology. San Diego:Academic Press; 1996:Vol. 51. Deeming, D. C.; Ferguson, M. W. J., eds. Egg Incubation. Its Effects on Embryonic Development in Birds and Reptiles. New York: Cambridge University Press; 1992. Freeman, B. M.; Vince, M. A. Development of the Avian Embryo. New York: Wiley; 1974. Hamburger, V. A Manual of Experimental Embryology. Chicago: University of Chicago Press; 1960. Hamburger, V.; Hamilton, H. L. A series of normal stages in the development of the chick embryo. Jour. Morphol. 88:49–92; 1951. Hamilton, H. L. Lillie’s Development of the Chick. 3d ed. New York: Holt, Rinehart and Winston; 1952. Mason, I. The avian embryo—an overview, and chick embryos—incubation and isolation. In: Sharpe, P. T.; Mason, I., eds. Molecular Embryology Methods and Protocols. Methods in Molecular Biology; Vol. 97. Totowa, NJ: Humana; 1999: 215–224. Metcalfe, J.; Stock, M. K.; Ingerman, R. I., eds. Development of the avian embryo. Jour. Exp. Zool. Supplement 1, 1987. (Also published as a book with the same listed title and editors by Alan R. Liss, New York; 1987.) Needham, J. A History of Embryology. 2d ed. New York: Abelard-Schuman; 1959. New, D. A.T. The Culture of Vertebrate Embryos. New York: Logos/Academic; 1966. Oppenheimer, J. M. Essays in the History of Embryology and Biology. Cambridge: M.I.T. Press; 1967. Patten, B. M. Early embryology of the chick. 5th ed. New York, McGraw-Hill; 1971. Romanoff, A. L. The Avian Embryo. New York: Macmillan; 1960. Rugh, R. Experimental Embryology. 3d ed. Minneapolis: Burgess; 1962. Schoenwolf, G. C. Reptiles and birds. In: Gilbert, S. F.; Raunio, A. M., eds. Embryology: Constructing the Organism. Sunderland, MA: Sinauer; 1997: 437–458. Stern, C. D.; Holland, P. D. H., eds. Essential Developmental Biology: A Practical Approach. Oxford: IRL Press; 1993. Stern, C. D. The chick. In: Bard, J. B. L., ed. Embryos: Color Atlas of Development. London: Wolfe; 1994: 167–182. Wolpert, L. The Triumph of the Embryo. Oxford: Oxford University Press; 1991.

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Reviews and Research Papers CHICK DEVELOPMENT PATTERNS Aldhous, P. Shells give chicks an early break. New Scientist 16 December: 18; 1995. Alvarez, I. S.; Schoenwolf, G. C. Patterns of neurepithelial cell rearrangement during avian neurulation are prior to notochordal inductive interactions. Dev. Biol. 143:78–92; 1991. Bellairs, R. Fertilization and early embryonic development in poultry. Poultry Science 72:874–881; 1993. Bellairs, R.The primitive streak. Anatomy and Embryology 174:1–14; 1986. Bond, G. M.; Board, R. G.; Scott,V. D.An account of the hatching strategies of birds. Biol. Rev. 63:395–415; 1988. Capdevila, J.; Vogan, K. J.;Tabin, C. J.; Izpisua Belmonte, J. C. Mechanisms of left-right determination in vertebrates. Cell 101:9–21; 2000. Deeming, D. C. Characteristics of unturned eggs: critical period, retarded embryonic growth and poor albumen utilisation. British Poultry Science 30:239–250; 1989. Dodd, J.; Jessell,T. J.; Placzek, M.The when and where of floor plate induction. Science 282:1654–1657; 1998. Eyal-Gilad, H.; Debby, A.; Harel, N. The posterior section of the chick’s area pellucida and its involvement in hypoblast and primitive streak formation. Development 116:819–830; 1992. Eyal-Giladi, H.; Kochav, S. From cleavage to primitive streak formation: a complementary normal table and a new look at the first stages of the development of the chick. Dev. Biol. 49:321–337; 1976. Garcia-Castro, M. I.;Vielmetter, E.; Bronner-Fraser, M. N-cadherin, a cell adhesion molecule involved in establishment of embryonic left-right asymmetry. Science 288:1047–1051; 2000. Hall, B. K.The embryonic development of bone. Amer. Sci. 76:174–181; 1988. Hatada, T.; Stern, C. D. A fate map of the epiblast of the early chick embryo. Development 120:2879–2889; 1994. Isaac, A.; Sargent, M. G.; Cooke, J. Control of vertebrate left-right asymmetry by a Snail-related zinc finger gene. Science 275:1301–1304; 1997. Khaner, O.; Eyal-Giladi, H. The chick’s marginal zone and primitive streak formation. I. Coordinative effect of induction and inhibition. Dev. Biol. 134:206–214; 1989. Landmesser, L. T. The avian embryo: an accessible system to study the genesis of complex tissue structure. In: Rossomando, E. F.; Alexander, S., eds. Morphogenesis. New York: Marcel Dekker; 1992. Lemaire, L.; Kessel, M. Gastrulation and homeobox genes in chick embryos. Mech. Devel. 67:3–16; 1997. Mitrani, E.; Eyal-Giladi, H. Hypoblastic cells can form a disk inducing an embryonic axis in chick epiblast. Nature 289:800–802; 1981. Murphy, M. J.; Clark, N. B. The avian embryo as a model for early developmental endocrinology. Jour. Exper. Zool. Suppl. 4:177–180; 1990. Nicolet, G. Avian gastrulation. Advances in Morphogenesis 9:231–262; 1971. Nieto, M. A.; Sargent, M. G.; Wilkinson, D. G.; Cooke, J. Control of cell behavior during vertebrate development by Slug, a zinc finger gene. Science 264:835–839; 1994. Robertson, E. J. Left-right asymmetry. Science 275:1280; 1997. Rosenquist, G. C.A radioautographic study of labeled grafts in the chick blastoderm. Development from primitivestreak stages to stage 12. Carnegie Contribs. Embryol. 38:71–110; 1966. Sanders, E. J. Embryonic cell invasiveness—an in vitro study of chick gastrulation. Jour. Cell Science 98:403–408; 1991. Schoenwolf, G. C.; Sheard, P. Fate mapping the avian epiblast with focal injections of a fluorescent-histochemical marker: ectodermal derivatives. Jour. Exp. Zool. 255:323–339; 1990. Schoenwolf, G. C.; Smith, J. L. Mechanisms of neurulation: traditional viewpoint and recent advances. Development 109:243–270; 1990. Schoenwolf, G. C.;Yuan, S. Experimental analysis of the rearrangement of ectodermal cells during gastrulation and neurulation in avian embryos. Cell Tissue Res. 280:243–251; 1995. Selleck, M. A. J.; Stern, C. D. Fate mapping and cell lineage analysis of Hensen’s node in the chick embryo. Development 112:615–626; 1991. Spratt, N. T., Jr.; Haas, H. Germ layer formation and the role of the primitive streak in the chick. I. Basic architecture and morphogenetic tissue movements. Jour. Exp. Zool. 158:9–38; 1965. Stern,C.D.;Canning,D.R.Gastrulation in birds—a model system for the study of animal morphogenesis.Experientia 44:651–656; 1988. Stern, C. D.; Canning, D. R. Origin of cells giving rise to mesoderm and endoderm in chick embryo. Nature 343:273–275; 1990. Stern, C. D. The marginal zone and its contribution to the hypoblast and primitive streak of the chick embryo. Development 109:667–682; 1990.

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Turner, J. S.The thermal energetics of an incubated chicken egg. Jour. Thermal Biol. 15:211–216; 1990. Vogel, G. Embryo’s organizational chart redrawn. Science 280:1838–1839; 1998. IN VITRO CULTURE Auerbach, R.; Kubai, L.; Knighton, D.; Folkman, J. A simple procedure for the long-term cultivation of chicken embryos. Dev. Biol. 41:391–394; 1974. DeHaan, R. L. Avian embryo culture. In: Wilt, F. H.; Wessells, N. K., eds. Methods in Developmental Biology. New York:Thomas Y. Crowell, 1967:401–412. Downie, J. R. Culturing chick embryos—a simplification of New’s method. Jour. Biol. Ed. 13:114–118; 1979. England, M. A.; Lawson, A. Natural wound formation: endodermal responses to experimental primary neural induction in the chick embryo. Anat. Rec. 236:710–720; 1993. Flamme, I.;Albach, K.; Muller, S.; Christ, B.; Jacob, H. J.Two-phase in vitro culture of explanted chick embryos. Anat. Rec. 229:427–433; 1991. Hornbruch,A. New culture. In: Sharpe, P.T.; Mason, I., eds. Molecular Embryology Methods and Protocols. Methods in Molecular Biology; Vol. 97.Totowa, NJ: Humana; 1999: 225–244. Howard, E. Some effects of NaCl concentration on the development of early chick blastoderms in culture. Jour. Cell. Comp. Physiol. 41:277–284; 1953. Jakobson, A. M.; Hahnenberger, R.; Magnusson, A. A simple method for shell-less cultivation of chick embryos. Pharmacol. & Toxicol. 64:193–195; 1989. Lash, J. W.; Gosfieled, E., III; Ostrovsky, D.; Bellairs, R. Migration of chick blastoderm under the vitelline membrane: the role of fibronectin. Dev. Biol. 139:407–416; 1990. Mitrani, E.; Shimoni, Y. Induction by soluble factors of organized axial structures in chick epiblasts. Science 247:1092–1094; 1990. New, D. A. T. A new technique for the cultivation of the chick embryo in vitro. Jour. Embryol. Exp. Morph. 3:326–331; 1955. Naito, M.; Nirasawa, K.; Oshi,T. An in vitro culture method for chick embryos obtained from the anterior portion of the magnum of oviduct. Brit. Poultry Science 36:161–164; 1995. Naito, M.; Nirasawa, K.; Oishi, T. Development in culture of the chick embryo from fertilized ovum to hatching. Jour. Exp. Zool. 254:322–326; 1990. Perry, M. M. A complete culture system for the chick embryo. Nature 331:70–72; 1988. Powell, J. A.; Pitkin, R. B. Laboratory on organ culture of chick heart embryos. Amer. Biol. Teacher 43:43–44; 1981. Rosenquist, G. C. The location of the pregut endoderm in the chick embryo at the primitive streak stage as determined by autoradiographic mapping. Dev. Biol. 26:323–335; 1971. Rowlett, K.; Simkiss, K.The surrogate egg. New Scientist 15 August: 42–44; 1985. Rowlett, K.; Simkiss, K. Explanted embryo culture: In vitro and in ovo techniques for domestic fowl. British Poultry Science 28:91–101; 1987. Scadding, S. R. How to culture chicken embryos in petri dishes. Amer. Biol. Teacher 43:382–383, 396; 1981. Simkiss, K. Embryo manipulation of the germplasm. Poultry Science 76:1093–1100; 1997. Simkiss, K. Surrogate eggs, chimeric embryos and transgenic birds. Comp. Biochem. Physiol. 104A:411–417; 1993. Spratt, N.T., Jr. Development in vitro of the early chick blastoderm explanted on yolk and albumen extract salineagar medium. Jour. Exp. Zool. 106:345–366; 1947. Wessells, N. K. Avian and mammalian organ culture. In: Wilt, F. H.; Wessells, N. K., eds. Methods in Developmental Biology. New York:Thomas Y. Crowell, 1967:445–456. SURGERY Bagnall, K. M. A method to increase the survival rate of early chick embryos in experiments involving surgical intervention. Teratology 38:75–77; 1988. Barnes, D. M. Bird brain switch leads to new song. Science 241:1434–1435; 1988. Barnes, D. M. Bird chimeras may be models for certain neurological diseases. Science 232:930–932; 1986. Brockes, J. We may not have a morphogen. Nature 350:15; 1991. Carlson, B. M.; Simandl, B. K.; Stocker, K. M.; Connelly,T. G.; Fallon, J. F. A method for combined gross skeletal staining and feulgen staining of embryonic chick tissues. Stain Technology 61:27–31; 1986. Degennaro, L. D. Origin of the avian glycogen body. 1. Effects of tail bud removal in the chick embryo. Growth, Devel. and Aging 55:19–26; 1991. Eichele, G. Retinoids induce duplications in developing vertebrate limbs. BioScience 36:534–540; 1986. Guyer, M. F. Staining the skeletons of cleared embryos. In Animal Micrology. 5th ed. Chicago: University of Chicago Press; 1953:148–149.

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Hamburger,V. Morphogenetic and axial self-differentiation of transplanted limb primordia of 2-day chick embryos. Jour. Exp. Zool. 77:379–400; 1938. Hinchliffe, J. R.; Samsom, A. The distribution of the polarizing zone (ZPA) in the leg bud of the chick embryo. J. Embryol. Exp. Morphol. 86:169–175; 1985. Hinni, J. B.;Watterson, R. L. Modified development of the duodenum of chick embryos hypophysectomized by partial decapitation. Jour. Morphol. 113:381–426; 1963. Honig, L. S.; Summerbell, D. Maps of strength of positional signaling activity in the developing chick wing bud. Jour. Embryol. Exp. Morphol. 87:163–174; 1985. Huber, J. F. Nerve roots and nuclear groups in the spinal cord of the pigeon. Jour. Comp. Neurology 65:43–91; 1936. Jensh, R. P.; Brent, R. L. Rapid schedules for KOH clearing and Alizarin Red S staining of fetal rat bone. Stain Technology 41:179–183; 1966. Kim, K. B.; Cho, B. K.; Chi, J. G.; Wang, K. C. Morphological study of surgically induced open neural tube defect in old (14 and 21 days) chick embryos. Neuroscience Letters 192:61–64; 1995. Kohn, M. J.; Izpisua-Belmonte, J. C.;Abud, H.; Heath, J. K.;Tickle, C. Fibroblast growth factors induce additional limb development from the flank of chick embryos. Cell 80:739–746; 1995. Kostakopoulou, K.;Vargesson, N.; Clarke, J. D.W.; Brickell, P. M.;Tickle, C. Local origin of cells in FGF-4-induced outgrowth of amputated chick wing stumps. Int. J. Dev. Biol. 41:747–750; 1997. Murray, B. M.; Wilson, D. J. A scanning electron microscope study of the normal development of the chick wing from stages 19 to 36. Anat. Embryol. 189:147–155; 1994. O’Farrell, P. J. Unanimity waits in the wings. Nature 368:188–189; 1994. Rawles, M. E. Origin of melanophores and their roles in development of color patterns in vertebrates. Physiol. Rev. 28:383–408; 1944. Rennie, J. Super Sonic. Sci. Amer. April: 20; 1994. Riddle, R. D.; Johnson, R. L.; Lanfer, E.; Tabin, C. Sonic hedgehog mediates the polarizing activity of the ZPA. Cell. 75:1401–1416; 1993. Riddle, R. D.;Tabin, C. J. How limbs develop. Sci. Amer. February: 74–79; 1999. Saunders, J. W., Jr. Cell death in embryonic systems. Science 154:604–612; 1966. Saunders, J. W., Jr. Developmental Biology. New York: Macmillan; 1982. Saunders, J. W., Jr.; Fallon, J. F. Cell death in morphogenesis. In: Locke, M., ed. Major Problems in Developmental Biology. New York: Academic Press; 1967. Saunders, J. W., Jr.; Gasseling, M. T.; Saunders, L. C. Cellular death in morphogenesis of the avian wing. Dev. Biol. 5:147–178; 1962. Saunders, J. W., Jr. The proximal-distal sequence of origin of the parts of the chick wing and the role of the ectoderm. Jour. Exp. Zool. 108:363–404; 1948. Smith, S. M.; Pang, K.; Sundin, O.;Wedden, S. E.;Thaller, C.; Eichele, G. Molecular approaches to vertebrate limb morphogenesis. Development Supplement: 121–131; 1989. Summerbell, D.; Lewis, J. H. Time, place and positional value in the chick limb bud. Jour. Embryol. Exp. Morphol. 33:621–643; 1975. Summerbell, D.The effect of local application of retinoic acid to the anterior margin of the developing chick limb. J. Embryol. Exp. Morph. 78:269–289; 1983. Tabin, C. J. Retinoids, homeoboxes, and growth factors: Toward molecular models for limb development. Cell 66:199–217; 1991. Taylor, G. P.;Anderson, R.; Reginelli,A. D.; Muneoka, K. FGF-2 induces regeneration of the chick limb bud. Dev. Biol. 163:282–284; 1994. Teillet, M.A.; Ziller, C.; LeDourain, N. M. Quail-chick chimeras. In: Sharpe, P.T.; Mason, I., eds. Molecular Embryology Methods and Protocols. Methods in Molecular Biology; Vol. 97.Totowa, NJ: Humana; 1999: 305–318. Thevenet, A.; Sengel, P. Naturally occurring wounds and wound healing in chick embryo wings. Roux’s Archives of Developmental Biology 195:345–354; 1986. Thommes, R. C.; Jameson, K. M. Hypothalamo-adenohypophyseal-thyroid interrelationships in the chick embryo. III.Total T4 levels in the plasma of decapitated chick embryos with adenohypophyseal transplants. Gen. Comp. Endocrinol. 42:267–269; 1980. Thommes, R. C. Ontogenesis of thyroid function and regulation in the developing chick embryo. Jour. Exp. Zool. Suppl. 1:273–279; 1987. (See Metcalfe et al. 1987 earlier.) Tickle, C. Retinoic acid and chick limb bud development. Development Supplement 1:113–121; 1991. Tickle, C.;Alberts, B.;Wolpert, L.; Lee, J. Local application of retinoic acid to the limb bud mimics the action of the polarizing region. Nature 296:564–566; 1982. Vaux, D. L.; Haecker, G.; Strasser, A. An evolutionary perspective on apoptosis. Cell. 76:777–779; 1994.

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Watterson, R. L. Development of the glycogen body of the chick spinal cord. I. Normal morphogenesis, vasculogenesis, and anatomical relationships. Jour. Morphol. 85:337–390; 1949. Weston, J. A. A radioautographic analysis of the migration and localization of trunk neural crest cells in the chick. Devel. Biol. 6:279–310; 1963. Weston, J.A.The migration and differentiation of neural crest cells. Advances in Morphogenesis 1:301–330; 1970. Wolpert, Lewis.The shape of things to come. New Scientist 27 June: 38–42; 1992. Wolpert, L. Pattern formation in epithelial development: the vertebrate limb and feather bud spacing. Phil. Trans. R. Soc. Lond. B 353:871–875; 1998. Zwilling, E. Limb morphogenesis. Advances in Morphogenesis 1:301–330; 1961. CHORIOALLANTOIC MEMBRANE GRAFTING Bertossi, M.;Virgintino, D.; Coltey, P.; Errede, M.; Mancini, L.; Roncalli, L.Vascularization of embryonic adrenal gland grafted onto chorioallantoic membrane. Anatomy and Embryology 198:267–275; 1998. Burton, G. J.; Palmer, M. E. Development of the chick chorioallantoic capillary plexus under normoxic and normobaric hypoxic and hyperoxic conditions: a morphometric study. Jour. Exp. Zool. 262:291–298; 1992. Coulombre, A. J. Grafting of embryonic rudiments. In: Wilt, F. H.; Wessells, N. K., eds. Methods in Developmental Biology. New York:Thomas Y. Crowell, 457–469; 1967. Dossel, W. New method of intracoelomic grafting. Science 120:262–263; 1954. Flamme, I.; Schulze-Osthoff, K.; Jacob, H. J. Mitogenic activity of chicken chorioallantoic fluid is temporally correlated to vascular growth in the chorioallantoic membrane and related to fibroblast growth factors. Development 111:683–690; 1991. Fuenzalida, M.; Lemus, R.; Romero, S.; Fernandez-Valencia, R.; Lemus, D. Behavior of rabbit dental tissues in heterospecific association with embryonic quail ectoderm. Jour. Exp. Zool. 256:264–272; 1990. Gajovic, S.; Gruss, P. Differentiation of the mouse embryoid bodies grafted on the chorioallantoic membrane of the chick embryo. Int. J. Dev. Biol. 42:225–228; 1998. Hardy, M. H.; Dhouailly, D.; Torma, H.; Vahlquist, A. Either chick embryo dermis or retinoid-treated mouse dermis can initiate glandular morphogenesis from mammalian epidermal tissue. Jour. Exp. Zool. 256:279–289; 1990. Rawles, M. E.A study of the localization of organ-forming areas in the chick blastoderm. Jour. Exp. Zool. 72:271–315; 1936. Rawles, M. E. On transplantation of normal embryonic tissues. Ann. N.Y. Acad. Sci. 55:302–312; 1952. Rawles, M. E.The heart-forming areas of the early chick blastoderm. Physiol. Zool. 16:22–42; 1943. Rawles, M. E.Tissue interactions in scale and feather development as studied in dermal-epidermal recombinations. Jour. Embryol. Exp. Morph. 11:765–789; 1963. Ribatti, D.; Loria, M. P.; Tursi, A. Lymphocyte-induced angiogenesis—a morphometric study in the chick embryo chorioallantoic membrane. Acta Anatomica 142:334–338; 1991. Ribatti, D.;Vacca,A.; Roncali, L.; Dammacco, F.The chick embryo chorioallantoic membrane as a model for in vivo research on angiogenesis. Int. J. Dev. Biol. 40:1189–1197; 1996. Sikorski, R.; Peters, R. Metastasis in eggs. Science 281:1823; 1998. Tuan, R. S.; Ono, T. Regulation of extraembryonic calcium mobilization by the developing chick embryo. Jour. Embryol. Exp. Morph. 97:63–74; 1986. Wanek, N.; Gardiner, D. M.; Muneoka, K.; Bryant, S.V. Conversion by retinoic acid of anterior cells into ZPA cells in the chick wing bud. Nature 350:81–83; 1991. Weiss, P.;Taylor,A. C. Reconstitution of complete organs from single-cell suspensions of chick embryos in advanced stages of differentiation. Proc. Nat. Acad. Sci. 46:1177–1185; 1960. West, D. C.; Hampson, I. N.; Arnold, F.; Kumar, S. Angiogenesis induced by degradation products of hyaluronic acid. Science 228:1324–1326; 1985. Zwilling, E. A modified choriollantoic grafting procedure. Transplantation Bulletin 6:115–116; 1958.

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IV. Fern Gametophyte Development

14. Patterns of Fern Gametophyte Development


L A B O R A T O R Y

14 Patterns of Fern Gametophyte Development

Because of the relative ease with which the phases of fern life histories can be observed, investigations of fern reproduction played an important historical role in the elucidation of general reproductive patterns in terrestrial plants. Ferns pose interesting developmental questions because, in the fern life cycle, a single organism produces two very different, independently living plants. Before you begin work on fern development, it would be helpful for you to review the major features of the fern life cycle. A simplified, diagrammatic outline of the life history of a typical fern is represented in figure 14.1.The sporophyte, the spore-producing phase of the fern life history, is the familiar “fern plant” seen growing both in gardens and in the wild.The fern gametophyte (gamete-producing plant) is a relatively inconspicuous, independent plant, the prothallus, upon which antheridia (spermproducing organs) and archegonia (egg-producing organs) develop. In the bracken fern, Pteridium aquilinum, and in Woodwardia sp. and several other commonly investigated fern species, both types of reproductive organs are borne on the same prothallus, but antheridia are produced before archegonia on any given prothallus. The gametophytes of many ferns grow well in laboratory cultures, so the early stages of their development can be traced by direct microscopic observation following transfer of spores to a suitable medium.This direct observation makes it relatively easy to assess the effects of experimental treatments, and a number of developmental biologists have investigated fern gametophyte development. In this laboratory, we will examine the early development of Pteridium aquilinum or a fern that has similar developmental patterns.Your work in this laboratory will be focused on development of the gametophyte phase of the fern’s life cycle. However, if you can maintain your cultures long enough, you will also be able to observe development of young sporophytes.

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Vegetative reproduction

Sporophyte

sori on sporophyll

sorus (cluster of sporangia)

“fiddle head” (developing frond) young sporophyte growing on gametophyte

Meiosis rhizome (underground stem)

roots

mature sporangium with spores

haploid spore

germinating spore two-dimensional growth

sporophyte embryo growing in archegonium

rhizoid

egg archegonium

neck of archegonium sperm young prothallus antheridium antheridium

rhizoids Mature gametophyte (prothallus) Vegetative reproduction

FIGURE 14.1 Generalized fern life cycle.

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Techniques Preparation of Cultures You may grow fern gametophytes from spores or your instructor may elect to start the cultures in advance of the first laboratory period. 1. Fern gametophytes grow well in a liquid culture medium developed by Davis and Postlethwait.The medium contains a mixture of inorganic salts, and its composition is given in Appendix F.After the liquid medium has been prepared, fill your culture dishes about half full. Mark the fluid level on the outside of each dish so that you can add distilled water to the dishes later if evaporation occurs. It is best to position the culture dishes where the plants are to be grown before spores are added. This eliminates handling that tends to wash spores up on the edges of the dishes where they dry out or develop poorly. Sprinkle a very light, scattered layer of spores on the surface of the medium. Do not cover the surface with a dense mat of spores, since overcrowding inhibits development. If you must handle the dishes after the spores are sprinkled, move them carefully with a minimum amount of disturbance of the medium. If you use flasks, stopper them with cotton plugs or cover them with aluminum foil. 2. Keep cultures in an area that receives a moderate amount of light during the day.A north-facing window ledge is a good site, or “soft white” fluorescent desk-lamp bulbs can be used to provide artificial light.A desk lamp with two 15-watt fluorescent bulbs set at a height of 25 to 30 cm above the culture supports good development if the lights are on continuously or for a specified photoperiod of at least 8 to 10 hours per day. Fern gametopyhtes do not grow well under the bright light of sunny, south-facing windows or under very bright lights in certain types of controlled environment chambers. 3. Your culture can be maintained indefinitely and usually requires only occasional checks on fluid level. Contamination usually will not be a serious problem when you use this medium, but if some parts of the culture become contaminated, attempt to “rescue” plants from other areas and transfer them to fresh culture dishes.

Observations Your observations should be made critically and recorded carefully since they will constitute the normal control developmental pattern against which you will make comparisons in the next experimental laboratory. 1. After various time intervals, remove plants from your culture for observation. Since you will probably have a very large number of gametophytes in your culture, do not attempt to return plants to the culture after you have observed them. During early development, you can transfer plants to a slide for observation, using either a bacteriological loop or a pipette. As they grow larger, it may become necessary to handle prothallia gently with a spatula and forceps. Place the gametophytes in a drop of water on the slide, cover the slide with a coverslip, and make observations using the low-power objective of a compound microscope.

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(b)

(a)

(d) (c) FIGURE 14.2 (a) One-dimensional gametophytes with rhizoids growing out of germinated spores. (b) Early two-dimensional gametophyte. (c ) Heart-shaped prothallus showing distinct apical notch. (d ) Enlargement of a portion of prothallus showing several antheridia along the margin.

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2. Check your cultures at daily intervals.When you find germinating spores, begin to record pertinent facts about early development, such as the approximate percentage of germination, the time of emergence, the appearance of the first cell of the protonema (the filamentous chain of cells that will grow out from the spore), and the development of the first rhizoids (see fig. 14.2a). 3. A critically important step in the early development of the gametophyte is an abrupt alteration in growth form. Early growth produces a protonema, a chain of cells lying end to end (one-dimensional growth). Eventually, after a few days of development under ordinary white-light conditions, the division plane of the cell at the end of the filament is oriented differently from previous divisions so that the two daughter cells produced by division come to lie side by side at oblique angles to the linear chain of earlier cells.This change in division pattern marks the beginning of the flat, platelike gametophyte prothallus (two-dimensional growth) (see fig. 14.2b). Record the time sequence of this transition in your culture and note the number of protonema cells present at the time of conversion from one-dimensional to two-dimensional growth. If your microscope is equipped so that you can make the necessary measurements, your instructor may also ask you to determine length-to-width ratios for individual cells or for the entire chain of cells in the protonema at several stages leading up to the time when growth becomes two-dimensional.Volume is also a useful means of quantitative comparison that you can use between controls and plants that you might grow under various experimental conditions. Cells of the protonema are approximately cylindrical, so you can calculate individual cell volumes or total protonemal volume using this formula for the volume of a cylinder:

冢

Mean width Volume ⬵ ␲ ᎏᎏ 2

冣

2

Length

4. As you follow development of young gametophytes, make sketches to record general appearance at various stages. As most areas of the gametophyte are only one cell thick, it is possible to obtain reasonably accurate counts of total cell numbers as development proceeds. Plot these data against time so that you have a reference curve for cell proliferation under your culture conditions that can be used for comparisons later when you grow gametophytes under experimental conditions. You will encounter considerable individual variability, but your results will nevertheless be representative of the general sequence of events in prothallus development. Watch also for differences in cell size and shape in different parts of the prothallus. Such features as chloroplast density and distribution within cells should also be recorded. As the prothallus continues to grow, the appearance of the apical notch (fig. 14.2c) gives it the characteristic heart-shaped form. This apical notch is a meristematic region, that is, a site of continued active cell division that adds cells to the growing prothallus. 5. When the prothalli attain maturity, watch for the development of antheridia.The time in culture required for antheridium formation varies, but depending upon specific culture conditions, you may expect to find antheridia after several weeks in culture. They appear as protuberances in the area of the pointed end of the prothallus (fig. 14.2d ). You can observe the motile fern sperm by transferring the gametophyte with antheridia to a drop of water on a slide. Focus up and down on an antheridium and look for movements of sperm within it. Some antheridia may discharge sperm spontaneously while you are watching, or you may be able to squeeze sperm out of the antheridia by pressing the coverslip gently. Note the corkscrew shape and erratic, spiraling swimming pattern of the sperm. 6. In cultures maintained for longer periods of time, there may be complex outgrowths from individual prothalli that can actually produce clusters of attached prothalli. Examine your cultures occasionally for development of these more complex growth patterns. 7. Development of archegonia may take up to several months under most culture conditions. If you maintain cultures for long periods of time, it would be advisable to transfer at least some of the plants to fresh culture dishes. Repeated examination of your plants for archegonium formation probably would damage them. If you lack large numbers of plants, you may simply want to wait for signs of growth of the young sporophyte plant.

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Materials EQUIPMENT Petri dishes (small Erlenmeyer flasks also may be used for culturing gametophytes) Camel’s hair brush (for shaking spores onto culture surface) Clean microscope slides and coverslips Bacteriological loops or pipettes (ordinary “medicine droppers” also work well) Forceps Small spatula (useful, but not essential for handling older prothalli) Compound microscope Light source for cultures (see Techniques section) SOLUTIONS

AND

CHEMICALS

Fern culture medium LIVING MATERIAL Spores of Pteridium aquilinum, Woodwardia sp., or other fern species whose gametophyte development begins with a linear chain of cells (a protonema)

Some Useful Information Sources www.perspective.com/nature/plantae/ferns.html This website contains beautiful images of developing fern gametophytes and young sporophytes. www.bbg.org/gardening/plants/ferns/growing.html This is a methods website on growing ferns from spores prepared at Brooklyn Botanical Garden by Judith I. Jones. http://amerfernsoc.org This website of the American Fern Society has a number of interesting links. www.bio.utk.edu/cfern/index.html This is an educational website devoted to the C-fern, prepared at the University of Tennessee-Knoxville.

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15. Environmental Effects on Fern Gametophyte Development


L A B O R A T O R Y

15 Environmental Effects on Fern Gametophyte Development

From the earliest stages of their development, and throughout their lives, plants are involved in a continuing series of interactions with their environment. Developing fern gametophytes are no exception to this rule, and in this laboratory, you will have the opportunity to investigate their responses to several environmental influences.These responses are (1) developmental response to light of different wavelengths (following), (2) regenerative responses to loss of parts (p.159), and (3) growth of gametophyte structures from sporophyte tissue placed in an atypical environment (p. 161).

PHOTOMORPHOGENESIS IN FERN GAMETOPHYTES Because of the critical importance of light as the energy source for photosynthesis, we might expect light to be a key factor in plant development and growth, and indeed it is. However, it has long been known that there are direct light effects on plant development that are separate from light’s role in photosynthesis. These effects are grouped under the collective name photomorphogenesis. We can define photomorphogenesis as the control that light exerts over growth and differentiation of a plant, independent of photosynthesis. Many developmental responses are controlled by light intensity, light quality (wavelength), directional illumination, or the relative length of light and dark periods (photoperiodic responses). All of these aspects of illumination must be considered in the interpretation of photomorphogenetic phenomena. Some photomorphogenetic responses in plant development have been known and studied for many years. For example, Charles Darwin and his son, Francis Darwin, experimented on specific plant growth responses to unidirectional illumination. Another strikingly obvious light relationship of plant development that has been investigated for more than 100 years is etiolation. Flowering plants’seedlings, when grown in darkness or very weak light, tend to be exceptionally tall and spindly.They have small, unexpanded leaves and are very pale due to deficient chloroplast development. In some plants, very brief periods of light exposure are adequate to reverse the etiolation effect. In etiolation reversal, as well as in many other photomorphogenesis responses, the most effective part of the light spectrum is the red region.This red-light responsiveness is mediated by a specific light-absorbing pigment, phytochrome, which occurs in two forms that are interconvertible in response to red- and far-red light exposure. In some plant systems, phytochrome-mediated responses become fixed in particular patterns of differentiation after only one light exposure period, but in others, there is the potential for reversal if red-light treatment is followed by a period of exposure to far-red light. For example, in the germination of Grand Rapids lettuce seeds, the response can be switched back and forth repeatedly by alternating red- and far-red illumination.

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Laboratory 15







Light also exerts control over the growth and development of fern gametophytes. In some ferns, for example, light is required for spore germination. Another particularly obvious light control relationship is in the switch from one-dimensional to two-dimensional growth. Gametophytes grown in red light tend to continue the filamentous protonemal growth form and do not acquire the new plane of cell division necessary to produce the flat prothallus. However, this abnormal development in red light is due to disruption of complex light relationships and cannot be interpreted in terms of the phytochrome model (see “Suggestions for Further Investigation” section on p. 169). Nevertheless, this is an interesting example of photomorphogenetic control in plants, and this laboratory will introduce you to the phenomenon. Techniques It is important to begin experiments on photomorphogenesis with caution, because the experiments suggested here can be conducted at several levels of sophistication, as can all experiments on photomorphogenesis.Accurate measurements of light intensity and spectral qualities are possible when proper instruments are available and the wavelength emission of light sources can be specifically controlled. In order to make repeatable quantitative measurements and to draw firm conclusions about a developmental response involving photomorphogenesis, it is essential that such critical methods be employed. However, you can obtain interesting qualitative results from work done at a less sophisticated level, and these procedures are written with that in mind. It would be desirable to make each of the observations in the suggested experiments with large numbers of gametophytes, but you may be somewhat limited by the numbers of plants actually available.Your instructor will make some suggestions about the numbers of observations to be made. Growth in Red Light 1. Follow the same procedure that you used in Laboratory 14 to set up control and experimental cultures. Sow enough spores on the medium to allow for later removal of some gametophytes for transfer without depletion of the red-light culture. Your instructor may elect to schedule these experiments simultaneously with your basic observations on fern gametophyte development. In that case, the term “control culture” may refer simply to your initial gametophyte cultures.You must take great care to shelter your red-light cultures from any other light sources because a small amount of “leaked” white or blue light can have drastic consequences for your results. You may provide either a specified daily photoperiod or continuous illumination for your red-light plants, but you should try to make the experimental and control situations as nearly comparable as possible.Take the same precautions about contamination and fluid level with your red-light cultures as you do with your control cultures and use the same techniques to remove plants for examination. 2. Check your cultures at daily intervals and make the comparisons suggested under Analysis of Results, which follows. Consult your instructor concerning the numbers of plants that should be examined each day.

Transfer from Red Light to White Light You should make the transfer from red light to white light when plants in your red-light cultures have at least as many cells as control plants have at the time that they switch to two-dimensional growth. Your instructor may suggest a specific time for this transfer or may ask you to make a series of transfers at various stages if you have enough plants available. Transfer some plants from your red-light cultures to a new culture that can be moved to the white-light area. Follow the further development of these transferred plants carefully and record the results for comparison with other groups of plants.As another control for this experiment, you may wish to transfer some plants to fresh medium and leave them under red light. Transfer from White Light to Red Light As a reciprocal experiment to the one suggested above, transfer some of the control plants to the red-light area.You should make this transfer after the controls are well into the two-dimensional growth phase.Your instructor may suggest a specific time or leave this choice to you.A control culture of plants transferred to fresh medium, but left under white light, would also be desirable.

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Observations It probably would be most convenient to keep results from the several types of experiments separate from one another as you are taking them, but later you will want to combine data of various kinds in different ways for purposes of comparison and discussion. 1. For all experimental groups, record information on: a. Spore germination and early growth of protonemata and rhizoids. b. General appearance of plants (sketches are very useful). c. Number of cells per plant. d. Cell sizes, shapes, and other characteristics at all stages. e. Chloroplast density and distribution within cells. f. Length-to-width ratios and volume determinations (if you are able to make appropriate measurements). g. One-dimensional and two-dimensional growth relationships and the characteristics of the plants at the time of their switchover. 2. Growth in red light. Make the general observations suggested previously at a series of intervals. Continue observations on the red-light plants at least as long as it takes for control plants to become well-developed plates of cells.You and your instructor may agree to carry the observations on for longer periods of time in order to see if the red-light effects can be maintained more or less indefinitely. 3. Transfer from red light to white light. In transfer experiments, the time course of developmental responses to the change is of vital importance.Thus, it is essential to record all observations in relation to time since transfer. The switch to two-dimensional growth involves changes in cell behavior during cell division, but you should be able to observe changes in cell shape, and possibly, in other characteristics that precede the first oblique cell division. 4. Transfer from white light to red light. Once again, time sequence is critical and times of all observations should be carefully recorded. Begin to examine small samples of plants daily after 3 days in red light. Watch for evidence of reversion to filamentous growths of the type seen in your initial red-light cultures. If you find any evidence of filamentous growth, examine larger samples at regular intervals. Sketch results and record specific information on changes in cell characteristics.

You may encounter a number of difficulties in conducting these experiments, or you may obtain results in one experiment that are difficult to reconcile with those in another, but frequently such difficulties are instructive since careful consideration can provide new insight into the problems of designing, conducting, and interpreting experiments on photomorphogenesis. Some of the variables that you will need to bear in mind while examining and discussing your experimental results are (a) the vastly different effects on photomorphogenetic responses of comparable total light energies at different wavelengths, and uncertainties about the actual spectral qualities of light in your particular experimental situation, ( b) the interaction of light with other environmental factors, (c) the total radiant energy actually reaching plants in any given experiment, and (d) the question of photosynthetic rates and their bearing on growth rates in various experiments.

REGENERATION BY FERN GAMETOPHYTES As with many other plants, the gametophytes of many fern species show regenerative growth in response to damage or loss of parts. Pieces of the gametophytes of some species regenerate complete prothalli after they are cut into pieces with a blade or even after they are fragmented in a homogenizer. In this laboratory, you will have an opportunity to design and conduct experiments on fern gametophyte regeneration. Techniques 1. Plan an experiment to test the regenerative capacity of different sections of the prothallus from its apex to its base and an experiment to test the effect of fragment size on regeneration.

Laboratory 15

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Vegetative reproduction

Sporophyte

sori on sporophyll

sorus (cluster of sporangia)

“fiddle head” (developing frond) young sporophyte growing on gametophyte

Meiosis rhizome (underground stem)

roots

Apospory (see text)

Apogamy (see text)

mature sporangium with spores

haploid spore

germinating spore two-dimensional growth

sporophyte embryo growing in archegonium

rhizoid

egg archegonium

neck of archegonium sperm young prothallus antheridium antheridium

rhizoids Mature gametophyte (prothallus) Vegetative reproduction

FIGURE 15.1 A generalized diagram showing how development by apogamy or apospory bypasses normal events in the fern life cycle.

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2. To initiate prothallus regeneration, use a razor blade to cut up several heart-shaped prothalli into fragments, examine them microscopically, record their sizes, sketch their shapes, and transfer them to separate culture dishes where you can follow regenerative growth. Be sure to record data concerning which portion of the original prothallus each fragment came from. Place the cut fragments on moist mineral agar medium in a petri dish. ( It is much easier to follow regenerative growth of individual gametophytes if they are placed on agar rather than in liquid culture medium.)

Observations 1. How is new growth initiated on each fragment? When and where does new growth begin? Record data on position and time of initiation of new outgrowths. 2. Are there quantitative relationships between fragment size and pattern of regeneration? Do some fragments produce more than one regenerated prothallus? Record data on these relationships. 3. Are the patterns or extent of regeneration related to the portion of the prothallus from which a fragment comes? What are the relationships?

PROTHALLUS GROWTH FROM SPOROPHYTE TISSUE Ferns have been important subjects in research on the developmental basis of the production of the two distinct morphological expressions that characterize plant life histories. It was originally hypothesized that the gametophyte prothallus structure is the inevitable product of a growth pattern characteristic of haploid cells and that sporophyte generation cells develop structures characteristic of the sporophyte plant simply because they are diploid. Neither of these hypotheses were verified following experimental testing. It is now clear that environmental factors play a role in determining the developmental patterns that produce gametophytes or sporophytes and that those developmental patterns can be experimentally modified. For example, apogamy, the direct production of sporophyte plants from gametophytes without gamete fusion, can be induced in some species of ferns by adding sugar to a mineral agar medium on which the plants are grown (see “Suggestions for Further Investigations” section), and apospory, the direct growth of a prothallus from sporophyte tissue without intervening spore formation, can be induced in some ferns (fig. 15.1). In this experiment, you can attempt to induce apospory by physically altering the environment for part of a young fern sporophyte. Techniques 1. Obtain a fern gametophyte with an attached young sporophyte with its characteristic first leaf. Carefully detach the blade of the leaf. 2. Transfer the sporophyte leaf to a petri dish and lay it down flat on mineral agar medium that has been moistened by adding fern medium. 3. It is important that the leaf be in contact with the surface. If it curls, it may be necessary to place a small fragment of glass on it to hold it down against the agar. Your instructor will provide small fragments of broken coverslips if they are needed, but do not attempt to break up a coverslip yourself unless your instructor directs that you do so and provides necessary safety information.

Observations 1. Is there an outgrowth from the edges of the leaf ? If so, what form does it take? Is there one outgrowth, or are there several? Is there an end-to-end chain of cells resembling a protonema, or does each outgrowth involve a number of cells in some other arrangement? Record qualitative and quantitative data. 2. Do the outgrowths eventually come to resemble ordinary prothalli? Are they the same shape? How do they compare in size to ordinary prothalli at the time that apical notch development is evident? 3. If you are able to maintain your cultures long enough to do so, determine whether there is evidence of antheridium or archegonium development.

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Materials EQUIPMENT Petri dishes for liquid medium cultures (small Erlenmeyer flasks may also be used for culturing gametophytes) Clean microscope slides and coverslips Broken coverslip fragments ( Prothallus growth from sporophyte tissue experiment) Single-edged razor blades ( Regeneration and prothallus growth from sporophyte tissue experiments) Bacteriological loops and pipettes (ordinary “medicine droppers” work well) Compound microscope Dissecting microscope Light sources for control and red-light cultures SOLUTIONS

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CHEMICALS

Fern culture medium Mineral agar medium poured in petri dishes (Regeneration and prothallus growth from sporophyte tissue experiments) LIVING MATERIAL Spores of Pteridium aquilinum, Woodwardia sp., or other fern species whose gametophyte development begins with a linear chain of cells (a protonema) Fern gametophytes (prothallia) that are heart shaped with an apical notch but do not yet have antheridia or archegonia ( Regeneration experiment) Young fern sporophytes at the first-leaf stage ( Prothallus growth from sporophyte tissue experiment)

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16. Sexual Reproduction in the Water Fern, Marsilea


L A B O R A T O R Y

16 Sexual Reproduction in the Water Fern, Marsilea

Early phases of gametophyte development in Pteridium, Woodwardia, and other ferns with similar development provide good opportunities for observation of cellular details of developmental patterns. However, it takes several weeks (or more) for these ferns to develop from spore germination through differentiation of antheridia and archegonia to fertilization and the beginning of sporophyte development. It is possible to observe sexual reproduction in a much shorter period of time using aquatic ferns of the genus Marsilea, commonly called “water clover.” Development from spore germination to the beginning of sporophyte plant growth requires only a few days in Marsilea. Marsilea spores are borne inside small, hard structures called sporocarps that develop on small branches of petioles near the point where they emerge from the horizontal stem (fig. 16.1a). Unlike Pteridium and Woodwardia, which are homosporous (they produce only one type of spore), Marsilea is heterosporous.That is,it produces two distinctly different types of spores,megaspores and microspores. In Marsilea, each sporocarp contains many of each spore type (fig. 16.1b). Megaspores are large, eggshaped structures that are about 0.75 mm in length, while microspores are spherical and about onetenth the size of megaspores. Following germination, each megaspore produces a simple female gametophyte that includes an archegonium with an egg.The much smaller, round microspores develop into simple male gametophytes. A male gametophyte has two antheridia, each of which produces and releases sixteen sperm. After fertilization, the zygote’s development begins within the confines of the archegonium (fig. 16.2). Zygote development induces the gametophyte tissue to produce a bright green sheath around the growing first leaf. Within a couple of days, both a leaf and a root emerge, and development of a sporophyte plant proceeds rapidly. Techniques 1. Marsilea gametophyte development can be initiated by cutting a sporocarp in half and immersing it in fern medium in a petri dish. It is best to cut the sporocarp in an enclosed place because the parts of the dry, hard sporocarp can fly away and be lost. 2. Place your culture in a well-lighted place at room temperature. In a few minutes, colloidal material absorbs enough water so that a gelatinous ring with attached clusters of spores emerges. Once the spores have emerged, gametophyte development proceeds quickly, and fertilization in some species of Marsilea occurs in less than 10 hours time. When you are confident that fertilization has occurred (surely by 24 hours after hydration), transfer a number of the female gametophytes/megaspores to a dish containing fresh culture medium. You can do this using a

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leaf

sporocarp

rhizome

roots

(a)

gelatinous ring megaspore microspores

sporocarp wall

(b)

FIGURE 16.1 (a) Part of a Marsilea plant showing sporocarps that develop on short lateral branches of petioles. (b) Longitudinal section of a sporocarp. (Marsilea sporocarps are about 5 mm long.)

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embryonic leaf of sporophyte female gametophyte

embryonic root of sporophyte rhizoid of female gametophyte primary leaf of sporophyte

nutrient storage cell of female gametophyte

sporophyte root

(a)

(b)

FIGURE 16.2 (a) About 48 hours after fertilization, the greenish, triangular Marsilea embryo is evident inside the female gametophyte. The primordia of the first leaf and the first root are visible. The filaments emerging to the right are rhizoids of the gametophyte. (b) Within 4 or 5 days, the developing sporophyte has a well-developed, green primary leaf and a root. (In each diagram, the nutrient storage cell is about 0.75 mm long.)

Pasteur pipette. This step is helpful because bacteria growing on dead sperm and degenerating empty male gametophytes could foul the medium and might affect developing embryo sporophytes. Sporophytes develop rapidly and emerge from the gametophyte within 2 or 3 days. Observations of all of these developmental events can be made with a dissecting microscope.

Observations 1. How much time elapses between the cutting of the sporocarp tip and the emergence of the gelatinous ring bearing clusters of spores? 2. How many megaspores and microspores are there in each cluster? Are the number of spores in clusters uniform or variable? 3. How long do the spores remain clustered before they disperse in the medium? 4. When are swimming sperm released? Are there any evident changes in male gametophytes that precede sperm release? What sperm structural features can you observe in sperm transferred on a slide to a compound microscope? How long are free-swimming sperm present in the medium? 5. The megaspore develops into a small, simple female gametophyte at the tip of a larger nutrient storage cell. What is the approximate size ratio between gametophyte and storage cell? 6. A mass of gelatinous material secreted around the female gametophyte attracts numerous sperm that become embedded in it. Can you devise a means of visualizing this material around one gametophyte removed from your culture? 7. Details of early sporophyte development are difficult to observe through the wall of an intact gametophyte, but eventually a small triangular embryo becomes faintly visible (fig. 16.2).The developing embryo induces gametophyte tissue to develop into a sheath, the calyptra, around the growing leaf portion of the embryo.At about this time, surface cells of the gametophyte develop rhizoids, and chloroplasts develop in the calyptra and other gametophyte cells. How long after the times of culture initiation and sperm release can you first detect greening of gametophyte tissue? 8. When is an emerging root and/or green-sheathed leaf first visible? When do root hairs develop on the root?

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Once a root and leaf have emerged, you might wish to transfer the young sporophyte to the surface of a pot of very wet soil. It may be necessary to set a jar over the pot to keep the soil moist. It is usually possible to transplant Marsilea sporophytes later if they outgrow the space available. Additional Experiments with Marsilea If your instructor asks you to extend your work with Marsilea, here are some suggestions for experiments that you might add. In each case, consider the questions and hints provided. Then design a hypothesis and experimental protocol for your investigation. Decide on the types and frequency of observations that you will need to make to adequately test your hypothesis. Be certain to make provisions for appropriate control observations, set standards for quantification of your data, and give some preliminary thought to how you might wish to present results of your investigation. Some suggestions for additional experiments follow. Growth in Red Light or Darkness Questions: Does development proceed normally in red light? Do developing female gametophytes and embryo sporophytes appear to be structurally normal when they are maintained in red light or in darkness? Do developmental processes proceed at the same rate as they do under normal illumination? Are the gametophytes’ rhizoids normal? Is embryonic sporophyte root and shoot development normal? The embryonic sporophyte is green from very early in its development. Does its growth depend upon it being photosynthetically active, or can its growth remain normal, sustained by nutrients contained in the megaspore? Hints: Rhizoid lengths can be measured and calculated using an ocular micrometer, as can sporophyte root and shoot lengths, as well as their approximate volumes. Make certain that you begin with enough female gametophytes so that you can discard the ones that you remove from culture for detailed observation or measurement. Why is this necessary? Crowding Effects Questions: Do growing sporophytes influence the growth of their neighbors in positive or negative ways? That is, is there evidence of growth promotion or growth inhibition among developing sporophytes? If there should be evidence of inhibition, is it absolute (i.e., no growth), or is there only retarded growth of roots and shoots? Hints: You might be able to detect interactive effects by transferring various numbers of female gametophytes/megaspores after fertilization has occurred to small volumes of medium in small containers such as the wells of multiwell cell culture plates (“clusters”). In this case, you will need to make observations by scanning the various sets because removing individuals for observation would disrupt the crowding relationship. This assumes that these particular experiments would be done under normal illumination. Temperature Effects on Development Questions: How broad is the range of temperatures under which sporophyte development will occur? Do temperatures near the extremes of the organism’s temperature tolerance cause abnormal growth or growth retardation? Hints: It might be wise to avoid temperature shocks caused by abrupt transfer from the initial temperature under which fertilization has occurred to the various experimental temperatures that you choose. Problems can be avoided if culture dishes are placed in the experimental temperature environments just after organisms are transferred to them.Thus, the cultures come to the experimental temperatures gradually. You may wish to have data on how long this temperature change takes. Observations should be made efficiently so as to limit temperature change of the various cultures. You may wish to make repeated observations of specific individuals.This can be done if single individuals are isolated in individual small petri dishes or in the wells of multiwell cell culture plates.

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Materials EQUIPMENT Standard 100-mm or 60-mm petri dishes Razor blade Pasteur pipettes with bulbs Dissecting microscope Compound microscope Multiwell cell culture plates (“clusters”) for optional experiments SOLUTIONS

AND

CHEMICALS

Fern culture medium LIVING MATERIAL Marsilea sporocarps

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Suggestions for Further Investigation of Fern Gametophyte Development


SUGGESTIONS FOR FURTHER INVESTIGATION OF FERN GAMETOPHYTE DEVELOPMENT The fern life history, with its independent gametophyte generation, holds a special fascination for biologists, and ferns have proven to be very useful subjects for investigation of many aspects of vascular plant biology. Fern spores are easy to collect and, under proper conditions, can be stored for some time.Thus, fern gametophytes can be grown at will, which makes them convenient subjects for use in courses such as developmental biology. In the next several sections, you will find suggestions for further investigation of topics to which you have been introduced in the fern labs of your course. Fern Development Patterns Ferns have long been employed in analyses of plant reproductive patterns (Whittier 1977; Sheffield and Bell 1987;Whittier 1995) and continue to be investigated from an evolutionary perspective (for example, Wolf et al 1988; Mui 1991). Fern gametophyte development is a relatively simple model system for study of the cellular aspects of plant morphogenesis (for example, Dyer 1983; Cooke and Racusen 1988). Techniques for culturing fern gametophytes are well worked out (Laetsch 1967; Davis and Postlethwait 1966; Dyer 1983; Chilton and Graham 1988; Raghavan 1989) so this system is readily accessible to developmental biology students. The transition from linear protonemal growth to two-dimensional growth is one of the most interesting morphogenetic events in early fern gametophyte development (Cordero and Szewczak 1994). Many aspects of the transition have been thoroughly described (for example, see papers by B. D. Davis and his colleagues; Racusen and Cooke 1982), but details of some cellular aspects need further work, and diverse approaches and techniques (for example,Tilney, et al 1990; Grill 1999) promise interesting new insights. This pattern transition is also very susceptible to environmental influences (see next section). All aspects of gametophyte development present interesting questions for investigation: from spore germination (Raghavan 1992) to the activity of the apical cells (Reynolds 1979; Gifford 1983) that play a role similar to that of meristematic regions in more familiar developing plant systems, and even to hormonal control of gamete-producing structures (Stevens and Werth 1999). You might wish to repeat and extend some of the experiments that have been done on various life cycle phases. For example,Whittier and Steeves (1960) reported that apogamy (the direct production of sporophyte plants without gamete fusion) could be induced in sterile cultures of one strain of Pteridium grown on agar-mineral medium enriched with glucose. They found that 2.5% glucose was the optimal concentration for the induction of sporophyte development. The apogamous sporophytes ranged from structures that were morphologically intermediate between gametophyte and sporophyte to quite normallooking young sporophyte plants.The induction of apogamy in other strains of Pteridium required higher sugar concentrations and longer periods of exposure. If you have prothalli available, you could experiment with various glucose concentrations and exposure times to see if apogamy can be induced in the particular fern that you are studying (also consult Raghavan 1989, for additional references). In the context of biotechnology, there is considerable interest in study of plant cells that have had their cell walls removed since such cells (protoplasts) are much easier to manipulate genetically than cells with their walls intact.Techniques have been developed for removing fern cell walls (Partanen et al 1980; Sheffield and Bastin 1981), and you might find it very interesting to try your hand at preparing fern protoplasts and using them in experiments (Maeda et al 1990).You might also wish to study some details of the young sporophyte’s development (see White and Turner 1995). Environmental Effects The initial outgrowth of the fern protonema from the spore can be oriented by unidirectional illumination. In addition to testing this growth polarity, it is interesting to experiment with a possible phototropic response in which growing tips turn in another direction if the angle of illumination is changed. Such a response can also be tested by switching illumination from above to unidirectional illumination from the side. It is also possible to investigate the relative effectiveness of white and red

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light in the phototropic response (Dyer 1983). For these types of experiments, it is best to grow gametophytes on agar medium, and the techniques for doing so are described by Davis and Postlethwait (1966) and by Chilton and Graham (1988)—see Fern Development Patterns section. Davis (1968) did experiments to determine the minimal length of exposure to white light necessary to induce gametophytes previously grown in red light to switch to two-dimensional growth.Basically, his experiments involved growing gametophytes in red light until they showed obvious characteristics of red-light treatment, transferring samples to white light for various periods and then returning them to red light. Then he determined the percentage of each group that had entered the two-dimensional phase after their return to red light (actually the difference between the percentage of two-dimensional gametophytes at the end of the white-light exposure and at the end of 72 hours back in red light).You might find it interesting to experiment with this system using his methods. Although conditions vary, you might start your experiments with various white-light exposures between 12 and 60 hours in length. You can avoid some interpretation problems if you use continuous illumination throughout the experiment. Although the fern gametophyte’s switchover from one-dimensional to two-dimensional growth does not show the red–far-red reversibility characteristic of phytochrome-mediated responses, the photoorientation pattern of chloroplasts in protonemal cells changes in response to red–far-red illumination alterations (Kagawa et al 1994). Marsilea Development Reproductive and developmental processes of the water fern, Marsilea, differ in several ways from those of the ferns commonly studied in teaching laboratories. If you want additional general information about this interesting fern species, you might begin by consulting Laetsch (1967) and Settle (1980). The Marsilea sporocarp is a very hardy structure that is capable of remaining viable for a long time ( Johnson 1985). It is quiescent until it is cut or broken open in some other way. Then the megaspore begins the rapidly completed series of steps in megagametophyte development leading to egg production (Machlis and Rawitscher-Kunkel 1967; Bell 1985). Microgametophyte development and, eventually, sperm maturation (Myles and Hepler 1982) are also completed within a few hours, so that fertilization (Myles 1978) occurs within about 10 hours of sporocarp hydration. If you wish to read more about some of these processes, consult Settle (1980) and the references previously cited. If you want to do further experiments on Marsilea development, you might consult Mahlberg and Yarus (1977) and consider experimenting with the effects of environmental factors on megaspore germination and sporophyte formation. Rhizoid development (Bloom and Nichols 1972) is an easily observable aspect of megagametophyte development and might be an interesting process to investigate (see also Miller and Greany 1976). If the embryonic shoot of the young sporophyte is microsurgically removed,small apical fragments can be cultured (Laetsch 1967).Photomorphogenetic responses of the young sporophyte plant (sporeling) have been studied (Laetsch and Briggs 1962), and these responses could be investigated further.Another fern that has a number of interesting features for study is Ceratopteris richardii, which is marketed as the C-Fern (Renzaglia et al 1995; Hickok et al 1998). Fern Development References General Sources Bold, H. C. Morphology of plants. 5th ed. New York: Harper and Row; 1986. Burgess, J. An introduction to plant cell development. Cambridge: Cambridge University Press; 1985. Foster,A. S.; Gifford, E. M., Jr. Comparative morphology of vasular plants. San Francisco:W. H. Freeman & Co.; 1959. Laetsch, W. M. Ferns. In: Wilt, F. H.; Wessells, N. K., eds. Methods in developmental biology. New York: Thomas Y. Crowell, 1967: 319–328. Raghavan, V. Developmental biology of fern gametophytes. Cambridge: Cambridge University Press; 1989. Steeves,T. A.; Sussex, I. M. Patterns in plant development. 2d ed. Cambridge: Cambridge Univ. Press; 1989. Wareing, P. F.; Phillips, I. D. J. Growth and differentiation in plants. 3d ed. Oxford: Pergamon; 1981. Wilkins, M. Plantwatching: how plants remember, tell time, form partnerships and more. New York: Facts on File; 1988.

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Reviews and Research Papers FERN DEVELOPMENT PATTERNS Chilton, G.; Graham, L. C. Culturing fern gametophytes on solid mineral media for classroom study. Jour. Biol. Education 22:110–112; 1988. Cooke,T. J.; Racusen, R.H.The growth forms of developing fern gametophytes: a theoretical consideration of form and function relationships. Annals of Botany 62:633–641; 1988. Cordero, R. E.; Szewczak, C.A.The developmental importance of cell division. Amer. Biol.Teacher. 56:176–179; 1994. Davis, B. D. The transition from filamentous to two-dimensional growth in fern gametophytes. II. Kinetic studies on Pteridium aquilinum. Amer. Jour Bot. 56:1048–1053; 1969. Davis, B. D. The transition from filamentous to two-dimensional growth in fern gametophytes. III. Interaction of cell elongation and cell division. Amer. Jour. Bot. 58:212–217; 1971. Davis, B. D.; Chen, J.C.W.; Philpott, M.The transition from filamentous to two-dimensional growth in fern gametophytes. IV. Initial events. Amer. Jour. Bot. 61:722–729; 1971. Davis. B. D.; Postlethwait, S. N. Classroom experimentation using fern gametophytes. Amer. Biol.Teacher. 28:97–102; 1966. Dyer,A. F. Fern gametophytes in culture—a simple system for studying plant development and reproduction. Jour. Biol. Education 17:23–39; 1983. Gifford, E. M., Jr. Concept of apical cells in bryophytes and pteridophytes. Ann. Rev. Plant Physiol. 34:419–440; 1983. Maeda, M.; Sugimoto, Y.; Nakamura, M.; Masuda, K.; Kaneko, H.; Sugai, M. Division and gametophytic tissue formation from protoplasts of young sporophytes in fern Lygopodium japonicum. Plant Cell Reports 9:113–116; 1990. Mui, K.T. Evolutionary studies of the chloroplast genome in Pteridium. Plant Science 77:81–92; 1991. Partanen, C. R.; Power, J. B.; Cocking, E. C. Isolation and division of protoplasts of Pteridium aquilinum. Plant Science Letters 17:333–338; 1980. Racusen, R. H.; Cooke, T. J. Electrical changes in the apical cell of the fern gametophyte during irradiation with photomorphogenetically active light. Plant Physiol. 70:331–334; 1982. Raghavan, V. Germination of fern spores. Amer. Scientist 80:176–185; 1992. Reynolds,T. L. Apical dominance in Anemia phyllitidis gametophytes. Amer. Fern Jour. 69:92–96; 1979. Sheffield, E. Alternation of generations in ferns: mechanisms and significance. Biol. Rev. 69:331–343; 1994. Sheffield, E.; Bastin, J. H. Isolation and application of plant protoplasts. School Science Review 62:489–491;1981. Sheffield, E.; Bell, P. R. Current studies of the pteridophyte life cycle. Bot. Rev. 53:442–490; 1987. Stevens, R. D.;Werth, C. R. Interpopulational comparison of dose-mediated antheridiogen response in Onoclea sensibilis. Amer. Fern Jour. 89:221–231; 1999. Tilney, L. G.; Cooke,T. J.: Connelly, P. S.;Tilney, M. S.The distribution of plasmodesmata and its relationship to morphogenesis in fern gametophytes. Development 110:1209–1221; 1990. White, R. A.;Turner, M. D. Anatomy and development of the fern sporophyte. Bot. Rev. 61:281–305; 1995. Whittier, D. P. The value of ferns in an understanding of the alternations of generation. BioScience 21:225–227; 1971. (Five additional papers on “Ferns as Tools in Solving Biological Problems” follow this one. These valuable papers are on pp. 266, 271, 313, 317, and 323 in Vol. 21 of BioScience.) Whittier, D. P.; Steeves,T.A.The induction of apogamy in the bracken fern. Canadian Jour. Bot. 38:925–930; 1960. Wolf, P. G.; Haufler, C. H; Sheffield, E. Electrophoretic variation and mating system of the clonal weed Pteridium aquilinum (L. Kuhn). Evolution 42:1350–1354; 1988. ENVIRONMENTAL EFFECTS Bell, P. R.; Richards, B. M. Induced apospory in polypodiaceous ferns. Nature 182:1748–1749; 1958. Binding, H.; Mordhorst, G. Gametophyte regeneration and apospory from archegoniate protoplasts under conditions devised for higher plants. Bot. Acta 104:330–335; 1991. Davis, B. D. Effect of light quality on the transition to two-dimensional growth by gametophytes of Pteridium aquilinum. Bull. Torrey Bot. Club 95:31–36; 1968. Grill, R. Calcium requirement in blue-light-promoted and red-light-inhibited anteridiogenesis in the fern Anemia phyllitidis (L.) Sw. J. Plant Physiol. 145:285–290; 1995. Kawaga,T.; Kadota,A.;Wada, M. Phytochrome-mediated photoorientation of chloroplasts in the protonemal cells of the fern Adiantum can be induced by brief irradiation with red light. Plant Cell Physiol. 35:371–377; 1994. Partanen, C. R.; Power, J. B.; Cocking, E. C. Isolation and division of protoplasts of Pteridium aquilinum. Plant Science Letters 17:333–338; 1980.

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Sheffield, E.;Attree, S. M. Further experiments with ferns in culture: regeneration. Jour. Biol. Education 17:183–184; 1983. Sheffield, E.; Bell, P. R. Experimental studies of apospory in ferns. Annals of Botany 47:187–195; 1981. Smith, D. L.; Robinson, P. M.The effects of fungi on morphogenesis of gametophytes of Polypodium vulgare L. New Phytologist 68:113–122; 1969. Sobota, A. E.; Partanen, C. R. The growth and division of cells in relation to morphogenesis in fern gametophytes. I. Photomorphogenetic studies in Pteridium aquilinum. Canad. Jour. Bot. 44:497–506; 1966. Uchida, K.; Furuya, M. Control of the entry into S phase by phytochrome and blue-light receptor in the first cell cycle of fern spores. Plant Cell Physiol. 38:1075–1079; 1997. Whittier, D. P. The influence of growth substances on the induction of apogamy in Pteridium gametophytes. Amer. Jour. Bot. 53:882–886; 1966. Whittier, D. P.; Steeves,T. A.The induction of apogamy in the bracken fern. Can. Jour. Bot. 38:925–993; 1960. MARSILEA DEVELOPMENT Bell, P. R. Maturation of the megaspore in Marsilea vestita. Proc. Roy. Soc. Lond. 233B:485–494; 1985. Bloom,W.W.; Nichols, K. E. Rhizoid formation in megagametophytes of Marsilea in response to growth substances. Amer. Fern Jour. 62:24–46; 1972. Hickok, L. G.;Warne,T. R.; Baxter, S. L.; Melear, C.T. Sex and the C.-fern: not just another life cycle. Amer. Biol.Teacher 48:1031–1037; 1998. Johnson, D. M. New records for longevity of Marsilea sporocarps. Amer. Fern Jour. 75:30–31; 1985. Laetsch, W. M.; Briggs, W. R. Photomorphogenetic responses of sporelings of Marsilea vestita. Plant Physiol. 37:142–148; 1962. Machlis, L.; Rawitscher-Kunkel, E.The hydrated megaspore of Marsilea vestita. Amer. Jour. Bot. 54:689–694; 1967. Mahlberg, P. G.;Yarus, S. Effects of light, pH, temperature, and crowding on megaspore germination and sporophyte formation. Marsilea. Jour. Exp. Bot. 28:1137–1146; 1977. Miller, J. H.; Greany, R. H. Rhizoid differentiation in fern spores: experimental manipulation. Science. 193:687–689; 1976. Myles, D. G.The fine structure of fertilization in the fern Marsilea vestita. Jour. Cell Sci. 30:265–281; 1978. Myles, D. G.; Hepler, P. K. Shaping of the sperm nucleus in Marsilea: a distinction between factors responsible for shape generation and shape determination. Dev. Biol. 90:238–252; 1982. Renzaglia, K. S.; Warne, T. R.; Hickok, L. G. Plant development and the fern life cycle using Ceratopteris richardii. Amer. Biol. Teacher 57:438–442; 1995. Settle,W. J. Using water clover to demonstrate sexual reproduction in ferns. Amer. Biol. Teacher 42:295–297; 1980.

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V. Emerging and Regenerating Developmental Patterns

17. Pattern Regeneration and Reorganization in Protists and Animals


L A B O R A T O R Y

17 Pattern Regeneration and Reorganization in Protists and Animals

Much of the emphasis in developmental biology is on the initial development and differentiation of cells, tissues, and organisms, but the events involved in replacement of lost parts is also an important developmental biology topic. Regenerative capacities vary considerably among organisms. For example, we mammals have welldeveloped healing capacities but have a very limited ability to replace lost parts. On the other hand, many organisms can regenerate and reorganize following substantial losses. In this set of experiments, you will examine the capacities of several organisms to replace lost parts.

SPIROSTOMUM Members of the genus Spirostomum are unusually large ciliated protozoans.In nature,Spirostomum is often found in quiet ponds, especially in places where there are decaying leaves. A Spirostomum cell may be up to 3 or 4 mm in length,and a large,multilobed macronucleus stretches through about two-thirds of the cell’s length (fig. 17.1). Numerous micronuclei are scattered in the cytoplasm along the length of the macronucleus. Because of this arrangement, even a small cut fragment of a Spirostomum cell is likely to contain some micronuclei and a portion of the macronucleus. Membranelles (large modified cilia), arranged in rows, move particles into a food groove that leads to a cytostome where food particles are incorporated into food vacuoles. Near the posterior end of the cell, a large contractile vacuole is visible even when the organism is observed with a dissecting microscope. Techniques 1. Spirostomum can be observed and manipulated in a depression slide on the stage of a dissecting microscope. Transfer a cell to a drop of spring water or dilute Amphibian Ringer’s solution. Take some time to observe the organism’s structure and behavior. Note the prominent contractile vacuole visible near the posterior end of the cell. The presence of a contractile vacuole in each cell within a short time following cutting will serve as an easily observable marker of cell reorganization and regeneration.

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membranelles

cytostome

contractile vacuole

FIGURE 17.1 Sketch of a Spirostomum ambiguum cell showing various organelles. Spirostomum is a very large ciliated protozoan that ranges in size from about 1 mm up to about 3 or 4 mm in length. Note the size and location of the contractile vacuole. The zone of membranelles, which are modified cilia, usually can be seen, as can the cytostome where food particles are taken in. The beadlike macronucleus and scattered micronuclei can be seen with phase contrast or differential interference microscopy, but a dissecting microscope provides adequate magnification for general observations and for cutting Spirostomum cells.


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2. The Spirostomum cell can be cut by laying a cactus-spine needle across it and gently pressing down. Although a moving Spirostomum cell occasionally can be cut by one quick, well-aimed stroke, often the organism strongly contracts and avoids being cut. An easier technique takes advantage of the organism’s response to touch. Gently touch the organism and then quickly cut across it during the momentary pause that occurs when it stops moving and contracts strongly in response to being touched. 3. If one of the fragments you have cut is quite large, you might try to cut it in half so that you have a total of three fragments. 4. The cut cell fragments can be left in the depression slide if you place the slide in a petri dish lined with a folded, soaked paper towel, which will add humidity to the air and thus retard evaporation from the depression after the dish is covered.

Observations You can observe some features of the cut fragments and the reorganization process with a dissecting microscope, but more detailed observations will require use of a compound microscope. Phasecontrast optics or a dark-field arrangement will facilitate your observations. 1. Make note of the swimming behavior of the fragments at several time intervals after cutting. Do the fragments resume swimming immediately? Do they swim continuously? Does each have a definite anterior end that is directed forward most of the time? How do anterior and posterior ends of the fragments relate to their original orientation as part of the intact cell? 2. Can you see the contractile vacuole in the fragment derived from the posterior end of the cell? Are the membranelles actively beating? 3. When does a contractile vacuole become visible in the fragment (or fragments) that did not have one after the cell was cut? What other organizational changes can you detect in the fragments over time?

Materials EQUIPMENT Dissecting microscope Compound microscope Depression slides Petri dishes for moist chambers Cactus-spine microsurgical needles Pipettes and bulbs SOLUTIONS

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CHEMICALS

Spring water (or 10% Amphibian Ringer’s solution) LIVING MATERIAL Culture of Spirostomum

HYDRA Hydras are small freshwater cnidarians.A hydra has a simply organized, two-layered cylindrical body column. One end of a hydra is attached to a surface by an adhesive basal disc, or foot. At the opposite end of the body column is a mouth that is surrounded by a ring of tentacles. The cellular organization of a hydra’s body is always in a state of flux because cell replacement is a continuous process. Cells in each of the several cell lines present in the body continue to divide and replace lost cells.This cell division also contributes to asexual reproduction by budding. In well-fed hydras, buds regularly form in a region just below the midpoint of the body column. A bud grows, develops a mouth and tentacles, and begins to feed while still attached to the body column. It then detaches and begins life as an independent individual.

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Experiments on hydra regeneration and reorganization began with Abraham Trembley’s eighteenth century observations of the restitution of body organization in cut fragments of hydras. Many subsequent investigations have revealed a remarkable ability to reorganize body form following various disruptions, even including dissociation of the hydra into single cell suspensions. Other studies have focused on the roles of various diffusible regulator substances in determining patterns of hydra growth and regeneration. This experiment deals with basic aspects of form restitution in hydras cut into several fragments. Techniques 1. Observe a hydra in water in a small dish. Make a simple sketch of the body and note the number of tentacles present. 2. When the animal is quite relaxed and extended, use an iris scissors to snip off the upper third of the body.You will find that you need to use a slow, cautious approach and a fairly quick snip to cut a hydra, since it may contract very strongly if you disturb it before you make your cut. ( Please be very careful in handling the scissors to avoid damage to its sharp, but rather fragile blades.) Some students successfully cut hydras using two cactusspine microsurgical needles, but most prefer to use iris scissors. 3. If possible, cut the lower part of the body column again so that there are three fragments. This second cut is optional, however, and should not be attempted if it appears that it will cause massive damage to the body. 4. After you have noted or sketched the appearance of each fragment, cover the dish and set it aside for later examination. 5. If you maintain your hydras for several days, periodically remove some medium with a pipette and replace it with fresh medium.

Observations 1. Observe the fragments at regular intervals after cutting. How long do the fragments remain contracted? 2. Are there any changes in the tentacles of the upper fragment? 3. When do tentacles begin to appear on the other fragment(s)? How many tentacles develop? 4. When does the upper fragment appear to have a functional basal disk attached to the dish? 5. If you succeeded in cutting three fragments, can you make any judgments about polarity in the reorganizing middle fragment? 6. If Artemia (brine shrimp) larvae are available, you might wish to examine prey capture and feeding responses by the regenerated hydras, but food should be added only after all fragments have tentacles and are firmly attached to the bottom of the dish.

Materials EQUIPMENT Dissecting microscope Pipettes (with wider mouth than Pasteur pipettes) Small covered dishes (for example, 60-mm petri dishes) Iris scissors (iridectomy scissors) SOLUTIONS

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CHEMICALS

Spring water (or 10% Amphibian Ringer’s solution) LIVING MATERIAL Hydra culture Artemia larvae (optional)

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(a)

(b)

(c)

(d)

(e)

(f)

FIGURE 17.2 Some suggestions for planarian regeneration experiment (dashed lines represent cuts). In experiments involving partial separation, cutting will need to be repeated several times to prevent the parts from healing together.

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PLANARIANS Planarians are free-living, freshwater flatworms that usually are found in relatively clean, unpolluted water. They often inhabit small, shallow, slow-moving streams where they live under stones or leaves. Planarian worms have long been favored subjects in regeneration research. Even small fragments are capable of regenerating parts to complete the characteristic planarian body form with a head possessing two light-sensitive eyespots and a pair of lateral protuberances, called auricles, at one end and a tail that tapers to a point at the other. Regeneration at a cut surface is characterized by formation of a regeneration blastema, a region of cell division and progressive differentiation. Neoblasts, members of a population of “reserve cells” that apparently retain embryonic characteristics, gather in a forming blastema.The roles and relative importance of neoblasts in the regeneration process have been investigated and debated for years but are still not completely established. Polarity and other factors that guide the regeneration process have also been intensively studied in planarians. In this experiment, you will have opportunity to make basic observations of planarian regeneration and to investigate some aspects of the control of regenerative processes. Techniques 1. Choose a set of regeneration experiments from the experiments suggested in fig. 17.2. Your instructor may make some specific suggestions. 2. Prepare and label a set of small petri dishes containing spring water or dilute Amphibian Ringer’s solution to hold fragments during the regeneration process. 3. Obtain a petri dish containing ice and set it upside down on the stage of a dissecting microscope. Place a drop of spring water at the center of the bottom of the dish. 4. Use a pipette to transfer a planarian to the drop on the plate surface. 5. Wait until the worm is relaxed and spread on the dish and then quickly cut it with the blade provided. If you are making more than one cut, you may need to wait for the worm to relax before cutting it again. 6. Wash or brush the parts off the cutting surface into one of the petri dishes that you prepared earlier and place the dish in a cool, dark place. 7. Repeat Steps 4 to 6 for each animal to be cut. 8. In experiments in which worms are split, but not completely divided, it will probably be necessary to recut the worms several times during the first 2 or 3 days since the cut parts tend to heal together.This is especially true of experiments on head duplication. 9. Replace the solution in which the worms are regenerating every 2 or 3 days. Remove and discard any dead fragments that you find when you change the solution. 10. Do not feed the animals during regeneration.

Observations 1. Note movement of cut worms or worm fragments at various time intervals after cutting. 2. Watch for, and record, the time of development of a blastema at each cut surface. Usually, a blastema can be easily recognized because it has less pigment than regular body parts. 3. Record changes in the blastemas and any expansion of the regenerating parts. 4. How much time is required for recognizable new body parts, such as eyespots or auricles, to be formed? 5. When do regenerated parts become pigmented? 6. Are there any unusual results? That is, do any unexpected or “misplaced” body parts form? 7. Consult classmates and try to draw some general conclusions about survival rates and regenerative success following various experimental manipulations.

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Materials EQUIPMENT Dissecting microscope Single-edged razor blades or curved scalpel blades Pipettes (with wider mouth than Pasteur pipettes) Small covered dishes (for example, 60-mm petri dishes) SOLUTIONS

AND

CHEMICALS

Petri dish halves filled with ice (pour water into dishes, freeze, and keep frozen until needed) Spring water (or 10% Amphibian Ringer’s solution) LIVING MATERIAL Planaria

LUMBRICULUS Lumbriculus variegatus is an aquatic oligochaete annelid that superficially resembles an earthworm, though considerably smaller. Lumbriculus is particularly interesting because of its very strong regenerative capacity. This ability comes into play when body parts are lost, but it also is important in asexual reproduction. Lumbriculus individuals reproduce by fission. They break in half, and each half regenerates appropriate parts and produces a normal, complete body. Under some circumstances, Lumbriculus appears to reproduce primarily, or even exclusively, by fission. Under experimental conditions, it is relatively easy to demonstrate that even a very small fraction of a Lumbriculus body can regenerate and reorganize to form a complete worm. This spectacular regeneration from only a few segments necessarily involves morphallaxis.Morphallaxis is a technical name for conversion of the previously existing body part into a quite different part appropriate to the position that it assumes in the body of the newly organized individual. Morphallaxis during Lumbriculus regeneration entails very substantial structural and functional reorganization of the original segments in relation to the new segments that are regenerated anterior and posterior to them. In this experiment, you will have opportunity to examine Lumbriculus regeneration and to make some general observations of the phenomenon of morphallaxis. Techniques 1. Plan experiments to analyze regeneration by small fragments (about 20 to 30 segments) taken from various parts of the Lumbriculus body. It is often convenient to consider thirds of the worm so that you could examine regeneration of fragments taken from the first, second, or third portion of the body. 2. Place a piece of filter paper in a petri dish and soak it with distilled water. 3. Use a wide-mouth pipette to transfer a worm to the surface of the wet filter paper. 4. Use an iris scissors to make the necessary cuts in each worm body to obtain appropriate fragments for your experiment (please be very careful in handling the scissors to avoid damage to its sharp, but rather fragile blades). If you have difficulty making cuts because the worm is moving too actively, place the filter paper on the bottom of an inverted petri dish containing ice. Chilling should slow the worm’s movements. Make certain that all of the fragments have a cut surface at each end. Fragments containing the first or last segment of the body will produce results that complicate interpretations considerably. 5. Add enough distilled water to make it possible to use a pipette to pick up and transfer each fragment to a dish containing spring water or dilute Amphibian Ringer’s solution. Mark these dishes carefully with information about the third of the body represented and the approximate number of segments in the fragment or assign a specimen number relating to this information in your lab notebook.

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Observations 1. Make note of the amount of bleeding that occurs when each worm is cut. Observe movement by the fragments at various times after cutting and during the regeneration process. 2. Watch for evidence of new growth at each cut surface (new tissue will differ in color from the original fragment). 3. As new growth proceeds, look for signs of segment differentiation. Is segmentation evident from the beginning of new growth? Are new segments added one at a time? 4. Are there differences between anterior and posterior ends in terms of regeneration patterns? 5. How many new segments eventually form at each end of each fragment? Are there relationships between the numbers of new anterior or posterior segments and source (that is, body third) of the fragment? 6. Your instructor may suggest that you incorporate your data and the class data into a summarizing model of Lumbriculus regeneration.

Materials EQUIPMENT Dissecting microscope Pipettes (with wider mouth than Pasteur pipettes) Small covered dishes (for example, 60-mm petri dishes) Standard (100-mm or 90-mm) petri dishes Bowl or dish to hold ice water for chilling Filter paper Iris scissors (iridectomy scissors) SOLUTIONS

AND

CHEMICALS

Distilled water in squirt bottle Spring water (or 10% Amphibian Ringer’s solution) Petri dish halves filled with ice (pour water into dishes, freeze, and keep frozen until needed) LIVING MATERIAL Lubriculus culture

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18. Experimenting with Vascular Plant Development Patterns


L A B O R A T O R Y

18 Experimenting with Vascular Plant Development Patterns

Several special features of development in vascular plants are of particular interest to developmental biologists, and they present striking contrasts to familiar patterns of animal development. Plants have meristematic regions that are specific, localized areas where active cell division continues through the growing season. In a perennial plant, growth can continue for years, ending only at the death of the plant. Thus, the size of such a plant is indeterminate; the plant does not stop growing at a genetically determined size as do the vast majority of animals. Morphogenetic movement, which rearranges relative positions of various cells during animal development, does not occur in the development of vascular plants. Patterns of growth and differentiation in plants depend upon specifically oriented cell division planes and differential cell enlargement along specific axes. For example, many cells lengthen without substantial increases in girth. Eventually, complex structural arrangements develop with striking patterns of diverse cell differentiation associated with the cell’s relative position in the plant’s growing region. Because a plant is not free to move about, its entire life is a series of interactions with the environment in a single location. Plants are able to respond to physical factors such as gravity, light, and water availability and to make adjustments following damage or loss of body parts. All of these environmental interactions involve developmental processes of growth and differentiation. In this laboratory, you will have opportunities to observe and experiment with several features of vascular plant development, including cell elongation, meristematic regions and progressive differentiation, response to a physical environmental factor (gravity), and responses to loss of a body part.

POLLEN TUBE GROWTH IN VITRO Although pollen tube growth is a somewhat special case, it can, nevertheless, serve as a model for the cell enlargement along a single axis that plays such an important role in plant morphogenesis. After pollen grains land on the stigma, pollen tubes grow down through the style portion of the pistil to the ovary where they deliver sperm nuclei in the vicinity of female gametophytes inside the ovules. This pollen tube elongation is a late phase in male gametophyte development in flowering plants. Pollen grains of a number of species will germinate in vitro, and pollen tube elongation occurs rapidly enough to be observed during a single laboratory period. This in vitro growth can be spectacular in some cases, but tubes normally do not grow to be as long in vitro as they do inside the pistil of a flower of their own species.

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individual root hair region of maturation

region of elongation

apical meristematic region

root cap

FIGURE 18.1 Longitudinal section of a root tip. Cell detail shows progressive cell specialization in cells further from apical meristem. Root hairs develop as extensions of epidermal cells.

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Techniques 1. Place a folded piece of wet paper towel in a petri dish. This wet towel will keep the humidity high inside the dish and prevent evaporation of medium during pollen tube growth. 2. Place several drops of growth medium in the concavity of a depression slide and scrape pollen off an anther into the medium. Make note of the time. 3. If you are comparing pollen tube growth rates of several species, repeat Steps 1 and 2 for each type of flower. 4. Place each depression slide on a wet towel in a petri dish and add a cover to each dish.

Observations 1. Beginning about one-half hour after you have set up your culture slides, check each culture periodically for signs of pollen tube growth. The tube will first appear as a small translucent bulge on the side of a pollen grain. Pollen tube observations will be made easier if you use dark-field or phase-contrast microscopy. If you have neither available, stop down the iris diaphragm of your microscope’s condenser to increase contrast. Scan several fields of view each time you observe a culture because there may be variations in germination or tube growth in different areas of the culture. 2. Record data on the time of pollen tube emergence for each type of pollen that you have in culture. 3. Continue your observations by making observations or measurements of tube growth at regular intervals for several hours. You can make quantitative and graphical comparison of the growth rates of different species’ pollen tubes under these culture conditions.

Materials EQUIPMENT Compound microscope Depression slides Petri dishes for moist chambers SOLUTIONS

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CHEMICALS

Pollen germination medium LIVING MATERIAL Flowers with mature pollen on their anthers

MAPPING ROOT GROWTH AND DIFFERENTIATION Growth and differentiation at the tips (apices) of shoots and roots is a continuing process throughout the growing season. Near a root’s tip is a meristematic region, an area of active cell division. Some of the cells produced in this apical meristematic region are added to the root cap, a protective cover over the root tip. Root cap cells are ground off by the wear and tear of being pushed through spaces among rough soil particles and must be constantly replaced. Other cells produced by divisions in the meristematic region add to the length of the growing root. In a longitudinal section through a root tip, you can see regional differences among cells at various distances from the meristematic zone (fig. 18.1). Cells in and near the meristematic region are small, cuboidal, and fairly homogeneous in appearance, while those further from the center of the meristematic region are longer.These longer cells are in a zone of cell growth called the region of elongation. Although the meristematic region and the region of elongation take up only a few millimeters of a root’s total length, cell division and cell lengthening in these areas account entirely for the lengthening of the growing tip that pushes the root through the soil.

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FIGURE 18.2 Examples of the patterned thickenings (ornamentation) seen in the walls of differentiated xylem vessel cells.


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Further back from the tip, above the region of elongation, is a region where cell differentiation is establishing the structural pattern and functional relationships essential for specialized functioning of the root, the region of maturation. In the region of maturation, some epidermal cells produce root hairs, while cells in specific areas of the root interior produce cortex, endodermis, and elements of vascular tissue. Cellular aspects of this pattern of division, elongation, and differentiation can be examined in roots produced on cuttings of the common houseplant Zebrina pendula, called variously, the zebra plant, Wandering Jew, or simply, Zebrina. Roots grow out of the stem in a Zebrina cutting. Roots of this type that grow out of the stem after the plant’s basic root system has developed are called adventitious roots. Techniques 1. Remove a Zebrina root from the stem of a cutting and place it in a drop of water on a slide. 2. Cut three 2-mm pieces, beginning at the root tip end. Separate the three pieces so that you can keep track of which is which. 3. Keep them wet until you are ready to make a stained squash of each fragment using the Nickolas technique given on p. 232 in Appendix F.

Observations 1. Examine the squash of each fragment, in turn, to assess what percentage of it, if any, appears to be made up of the meristematic region and root cap, the region of elongation, and the region of maturation. Make estimates of the proportion of each squash that is occupied by cells characteristic of those root regions (see a–c following). a. The meristematic region is clearly identifiable because of the presence of dividing cells. Scan areas of your squash of the terminal 2-mm fragment for cuboidal cells. Then go to high power and look for cells in various mitotic stages. Root cap cells are larger, irregularly shaped cells. For purposes of mapping, you can lump meristematic regions and root cap into a single terminal region. b. The region of maturation is best identified in squashes by the presence of differentiated xylem vessel cells with their characteristically ornamented walls (fig. 18.2).When such cells are present, you may conclude that you are viewing tissue from the region of maturation. c. The region of elongation that lies between the other two contains elongated cells whose length is several times their width. To be classified as region of elongation in our scheme, however, it should contain no differentiated xylem cells. 2. Assume that each fragment is, in fact, 2 mm long and relate the proportions determined in Step 1 to the lengths of the cut fragments in order to map the growing end of the root. In other words, if it appears that about one-half of the third 2-mm fragment is assignable to the region of maturation, you can conclude that the region of maturation begins about 5 mm from the tip of the root. 3. You should squash, examine, and map several Zebrina roots. How do your results compare with those of classmates? How might the precision of these estimates be improved? 4. Your instructor might ask you to observe roots of several other species for purposes of comparison. In some species, the presence of root hairs is a clear external marker of the region of maturation. Your instructor might also suggest that you cut sections of some roots and stain them, using the techniques on p. 233 of Appendix F. In the cases of larger roots, you can also cut sections out of the root at various levels and squash the sections (using the same squash technique applied to root tip fragments) to determine what cell types are present.

Materials EQUIPMENT Compound microscope Single-edged razor blades Rulers Forceps Hot plate 100-ml beakers Slides and coverslips

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masking tape inverted petri dish bottom

seedling

petri dish cover wet paper towel

FIGURE 18.3 Petri dish container for seedlings in root growth experiments.


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SOLUTIONS

AND




CHEMICALS

1N HCl in dropping bottles Toluidine Blue O stain (0.5% in water for squashes; 0.05% in water for staining sections) Boiling stones or chips Nail hardener or Liquid Bandage LIVING MATERIAL Rooted Zebrina pendula cuttings Germinated seeds of other plants with emergent seedling roots (optional)

GRAVITROPISM Gravitropism (also sometimes called geotropism) is a directional growth response in which gravity is the orienting factor. Gravitropism is very important in early seedling growth because it orients seedlings properly in the soil. Stems are negatively gravitropic; that is, they curve away from the earth’s gravity center. Roots are positively gravitropic, curving toward it. The mechanisms that link gravity sensing with the expression of the gravitropic response of a root are complex, but we will limit work in this lab to studying the nature and timing of the gravitropic response and to experimenting with roots deprived of their primary gravity sensing area, the root cap. Techniques 1. Obtain germinated sweet corn seeds with growing roots that are 5 to 10 mm long. 2. Lay out several seeds inside the top of a petri dish with the side of the seed where the root has emerged away from the glass surface. Arrange the seedlings so that their roots are parallel to one another. Place a folded, wet paper towel over the seeds and set the petri dish bottom inside the cover with its bottom against the wet paper towel (fig. 18.3). Use masking tape to secure the two halves of the dish so that there is very gentle pressure on the paper towel and the seeds beneath it. Inspect the seedlings. Measure the roots and record their lengths. Trace the roots’ shapes on the outer surface of the cover with a marking pencil. (If the seeds do not seem stable in their positions, uncover them and attach them to the cover with a small piece of modeling clay before replacing the paper towel and dish bottom.) 3. Place the dishes in containers that will hold them upright on their sides.Add a small amount of water to these containers to prevent drying. Set the petri dishes so that the roots are horizontal, that is, their long axis is perpendicular to gravitational force. Put the dishes in a dark place. 4. Attempt to demonstrate experimentally that the gravity-detecting region of a root is separate from the root regions responsible for lengthening. The role of the root cap can be tested by cutting it off. In most roots, root cap tissue appears to have a different density than other root tissue when viewed with a dissecting microscope.With careful work, you may be able to strip off most of the root cap. Make note of how much you removed. Measure each root and record its length. Make note of the amount of tip removed from each and mount them in petri dishes as you did with the uncut seedlings in Step 2. Trace the shapes of the roots on the outer surface of the covers and place the dishes in the dark as in Step 3.

Observations 1. How long does it take to develop detectable downward turning in control roots? 2. How much time is required for roots to turn 90° and point straight down? 3. How much do roots increase in length during this gravitropic turning? 4. What region of the root actually curves? The meristematic region? The region of elongation? The region of maturation? If you are not able to draw conclusions about this, how might you obtain additional relevant data? 5. Do any or all of the roots with their root caps removed turn downward? Do they increase in length? Did they lengthen as much as the uncut seedlings did in the same time? Would you conclude that you have removed the meristematic region in some cases? How might you obtain data relevant to that question? 6. When you have gathered your basic data on gravitropism, you could extend your experiment by rotating uncut seedlings another 90° relative to gravity. Can you eventually cause roots to grow in a circle by repeated rotation relative to gravity?

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Materials EQUIPMENT Dissecting microscope Petri dishes Single-edged razor blades Forceps Marking pencils that will write on plastic or glass Rulers SOLUTIONS

AND

CHEMICALS

Modeling clay LIVING MATERIAL Geminated sweet corn seeds with 5 to 10-mm-long emergent seedling roots Germinated seeds of other species (optional)

ROOT REGENERATION RESPONSES Replacement of lost parts is a critical part of a plant’s interactions with its environment.Replacement of aboveground parts is a familiar process, for example, in plants eaten by herbivores (or subjected to a lawn mower). Plants also have the capacity to repair damage to roots and to replace lost root portions, but the replacement part or parts may differ from the original root. In this experiment, you will investigate the patterns established in roots after removal of various portions of their growing tip. Techniques 1. Prepare the number of liquid culture jars that you will need for your experiment.A liquid culture arrangement consists of a small jar wrapped with aluminum foil that is topped by a piece of fiberglass screen.The screen should have a 1-cm slit in its center where the seedling will be mounted.After you have pushed the center of the screen down about 1 cm below the rim of the jar, attach the screen to the jar with a rubber band around the lip of the jar. Make sure that the distinct concavity formed when you pushed the screen down is maintained as you apply the rubber band.This arrangement will allow you to keep the culture medium level up to the seed without filling the jar to the brim. Fill each jar with medium to the level of the slit in the center of the screen. 2. Obtain germinated pea (Early Alaska variety) seedlings with roots 35 to 45 mm long. 3. Cut off root tips as follows: Cut a tiny piece (less than 0.5 mm) off one root; cut between 1 mm and 2 mm off a second root; and finally, cut 4 to 5 mm off a third. After cutting, carefully measure the length of each root and record the data. 4. Transfer each cut pea seedling to a liquid culture jar. Part the screen at the slit as you insert the root to avoid damage to delicate root tissue. Make certain that the root is totally immersed in the medium. 5. Check the level of medium in your jars each day. Keep the culture level up to the bottom of the seed so that the root is completely immersed. 6. Place a seedling with an uncut root (control) in culture using exactly the same procedures. 7. If plants of another species are available, set up the same experiment with that plant.

Observations 1. Unless you are asked to make interim observations, examine your cultures after 7 days in culture. 2. Determine how each root has responded following cutting. Did growth resume at the cut surface of the root? Did this remaining tip split into several parts? Did new lateral branch roots develop that have no counterparts on control plants? At what angles do the lateral branches grow out from the original root? 3. Compare growth, in length, of the various experimental plants to that of the control plant. 4. Consult your instructor about using class data to draw conclusions about the pattern of root regeneration. 5. If you worked with a second species, collect data for it as in Steps 1–4.

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Materials EQUIPMENT Dissecting microscope Ruler Forceps Single-edged razor blades Small jars such as baby-food jars Fiberglass screening cut into 15 ⫻ 15 cm squares (approximate) with an approximately 1-cm slit cut in the center Aluminum foil SOLUTIONS

AND

CHEMICALS

Fern medium (or other appropriate plant culture medium) LIVING MATERIAL Germinated pea (preferably Early Alaska variety) seeds with 35 to 45-mm-long emergent seedling roots Germinated seeds of other species (optional)

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Suggestions: Further Invest. of Emerging & Regenerating Developmental Patterns


SUGGESTIONS FOR FURTHER INVESTIGATION OF EMERGING AND REGENERATING DEVELOPMENTAL PATTERNS Spirostomum Ciliated protozoans have long been subjects of cellular morphogenesis investigations (for example, Tartar 1961). The genetic control of pattern formation, particularly patterns of surface organelles, have been extensively researched (Nanney 1975; Sonneborn 1975;Aufderheide et al 1980). Research on internal details of cytoskeletal (Lynn 1981) and cortical (de Terra 1981) organization has shed further light on cellular patterns. If you would like to learn more about the current status of research on cell organization patterns and morphogenesis in these interesting and beautiful organisms or would like to experiment further with ciliate cells, consult reviews by Frankel (Frankel 1984 and 1989). Hydra A great deal of the fascinating, two hundred and fifty year history of research on hydra is discussed in publications by Howard and Sylvia Lenhoff.Their descriptions of the extensive and insightful research of the eighteenth century Swiss scientist Abraham Trembley are particularly interesting. Howard Lenhoff (1991) has also investigated the role that the work on hydra by Ethel Browne (Harvey) might have played in the development of Hans Spemann’s concept of embryonic induction (see p. 89). This is an intriguing bit of developmental biology history that you might enjoy reading.You can obtain some basic information about hydra reproduction and development in Grassi et al (1997) and Martin et al (1997). Some research on hydra pattern formation and regeneration has involved treatments that selectively destroy certain types of cells, such as interstitial cells (Diehl and Burnett 1964 and 1965) or nerve cells (Marcum et al 1977). While some of these cell depletion experiments involve use of very dangerous substances and are not suitable for student laboratories, others might be feasible if you wished to attempt them. If you do wish to undertake extensions of these or other studies on hydra, consult the papers by Grimmelikhuijzen and Schaller (1979), Lesh-Laurie (1982), Bode and Bode (1984), Technau and Holstein (1995) as well as Howard Lenhoff’s (1983) collection of methods for hydra research. One of the interesting themes of hydra research in the latter part of the twentieth century has been the role of diffusible regulators of morphogenetic events in hydra (Schaller et al 1989; Hobmayer et al 1990; Müller 1991; Javois and Frazier-Edwards 1991; Shimuzu et al 1993; Hassel et al 1993). Some of these regulators diffuse through intercellular spaces, but others diffuse via the cytoplasmic route through gap junctions (Fraser et al 1987). It appears that hydra will continue to be an important subject for investigation of diffusible morphogens (Wolpert 1991) in development. Hydra also has been subjected to dissociation and reaggregation experiments and has shown remarkable abilities to reconstitute complex forms after complete dispersal (Gierer et al 1972; Sato and Sawada 1989; Sato et al 1992;Technau and Holstein 1993; Lee and Javois 1993; Sato-Maeda and Tashiro 1999), and it might be feasible for you to replicate or extend some of these experiments. The role of environmental factors such as carbon dioxide tension in regulating hydra sexual reproduction has been studied by W. H. Loomis and others and continues to be the subject of active investigation (for example, Littlefield et al 1991). Thus, hydra research is also an important part of the study of general invertebrate reproductive biology. The cell biology of hydra morphogenesis is also a continuing active field of research (Teragawa and Bode 1990; Littlefield 1991). Before you begin additional work with hydra development, it would be helpful to consult Javois (1992) for a summary of hydra research up to the early 1990s. Planaria Regeneration of planarian worms is spectacular and relatively easy to observe. It is not surprising, therefore, that planarian regeneration has been extensively investigated for many years (Brønsted 1969). These regeneration studies have contributed to the understanding of repatterning processes necessary for reestablishment of substantial lost body portions of a relatively simply organized animal. They also Suggestions

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are of interest from a comparative viewpoint. The planarian regeneration blastema has no counterpart in hydra regeneration, but it does play a role similar to that of the blastema that functions in larval amphibian limb regeneration. Interpretations of the results of planarian regeneration experiments were instrumental in development of the concept of morphogenetic gradients by C. M. Child (1941) and his contemporaries.The gradient concept has been modified substantially, but it contributed to the development of the modern morphogen hypothesis (Slack 1987) and represents an important milestone in the history of developmental biology. Modern research on planarian regeneration includes work focused both on details of specific body structure regeneration (Asai 1991; Kurabuchi and Kishida 1992; Reuter et al 1996) and on morphometric (Mead 1991), cell biological (Martelly and Franquinet 1985), and genetic (Baguna et al 1994; Bayascas et al 1997; Bogdanova et al 1998; Bayascas et al 1998; Pineda et al 2000) aspects of regeneration. If you would like to do more experiments on planarian regeneration, you would do well to consult Hamburger (1960), where a number of regeneration experiments are suggested. Patterns of cell proliferation during regeneration could be investigated with the technique of Redi, Garagna, and Pellicciari (1982) or one of the earlier methods they cite. Lumbriculus It is important to note that Lumbriculus, an organism that displays interesting physiological and behavioral responses ( Lesiuk and Drewes 1999; Drewes and Cain 1999), routinely reproduces by fission and the subsequent regrowth of appropriate parts by the separated body halves. Thus, regeneration is not just a possible response to occasional injury, but part of a regularly employed reproductive strategy. Morgulis reported extensive investigations of Lumbriculus regeneration in 1907, and research on annelid regeneration continued throughout the first two-thirds of the twentieth century (Turner 1934; Berrill 1962; Herlant-Meewis 1964).These studies highlighted the general importance of morphological morphallaxis (p. 181) in regeneration by many organisms. Recently, Charlie Drewes and his colleagues have focused on functional morphallaxis in the reorganization of the regenerating nervous system.You could consult their papers in order to learn more about this very interesting organism and the spectacular reorganization that it undergoes during regeneration. Plant Patterns Further experiments with pollen germination could include comparisons of responses of various kinds of flower pollen and tests of the effects of modifying the ionic composition of the medium. For more information on basic techniques, consult Motten (1992), Schimpf (1992), and Tatina and Hohn (1994). Heslop-Harrison (1982), Chasan (1992), and Cheung (1995, 1996) provide information on specificity of pollen-stigma interactions and Palser et al (1992), Wang et al (1989), and Ray et al (1997) present data on the guidance of growing pollen tubes. There are a variety of sources of general information on plant growth and differentiation patterns (for example, Wareing and Phillips 1981; Steeves and Sussex 1989; Stebbins 1992; Burgess 1985; Sachs 1984) that you could consult, and Hershey (1994) describes simple and inexpensive hydroponics techniques. There have been a number of studies of gravitropism in corn (for example, Bostock et al 1991, Nantawisarakul and Newman 1992) that might provide ideas for further investigation (see also Haldeman and Gray 2000).You might also want to read articles on general principles of gravitropism (Moore 1984; Evans et al 1986) or on research on the mechanism of gravitropic bending (Cosgrove 1990; Barlow et al 1993; Moore and Maimon 1993). The gravitropic response has been investigated in a variety of systems, one of which you might find easy to study. Dandelion stems show a strong negative gravitropic response, and it is possible to investigate the cellular events that trigger this response with common lab instruments and a light microscope (Clifford and Oxlade 1991; Barclay and Clifford 1991). There are many potential variations on the basic experiments they suggest, and you could pursue them with ease and practically no expense during seasons when dandelions are flowering. As you work on gravi194

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tropism, however, you should be aware that there still are questions to be resolved regarding the sensing and signaling mechanisms in gravitropism (Salisbury 1993; Barlow 1995; Sack 1997; Kiss et al 1997; Rosen et al 1999).You can get some ideas for further investigation of gravitropism using Aradopsis mutants from Kiss et al (2000). There also has been some very interesting research on thermotropism, the oriented growth responses to temperature, by corn roots (Fortin and Poff 1990, 1991; Day 1991). It might be a little difficult to replicate or extend these experiments, but it could prove very interesting to do so. In this connection, it might also be interesting to look into direct temperature effects on corn root morphogenesis (Kiel and Stamp 1992). Reihman and Rost (1990) provide excellent data on regeneration responses in pea roots, as well as some references to other works on root regeneration responses. If you should wish to extend your basic experiments on root regeneration responses, you might consider a comparison of vascular bundle patterns of control roots to those of regenerated roots. Fuchs (1963) gives details of the technique for visualizing vascular bundles in roots used by Reihman and Rost (1990) that you could employ. If you have access to a fluorescence microscopy system, you might use the fluorescence technique, which can yield absolutely beautiful results (Hughes and McCully 1975; Gates 1991), to study sections of both control and regenerated roots. Regenerating squash roots show some very interesting responses to hormone treatments (Bohnsack 1989), in which calluslike growths develop at the ends of root branches. This callus formation is responsive to hormone balance, so you could undertake some kinds of experiments with this system that ordinarily would require complex tissue culture techniques. The experiments that Bohnsack describes can be done with roots growing in fern medium (p. 228). Pumpkin roots also give excellent results. For more ideas for experiments with developing plants, consult the collection of experimental approaches to plant development compiled by Singer (1992). Developmental Pattern and Regeneration References General Sources Cordero, R. E.The developmental importance of cell division. Amer. Biol. Teacher 56:176–179; 1994. Dinsmore, C. E. Animal regeneration—from fact to concept. BioScience 45:484–492; 1995. Dinsmore C. E., ed. A history of regeneration research. Milestones in the evolution of a science. New York: Cambridge Univ. Press; 1991. Jurgens, G.; Mayer, U. Arabidopsis. In: Bard, J. B. L., ed. Embryos: color atlas of development. London:Wolfe; 1994: 7–21. Malacinski, G. M.; Bryant, S. V., eds. Pattern formation: a primer in developmental biology. New York: Macmillan; 1984. Meyerowitz, E. M. Plants and the logic of development. Genetics 145:5–9; 1997. Rossomando, E. F.; Alexander, S., eds. Morphogenesis. New York: Marcel Dekker; 1992. Singer, S. R. Plant life cycles and angiosperm development. In: Gilbert, S. F.; Raunio, A. M., eds. Embryology: constructing the organism. Sunderland, MA: Sinauer; 1997:493–513. Stebbins, G. L. Comparative aspects of plant morphogenesis: a cellular, molecular, and evolutionary approach. Amer. Jour. Botany 79:589–598; 1992. Steeves,T. A.; Sussex, I. M. Patterns in plant development. 2d ed. Cambridge: Cambridge Univ. Press; 1989. Stocum, D. L.; Karr,T. L., eds. The cellular and molecular biology of pattern formation. Oxford: Oxford Univ. Press; 1990. Tsonis, P. A. Regeneration in vertebrates. Dev. Biol. 221:273–284; 2000. Wilkins, M. Plantwatching: how plants remember, tell time, form partnerships and more. New York: Facts on File; 1988. Wolpert, L. The triumph of the embryo. Oxford: Oxford Univ. Press; 1991.

Reviews and Research Papers SPIROSTOMUM Aufderheide, K. J.; Framel, J.; Nelson, E. M. Formation and positioning of surface-related structures in protozoa. Microbiol. Rev. 44:252–302; 1980.

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de Terra, N. Cortical control of macronuclear positioning in Stentor. Jour. Exp. Zool. 216:367–376; 1981. Frankel, J. Pattern formation in ciliated protozoans. In: Malacinski, G. M.; Bryant, S. V., eds. Pattern formation: a primer in developmental biology. New York: Macmillan; 1984:163–196. Frankel, J. Pattern formation: ciliate studies and models. New York: Oxford Univ. Press; 1989. Lynn, D. The organization and evolution of microtubular organelles in ciliated protozoa. Biol. Rev. 56:243–292; 1981. Nanny, D. L. Patterns of basal body addition in ciliary rows of Tetrahymena. J. Cell. Biol. 65:503–512; 1975. Sonneborn,T. M. Positional information and nearest neighbor interactions in relation to spatial patterns in ciliates. Ann. Biol. 14:565–584; 1975. Tartar, V. The biology of Stentor. London: Pergamon Press; 1961. HYDRA Bode, P. M.; Bode, H. R. Patterning in hydra. In: Malacinski, G. M.; Bryant, S. V., eds. Pattern formation: a primer in developmental biology. New York: Macmillan; 1984:213–240. Diehl, F. A.; Burnett, A. L. The role of interstitial cells in the maintenance of hydra. I. Specific destruction of interstitial cells in normal, asexual, non-budding animals. J. Exp. Zool. 155:253–260; 1964. Diehl, F. A.; Burnett, A. L. The role of interstitial cells in the maintenance of hydra. III. Regeneration of hypostome and tentacles. J. Exp. Zool. 155:299–318; 1965. Fraser, S. E.; Green, C. R.; Bode, H. R.; Gilula, N. B. Selective disruption of gap junctional communication interferes with a patterning process in hydra. Science 237:49–55; 1987. Gierer,A.; Berking, S.; Bode, H.; David, C. N.; Flick, K.; Hansmann, G.; Schaller, H.;Trenkner, E. Regeneration of hydra from reaggregated cells. Nature New Biology 239:98–101; 1972. Grassi, M.;Tardent, R.;Tardent, P. Quantitative data about gametogenesis and embryonic development in Hydra vulgaris Pall. (Cnidaria, Hydrozoa). Invert. Reprod. Dev. 27:219–232; 1995. Grimmelikhuijzen, C. J. P.; Schaller, H. C. Hydra as a model organism for the study of morphogenesis. Trends in Biochem. Sci. December: 265–267; 1979. Hassel, M.;Albert, K.; Hofheinz, S. Pattern formation in Hydra vulgaris is controlled by lithium-sensitive processes. Dev. Biol. 156:362–371; 1993. Hobmayer, B.; Holstein,T. W.; David, C. N. Stimulation of tentacle and bud formation by the neuropeptide head activator in Hydra magnipapillata. Dev. Biol. 183:1–8; 1997. Hobmayer, E.; Holstein,T.W.; David, C. N.Tentacle morphogenesis in hydra. I.The role of head activator. Development 109:887–895; 1990. Javois, L. C. Biological features and morphogenesis of Hydra. In: Rossomando, E. F.;Alexander, S., eds. Morphogenesis. New York: Marcel Dekker; 1992. Javois, L. C.; Frazier-Edwards, A. M. Simultaneous effects of head activator on the dynamics of apical and basal regeneration in Hydra vulgaris (formerly Hydra attenuata). Dev. Biol. 144:78–85; 1991. Lee, P.-C.; Javois, L. C. Patterning of heads and feet during regeneration of Hydra oligactis aggregates. Dev. Biol. 157:10–18; 1993. Lenhoff, H. M. Hydra: Research methods. New York: Plenum; 1983. Lenhoff, H. M. Ethel Browne, Hans Spemann, and the discovery of the organizer phenomenon. Biol. Bull. 181:72–80; 1991. Lenhoff, H.; Lenhoff, S.Tissue grafting in animals: its discovery in 1742, by Abraham Trembley as he experimented with hydra. Biol. Bull. 166:1–10; 1984. Lenhoff, H. M.; Lenhoff, S. G.Trembley’s polyps. Scientific American. May: 108–113; 1988. Lenhoff, H. M.; Lenhoff, S. G. Challenge to the specialist: Abraham Trembley’s approach to research on the organism—1744 and today. Amer. Zool. 29:1105–1117; 1989. Lesh-Laurie, G. E. Hydra. In: Harrison, F.W.; Cowden, R. R., eds. Developmental biology of freshwater invertebrates. New York: Alan R. Liss; 1982: 69–127. Littlefield, C. L. Cell lineages in Hydra: isolation and characterization of an interstitial stem cell restricted to egg production in Hydra oligactis. Dev. Biol. 143:378–388; 1991. Littlefield, C. L.; Finkemeier, C.; Bode, H. R. Spermatogenesis in Hydra oligactis. II. How temperature controls the reciprocity of sexual and asexual reproduction. Dev. Biol. 146:292–300; 1991. Marcum, B. A.; Campbell, R. D.; Romero, J. Polarity reversal in nerve-free Hydra. Science 197:771–772; 1977. Martin, V. J.; Littlefield, C. L.; Archer, W. A.; Bode, H. R. Embryogenesis in hydra. Biol. Bull. 192:345–363; 1997. Müller, W. A. Stimulation of head-specific nerve cell formation in Hydra by pulses of diacylglycerol. Dev. Biol. 147:460–463; 1991.

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Sato-Maeda, M.; Tashiro, H. Development of oriented motion in regenerating hydra cell aggregates. Zool. Sci. 16:327–334; 1999. Sato, M.; Sawada,Y. Regulation in the numbers of tentacles of aggregated Hydra cells. Dev. Biol. 133:119–127; 1989. Sato, M.;Tashiro, H.; Oikawa,A.; Sawada,Y. Patterning in Hydra cell aggregates without the sorting of cells from different axial origins. Dev. Biol. 151:111–116; 1992. Schaller, H. C.; Hoffmeister, S.A. H.; Dübel, S. Role of the neuropeptide head activator for growth and development in hydra and mammals. Development 1989 Suppl. 99–107; 1989. Shimizu, H.; Sawada,Y.; Sigiyama,T. Minimum tissue required for hydra regeneration. Dev. Biol. 155:287–296; 1993. Technau, U.; Holstein, T. W. Cell sorting during the regeneration of Hydra from reaggregated cells. Dev. Biol. 151:117–127; 1992. Technau, U.; Holstein, T. W. Head formation in Hydra is different at apical and basal levels. Development 121:1273–1282; 1995. Teragawa, C. K.; Bode, H. R. Spatial and temporal patterns of interstitial cell migration in Hydra vulgaris. Dev. Biol. 138:63–81; 1990. PLANARIA Agate, K., Watanabe, K. Molecular and cellular aspects of planarian regeneration. Seminars in Cell & Dev. Biol. 10:337–384; 1999. Asai, E. Regeneration of the pharynx in a freshwater planarian—An electron-microscopic study with special reference to the formation of the pharyngeal cavity and pharyngeal lumen. Zool. Sci. 8:775–784; 1991. Baguna, J.; Salo, E.; Romero, R.; Garcia-Fernandez, J., Bueno, J.; Munoz-Marmol, A. M.; Bayascas-Ramirez, J. R.; Casali,A. Regeneration and pattern formation in planarians: cells, molecules and genes. Zool. Sci. 11:781–795; 1994. Bayascas, J. R.; Castillo, E.; Munoz-Marmol, A. M.; Salo, E. Planarian Hox genes: novel patterns of expression during regeneration. Development 124:141–148; 1997. Bayascas, J. R.; Castillo, E.; Munoz-Marmol, A. M.; Salo, E. Platyhelminthes have a Hox code differentially activated during regeneration, with genes closely related to those of spiralian protostomes. Dev., Genes Evol. 208:467–473; 1998. Bogdanova, E.; Matz, M.; Tarabykin, V.; Usman, N.; Shagin, D.; Zaraisky, A.; Lukyanov, S. Inductive interactions regulating body patterning in planarian, revealed by analysis of expression of novel gene scarf. Dev. Biol. 194:172–181; 1998. Brønsted, H. V. Planarian regeneration. London: Pergamon; 1969. Ellis, C. H., Jr.; Fausto-Sterling, A. Platyhelminthes, the flatworms. In: Gilbert, S. F.; Raunio, A. M., eds. Embryology: constructing the organism. Sunderland, MA: Sinauer; 1997: 115–130. Kurabuchi, S.; Kishida,Y. Effect of delay in anterior or posterior amputation on regeneration of short fragments of planaria. Zool. Sci. 9:575–581; 1992. Martelly, I.; Franquinet, R. Planarian regeneration as a model for cellular activation studies. Trends in Biochem. Sci. 9:468–470; 1985. Mead. R. W. Proportioning of body regions in the planarian Dugesia tigrina as a function of the length:width ratio of the regenerating fragment. Jour. Exp. Zool. 259:69–77; 1991. Pineda, D.; Gonzalez, J.; Callaerts, P.; Ikeo, K.; Gehring, W. J.; Salo, E. Searching for the prototypic eye genetic network: Sine oculis is essential for eye regeneration in planarians. Proc. Natl. Acad. Sci. 97:4525–4529; 2000. Reuter, M.; Gustafsson, M. Neuronal signal substances in asexual multiplication and development in flatworms. Cell. Mol. Neurobiol. 16:591–616; 1996. Reuter, M.; Sheiman, I. M.; Gustafsson, M. K. S.; Halton, D.W.; Maule,A. G.; Shaw, C. Development of the nervous system in Dugesia tigrina during regeneration after fission and decapitation. Invert. Reprod. Dev. 29:199–211; 1996. Romero, R.; Baguna, J.; Calow, P. Intraspecific variation in somatic cell turnover and regenerative rate in the freshwater planarian Dendrocoelum lacteum. Invert. Reprod. and Dev. 20:107–114; 1991. Slack, J. M. W. Morphogenetic gradients—past and present. Trends in Biochem. Sci. 12:200–204; 1987. LUMBRICULUS Berrill, N. J. Regeneration and budding in worms. Biol. Rev. 27:401–438; 1962. Drewes, C.; Cain, K. As the worm turns. Amer. Biol Teacher 61:438–442; 1999. Drewes, C. D.Tell-tail adaptations for respiration and rapid escape in a freshwater oligochaete (Lumbriculus variegatus Müll). Jour. Iowa Acad. Sci. 97:112–114; 1990.

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Drewes, C. D.; Fourtner, C. R. Hindsight and rapid escape in a freshwater oligochaete. Biol. Bull. 177:363–371; 1989. Drewes, C. D.; Fourtner, C. R. Morphallaxis in an aquatic oligochaete, Lumbriculus variegatus: reorganization of escape reflexes in regenerating body fragments. Dev. Biol. 138:94–103; 1990. Drewes, C. D.; Fourtner, C. R. Reorganization of escape reflexes during asexual fission in an aquatic oligochaete, Dero digitata. Jour. Exp. Zool. 260:170–180; 1991. Drewes, C. D.; Vining, E.P.; Zoran, M. J. Regeneration of rapid escape reflex pathways in earthworms. Amer. Zool. 28:1077–1089; 1988. Herlant-Meewis, H. Regeneration in annelids. In: Abercrombie, M.; Brachet, J., eds. Advances in morphogenesis; Vol. 4. New York: Academic Press; 1964: 155–215. Lesiuk, N. M.; Drewes, C. D. Blackworms, blood vessel pulsations and drug effects. Amer. Biol. Teacher 61:48–53; 1999. Morgulis, S. Observations and experiments on regeneration in Lumbriculus. Jour. Exp. Zool. 4:549–574; 1907. Turner, C. D.The effects of x-rays on posterior regeneration in Lumbriculus inconstans. Jour. Exp. Zool. 68:95–115; 1934. PLANT PATTERNS Barclay, G. F.; Clifford, P. E. Using light microscopy to study geotropism. Amer. Biol. Teacher 53:285–286; 1991. Barlow, P.W. Gravity perception in plants: a multiplicity of systems derived from evolution? Plant, Cell Environment 18:951–962; 1995. Barlow, P.W.; Parker, J. S.; Butler, R.; Brain, P. Gravitropism of primary roots of Zea mays L. at different displacement angles. Ann. Bot. 71:383–388; 1993. Bohnsack, C. W. Cytokinin induced cell division and differentiation using intact plants. Amer. Biol. Teacher 51:106–108; 1989. Bostock, R. M.; Kuc, J. A.; Laine, R. A. Geotropism in corn roots: evidence for its mediation by differential acid efflux. Science 212:70–71; 1991. Burgess, J. An introduction to plant cell development. Cambridge: Cambridge Univ. Press; 1985. Chasan, R. Racing pollen tubes. Plant Cell 4:747–750; 1992. Cheung, A.Y. Pollen-pistil interactions in compatible pollination. Proc. Natl. Acad. Sci. 92:3077–3080; 1995. Cheung, A.Y. Pollen-pistil interactions during pollen-tube growth. Trends Plant Sci. 1:45–51; 1996. Cosgrove, D. J. Gravitropism of cucumber hypocotyls: biophysical mechanism of altered growth. Plant Cell and Environment 13:235–241; 1990. Day, S. Rooting around for the right temperature. New Scientist (9 February): 29; 1991. Evans, M. L.; Moore, R.; Hasenstein, K.-H. How roots respond to gravity. Sci. Amer. December: 112–119; 1986. Fortin, M.-C. A.; Poff, K. L.Temperature sensing by primary roots of maize. Plant Physiol. 94:367–369; 1990. Fortin, M.-C. A.; Poff, K. L. Characterization of thermotropism in primary roots of maize: dependence on temperature and temperature gradient, and interaction with gravitropism. Planta 184:410–414; 1991. Fuchs, C. Fuchsin staining with NaOH clearing for lignified elements of whole plants or plant organs. Stain Technology 38:141–144; 1963. Gates, P.The plant anatomy light show. New Scientist (9 February): 42–43; 1991. Haldeman, J. H.; Gray, M. S. Experiments with corn to demonstrate plant growth and development. Amer. Biol. Teacher 62:297–302; 2000. Hershey, D. R. Solution culture hydroponics: history and inexpensive equipment. Amer. Biol. Teacher 56:111–118; 1994. Heslop-Harrison, J. Pollen-stigma interaction and cross-incompatibility in the grasses. Science 215:1358–1371; 1982. Hughes, J.; McCully, M. E. The use of an optical brightener in the study of plant structure. Stain Technology 50:319–329; 1975. Kiel, C.; Stamp, P. Internal root anatomy of maize seedlings (Zea mays L.) as influenced by temperature and genotype. Ann. Botany 70:125–128; 1992. Kiss, J. Z.; Guisinger, M. M.; Miller,A. J.; Stackhouse, K. S. Reduced gravitropism in hypocotyls of starch-deficient mutants of Arabidopsis. Plant Cell Physiol. 38:318–325; 1997. Kiss, J. Z.; Weise, S. E.; Kiss, H. G. How can plants tell which way is up? Amer. Biol. Teacher 62:59–63; 2000. Moore, R.; Maimon, E. Signal transmission during gravitropic curvature of primary roots of Zea mays. Plant Cell and Environment 16:105–108; 1993. Motten, A. F. A simplified experimental system for observing pollen tube growth in styles. Amer. Biol. Teacher 54:173–176; 1992. Nantawisarakul,T.; Newman, I.A. Growth and gravitropism of corn roots in solution. Plant Cell and Environment 15:693–701; 1992.

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Okada, K.; Shimura, Y. Aspects of recent developments in mutational studies of plant signaling pathways. Cell 70:369–372; 1992. Palser, B. F.; Rouse, J. L.; Williams, E. G. A scanning electron microscope study of pollen tube pathway in pistils of Rhododendron. Can. J. Bot. 70:1039–1059; 1992. Ray, S.; Park, S.-S.; Ray, A. Pollen tube guidance by the female gametophyte. Development 124:2489–2498; 1997. Reihman, M. A.; Rost, T. L. Regeneration responses in peas roots after tip excision at different levels. Amer. J. Bot. 77:1159–1167; 1990. Rosen, E.; Chen, R. J.; Masson, P. H. Root gravitropism: a complex response to a simple stimulus? Trends in Plant Science 10:407–412; 1999. Sachs,Tsvi. Controls of cell patterns in plants. In: Malacinski, G. M.; Bryant, S. V., eds. Pattern formation: a primer in developmental biology. New York, Macmillan, 1984: 367–391. Sack, F. D. Plastids and gravitropic sensing. Planta 203:S63–S68; 1997. Salisbury, F. D. Gravitropism: changing ideas. Horticultural Reviews 15:233–278; 1993. Schmipf, D. J. Rapid germination of pollen in vitro. Amer. Biol. Teacher 54:168–169; 1992. Singer, S. R. Plant development lab collection. Northfield, MN: Society for Developmental Biology; 1992. (Copies can be obtained for a modest copying and shipping charge from S. R. Singer, Biology Dept., Carleton College, Northfield, MN 55057; phone 507-663-4391.) Tatina, R.; Hohn, K. A technique for staining pollen nuclei. Amer. Biol. Teacher 56:174–175; 1994. Wang, C.; Rathore, K. S.; Robinson, K. R.The responses of pollen to applied electrical fields. Dev. Biol. 136:405–410; 1989. Wareing, P. F.; Phillips, I. D. J. Growth and differentiation in plants. 3d ed. Oxford: Pergamon: 1981.

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Appendices

A: Preparation of Materials for Experimentation on Sea Urchin Embryos


A P P E N D I X

A Preparation of Materials for Experimentation on Sea Urchin Embryos

Consult the Materials section at the end of each Laboratory for a list of materials needed for that lab. This appendix provides formulae and information on sources of organisms as well as information on acquiring or making some of the listed materials that are not standard stockroom items. Additional information on laboratory work with sea urchin embryos can be obtained from a number of sources, including Hinegardner (1967 ), Schroeder (1986 ), and Strathmann (1987 ), cited on page 29, and D. P. Costello and C. Henley, Methods for Obtaining and Handling Marine Eggs and Embryos, Woods Hole, MA: Marine Biological Laboratory, 1971.This has been updated and republished on the Web: Cohen,W. D. 1999. Egg characteristics and breeding season for Woods Hole species. In Biological Bulletin Compendia (Online): Biological Bulletin publications: www.mbl.edu/Biological Bulletin. A. Sea Urchins and Sand Dollars Used in Teaching Several species of sea urchins are commonly used in developmental biology teaching labs in North America, and each of them presents some advantages and some disadvantages. Possibly the most commonly used species has been the West Coast purple urchin, Strongylocentrotus purpuratus, but other species have become increasingly popular for use in teaching. S. purpuratus (average egg diameter about 80 ␮m) is ripe from December to April, with January and February usually being the best months for use. However, I have used S. purpuratus successfully during the first week of December a number of times and have had fair success as late as the final week in March. S. purpuratus develops best at temperatures several degrees cooler than room temperature. Temperatures around 15° C usually give good results, but a few embryos may develop into pluteus larvae at cool room temperatures. Another Strongylocentrotus species, S. droebachiensis (average egg diameter about 160 ␮m) that is found in northern areas along both coasts, is ripe during approximately the same months as S. purpuratus. S. droebachiensis is somewhat more difficult to use in teaching because it is a cold-water urchin and its embryos usually must be kept at temperatures no higher than 10–12° C for normal development to the pluteus stage.

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Strongylocentrotus species are shipped in seawater-soaked newspaper packed with ice bags or frozen picnic-cooler blocks. Adults can be kept in good condition for a day or two if they are refrigerated or placed in a cool to cold (e.g., 10° C) growth chamber in the styrofoam box in which they arrive.They usually don’t fare well in aquaria, even well-maintained, refrigerated ones. Lytechinus pictus (average egg diameter about 110 ␮m) is a Pacific urchin that has developmental temperature relationships similar to those of S. purpuratus, but its breeding season is from May to September, so it is useful mainly very late in the spring term and in summer courses. L. pictus has beautifully clear eggs and displays developmental stages and processes with text photo clarity. Lytechinus variegatus (average egg diameter about 100 to 105 ␮m) is a warmer-water urchin that gives great results at room temperature, but animals from the northern Gulf of Mexico breed May through September and sometimes into October and November. However, specially collected animals can be induced to shed in the winter. The Lytechinus species are shipped in plastic bags of seawater and should be transferred to covered finger bowls, small fish bowls, or small aquaria if they must be held for a day or two. Both species fare well in growth chambers between 10° and 15° C, but animals should be given time to come to temperature gradually if they have been shipped with ice. Arbacia punctulata (average egg diameter about 75 ␮m), the subject of many classic investigations, is a widely distributed species that has different breeding seasons in various parts of its range. Southern populations breed during the academic year (October to April), but animals around Cape Cod breed during the summer months ( June to August). Arbacia is shipped in plastic bags of seawater, and adults should be handled like the Lytechinus species. Arbacia is a great animal with which to work, but its eggs are heavily pigmented, making some observations a bit difficult for some students. Eucidaris tribuloides (average egg diameter about 95 ␮m) is a cidaroid urchin with relatively clear eggs, whose development differs slightly from the other urchins described here (see the “Suggestions for Further Investigation of Echinoderm Development” section). Eucidaris adults are shipped and handled like the Lytechinus species. The West Coast sand dollar (irregular urchin), Dendraster excentricus (average egg diameter about 115 to 120 ␮m), can be used satisfactorily from late spring through most of the summer.Adult sand dollars are generally hardier than regular urchins under laboratory conditions and can be maintained for several days at cool temperatures (about 15° C) if kept uncrowded in clear seawater. Offspring of D. excentricus from intertidal populations are very hardy and develop normally at room temperature. To inject a sand dollar with KCl, insert the needle at a very low angle very near the mouth. It may take injections in several directions to induce shedding, but the total volume of KCl required is less than that needed for most of the regular urchins. A common sand dollar of the northern Atlantic coast, Echinarachnius parma (average egg diameter about 140 ␮m), might be a good alternative choice in some cases. Its breeding season around Woods Hole is from late March to mid- or late July. E. parma embryos require cool temperatures for normal development. B. Commercial Sources of Sea Urchins and Sand Dollars The addresses of some of the established commercial collectors and/or suppliers of sea urchins and sand dollars follow. Gulf Specimen Marine Laboratories, Inc. P. O. Box 237 Panacea, FL 32346 (850) 984–5297 Arbacia punctulata (Oct–April) Lytechinus variegatus (May–Sept) Both species develop well at room temperature.

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Marinus, Inc. 1500 Pier C Street Long Beach, CA 90813 (562) 435–6522 Strongylocentrotus purpuratus (Dec–April) Lytechinus pictus (May–Sept) Dendraster excentricus (sand dollar) (May–July) S. purpuratus and L . pictus develop best at temperatures several degrees cooler than room temperature, but some embryos will develop normally at cool room temperatures. Dendraster embryos develop at room temperature. Pacific Bio-Marine P. O. Box 1348 Venice, CA 90294 (310) 677–1057 or (310) 822–5757 Strongylocentrotus purpuratus (Dec–April) Lytechinus pictus (May–Sept) Dendraster excentricus (sand dollar) (May–July) S. purpuratus and L . pictus develop best at temperatures several degrees cooler than room temperature, but some embryos will develop normally at cool room temperatures. Dendraster embryos develop at room temperature. Marine Biological Laboratory Department of Marine Resources* Woods Hole, MA 02543 (508) 548–3705, ext. 7375 Arbacia punctulata ( June–Aug) S. droebachiensis (Nov–March) S. droebachiensis embryos must be maintained at temperatures of 12° C or lower for normal development and may be more difficult to use in teaching. *The MBL Marine Resources Dept. gives top priority to collecting for MBL investigators, but they are eager to try to meet the needs of others as well. Susan Decker 14140 S. W. 22nd Place Davie, FL 33325 (954) 424–2620 Leave message; she will return calls promptly. Ms. Decker supplies L. variegatus from late fall to late spring. She prefers to ship by air freight rather than UPS or Fed Ex. She ships only on Sundays so could supply labs early in the week. Ocean Resources, Inc. P. O. Box 19B Isle au Haut, ME 04645 (207 ) 335–2600 S. droebachiensis (spring). Echinarachnius parma (sand dollar) (spring and early summer). Note again that S. droebachiensis and E. parma do not develop normally at common room temperatures, though E. parma will develop normally in a cool room. Carolina Biological Supply 2700 York Road Burlington, NC 27215 (800) 334–5551 Carolina lists L. variegatus or E. tribuloides (their choice) and makes them available October through March only. Eucidaris is a cidaroid urchin and several features of its development vary from other species listed here. Connecticut Valley Biological 82 Valley Road P. O. Box 326 Southampton, MA 01073 (800) 628–7748 Connecticut Valley offers a “North Atlantic” sea urchin embryology kit with S. droebachiensis that they make available from December to mid-April and a “Mid-Atlantic” kit with A. punctulata or L . variegatus that is available April through November.

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Ward’s Natural Science Establishment 5100 West Henrietta Road P. O. Box 92912 Rochester, NY 14692–9012 (800) 962–2660 Ward’s offers a sea urchin embryology set that they state is available year round. The species of urchin supplied varies with the season.

C. Seawater There are several formulae for the preparation of synthetic seawater. I recommend the one used at the Marine Biological Laboratory at Woods Hole.Another widely used synthetic seawater formula was devised some years ago by Daniel Mazia at the University of California, Berkeley.These formulae are presented below. Reagent grade chemicals should be used, and the sodium bicarbonate ( NaHCO3) should be added after all the other salts are dissolved to avoid solubility problems.The synthetic seawater can be made up in glass-distilled water, or water prepared by other purification methods, but simply deionized water should not be used.

Synthetic Seawater (Woods Hole MBL) NaCl KCl CaCl2 • 2H2O MgCl2 • 6H2O MgSO4 • 7H2O NaHCO3 Distilled H2O

24.72 g 0.67 g 1.36 g 4.66 g 6.29 g 0.18 g To make 1 L volume

Source: Cavanaugh, G. M. Formulae and Methods V. Marine Biological Laboratory Chemical Room. Woods Hole, MA: Marine Biological Laboratory; 1964.

Synthetic Seawater (Mazia) NaCl KCl MgCl2 • 6H2O MgSO4 • 7H2O CaCl2 NaHCO3 Distilled H2O

28.32 g 0.77 g 5.41 g 7.13 g 1.18 g 0.2 g To make 1 L volume

There are a number of synthetic seawater salt mixtures on the market. Several that are recommended by many workers for use in culturing embryos and larvae of marine invertebrates are prepared by the Jamarin Laboratory, Osaka, Japan. For information, consult the Chemical Room, Marine Biological Laboratory,Woods Hole, MA 02543 (508–548–3705).They distribute the “Jamarin U” seawater salts mixture in units that will make 20 liters of synthetic seawater. Some colleagues use Instant Ocean for work with sea urchin gametes and embryos. In my experience, Instant Ocean has been very adequate for maintenance of adult urchins and sand dollars, but I have obtained very poor results using Instant Ocean as a medium for gametes and developing embryos.

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Calcium-Free Seawater* NaCl KCl MgSO4 • 7H2O NaHCO3 Distilled H2O

26.5 g 0.7 g 11.9 g 0.5 g To make 1 L volume

Calcium and Magnesium-Free Seawater NaCl KCl NaHCO3 Na2SO4 Distilled H2O


*Formulae for calcium-free seawater and calcium- and magnesium-free seawater are from D. R. McClay (1986) in Schroeder (1986, see p. 29).

D. Solutions: KCl, Hypertonic Seawater, PABA, LiCl A solution of 0.5 M KCl in distilled water is used to induce shedding of gametes.The formula weight of KCl is 74.56.Thus, 1 liter of 0.5 M KCl solution contains 37.28 g KCl, and 100 ml of 0.5 M KCl contains 3.73 g KCl. The solution of KCl should be injected in several places around the peristomial membrane. The needle must be inserted far enough to ensure that the solution enters the perivisceral coelom, not the lantern coelom. Some familiarity with the anatomy of the adult sea urchin is helpful. On occasion, the injection of KCl may not induce shedding. Sometimes shaking the urchin after injection will promote shedding. In the event of continued failure, dissection, removal, and opening of the gonads may yield usable gametes, but laboratory work can be unsatisfactory when gametes from animals unresponsive to KCl injection are used. Methods for stimulating spawning by electric shock are also described by Hinegardner in his article in Wilt and Wessells (1967 ). Hypertonic seawater can be prepared by adding 30 g NaCl per liter to seawater (E. B. Harvey 1956). A solution of 10 mM p-aminobenzoic acid ( PABA) in seawater is used for preventing fertilization membrane hardening. (Be certain to use the sodium salt of PABA, not the free acid.) The formula weight of PABA is 159.1; 10 mM PABA contains 1.59 g PABA per liter of seawater. A seawater solution of 60 mM LiCl (lithium chloride) is made up with 2.54 g LiCl per liter of seawater ( LiCl’s formula weight ⫽ 42.39). E. Solutions: Alkaline Phosphatase Staining A substrate and staining solution for localization of alkaline phosphatase activity can be made up using NBT/BCIP ready-to-use Tablets (catalog # 1 697 471), Roche Molecular Biochemicals, 9115 Hague Road, P. O. Box 50414, Indianapolis, IN 46250 ( Tel. 800–262–1640 or www.IbuyRMB.com). Each tablet contains substrates and buffer components, and when dissolved in 10 ml of distilled water, is adequate for one cover-glass staining jar. The pH 9.1 buffer solution and a phosphate buffered saline solution are made up using the formulae in the tables following.The pH 9.1 buffer utilizes Trizma Pre-set Crystals (Sigma catalog # T- 6128).

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pH 9.1 Buffer Trizma Pre-set Crystals MgCl2 • 6H2O NaCl Distilled H2O

6.15 g 1.017 g 5.844 g To make 1 L volume

Phosphate Buffered Saline NaCl KH2PO4 Na2HPO4 KCl Distilled H2O


Adjust pH to 7.4

Cover-glass staining jars (catalog # 72242-24 for 4/pk with screw caps or catalog # 72242-04 for 4/pk with glass lids) can be obtained from Electron Microscopy Sciences, 321 Morris Rd. # A, Box 251, Fort Washington, PA 19034 ( Tel. 800–523–5874 or 215–646–1566). Electron Microscopy Sciences also sells a coverglass staining rack (catalog # 72240) designed to be used for dipping cover glasses into fullsize Coplin staining jars. F. Microscopy Techniques Darkfield microscopy is often very useful for studying small, translucent objects such as sea urchin embryos (see “Using darkfield microscopy to enhance contrast,” C. K. Omoto and J.A. Folwell, Amer. Biol. Teacher 61(8):621–624, October, 1999).Views of sperm-egg interactions are also enhanced in darkfield. A darkfield arrangement is often available on teaching microscopes, or appropriate darkfield stops can be purchased.With only a little effort, darkfield stops can be made from readily available materials, and sometimes students gain new insight into microscopy by setting up their own darkfield arrangement. There is a filter slot or a snap-off filter holder ring at the bottom of most types of condensors that will accommodate a darkfield stop. A darkf ield stop has a circular opaque area set in the center of a translucent mount that is cut to fit the condenser filter slot or a filter holder ring.A heavy, clear overhead transparency sheet can be cut to the appropriate size and shape, and the dark, circular center can be made of black Scotch Plastic Tape or electrician’s tape. With a microscope that has a filter slot, the stop is more convenient to use if the clear carrier is cut with a small “handle” extending from one side.The size of the dark center of the stop that is required depends on the power of the objective being used and, to some extent, on the point where the stop is inserted into the light path. It will take only a little experimentation by you or your students to determine the proper size for the opaque center of a darkfield stop that works well with a low-power (10 ⫻ to 12.5 ⫻ range) objective. I suggest that if your microscopes have filter slots or filter holder rings that accommodate 3.5 cm filters, a centered opaque spot about 1.5 cm in diameter would be a good starting point for your experimentation. I find that students generally like to tinker with darkfield stops, and I think it’s worth some lab time because it involves them in microscopy in a way that’s new for most of them. For the darkfield technique to work, a microscope’s condenser must be racked up to its top position with its iris diaphragm wide open. At first it might seem counterintuitive, but achieving the darkfield effect at higher magnifications such as using high-power (around 40 ⫻) objectives requires use of larger darkfield stops. Most observations of sea urchin or sand dollar development can be made, however, with low-power (10 ⫻ to 12.5 ⫻) objectives.There also are some interesting effects that can be obtained with off-center or oval 206

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darkfield stops. Finally, darkf ield stops with a small “pie piece” cut out (“Pac-Man filters”) sometimes produce some striking interference effects. Some students take considerable pride in achieving spectacular effects with their own (sometimes highly modified!) filters. Rheinberg illumination is a variation on the darkf ield technique in which the center of the light column is only partly occluded. Rheinberg illumination gives very striking color contrast effects even with inherently low-contrast objects such as sea urchin and sand dollar gametes and embryos (see two papers by Morton Abramovitz: Scientific American, April 1968, pp. 125–130, and American Laboratory, April 1983, pp. 38–41 for more background information). It is well worth the effort to introduce students to this historical, beautiful, and effective form of illumination. A Rheinberg filter is constructed of three parts: a translucent plastic mount of the same diameter as the darkfield mounts that fit your microscopes; a circular piece of gummed white paper (a piece of a mailing label works well); and a circular piece of translucent, but darker-colored plastic. The colored plastic presentation folders that are sometimes used for papers and reports make excellent sources of material for constructing Rheinberg filters. Cut a small hole (a little less than 1 cm in diameter) in the center of the mount. Cut circular pieces of the mailing label and the darker plastic that are about 1 cm in diameter.Then apply them to opposite sides of the mount so that the sticky side of the label paper contacts the darker plastic through the hole in the mount (fig. A.1). Then press them together firmly. Once again, you or your students may need to experiment a bit to determine an optimal size for the central elements of the filter for use with your microscopes. Rheinberg microscopy also requires that a microscope’s condenser be racked up to its top position with its iris diaphragm wide open. 7 mm 12 mm

(a)

Darker plastic

Lighter plastic

Gummed label paper

(b)

FIGURE A.1 Assembling a Rheinberg filter. a) The largest part is a piece of light-colored plastic cut to fit the microscope’s filter holder. Cut a hole about 7 mm in diameter in its center. The other parts are a piece of darker-colored plastic and a piece of gummed label paper. Each is about 11 or 12 mm in diameter. b) The smaller parts are applied from opposite sides of the larger plastic and pressed into place so that the gummed paper holds all three together.

The darker plastic at the center contributes the background color, and the mount contributes a bright, contrasting color to surfaces. A red center on a light-yellow mount is very effective for viewing flagella and cilia.A blue center on a clear mount produces soothing and aesthetically pleasing views of embryos and larvae. It’s fun to experiment with color combinations and component sizes for Rheinberg filters.The results can be very striking. Polarizing microscopy produces beautiful and informative images of the developing skeletal spicules of sea urchin larvae, which contain crystalline arrays that rotate plane polarized light. Some microscopes are equipped for polarizing microscopy, but if your microscopes are not, it is fairly easy to set up a simple arrangement that works effectively. A circular piece of a polarizing plastic sheet cut to the proper diameter can be inserted in the filter holder of the condenser (the polarizer), and another piece cut to the proper diameter can be set on the outer surface of an eyepiece lens where it can be Appendix A

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rotated as the analyzer component.This works quite well, so it is not necessary to attempt to place the filter inside the eyepiece. In fact, some attempts to do so can turn into disasters! Be certain to remove any thin protective sheets that may be present on the surfaces before using the polarizing plastic. Polarizing plastic can be obtained from the manufacturer International Polarizer, Inc., 320 Elm Street, Marlborough, MA 01752 ( Tel. 508–481–7495).They normally require a fairly substantial minimum order, but in the past, they have been willing to sell smaller quantities (2–17⬙ ⫻ 39⬙ sheets minimum order) to educational institutions via purchase orders. Smaller quantities of polarizing plastic can be obtained from Edmund Scientific and similar suppliers. “Feet of clay” is a technique often used to elevate cover glasses slightly so that they better accommodate eggs and embryos without compression.The technique involves gently dragging each corner of a cover glass across a piece of modeling clay so that a tiny piece of clay is stuck to each corner before the cover glass is placed on a drop on a slide.This technique nicely replaces the use of strips of modeling clay suggested for observing fertilization in Laboratory 1, for example. G. Filters and Filter Holders Filters to be used for fertilization membrane removal, blastomere separation, and culture concentration (with turkey baster) can be obtained from a variety of sources that supply plankton nets or the nylon netting for constructing plankton nets. Carolina Biological Supply offers 30-cm square nylon laboratory sifters in several useful mesh sizes (their basic catalog number is 65–2222, with an added letter to designate mesh size).Wildlife Supply Company,301 Cass Street,Saginaw,MI 48602 ( Tel.517–799–8100; website: www.wildco.com) sells Nitex plankton netting in one-third yard quantities. Aquaculture Supply, 33418 Old Saint Joe Road, Dade City, FL 33525 ( Tel. 352-567–8540; website: www.aquaculturesupply.com) offers polyester filter cloth that is somewhat less expensive than Nitex netting.Another primary source of Nitex netting is Tetco, Inc., 95 Valley Street, Bristol, CT 06010 ( Tel. 860–583–1209), but they may require large minimum orders. Mesh sizes of around 70 ␮m usually are effective for removing the fertilization membrane and separating blastomeres of Lytechinus and Dendraster, but somewhat smaller mesh sizes (48 to 53 ␮m) are usually required for S. purpuratus and Arbacia, while the considerably larger eggs of S. droebachiensis would be damaged by filters with these mesh sizes. I would suggest trying a mesh size of 120 ␮m for S. droebachiensis. However, the best way to avoid disappointing results is to have available various filters of several mesh sizes and to do preliminary demembraning trials before beginning a complex experiment. A filter holder can be made by cutting off the bottom of a 50-ml plastic syringe or a 50-ml plastic beaker. A 10-cm (4⬙) piece of PVC pipe with an inside diameter of 2.5 cm (1⬙) also works well. A filter can simply be held firmly in place while the eggs are poured through, or the filter can be fastened in place with a rubber band. A filter should be thoroughly washed and rinsed after each use, since students almost invariably manage to turn a filter over between batches of eggs. Also, a filter can quickly become occluded if debris is allowed to dry in it. A pipe with a filter-covered end is suggested for concentrating cultures (with a turkey baster) in Laboratories 2 and 3 and for dipping embryos or larvae in an alternative method of deciliation in Laboratory 2.To make this device, cut a piece of 35–40 ␮m mesh filter. ( I routinely use the 37 ␮m mesh that Carolina Biological supplies.) Then glue the piece of filter to the end of the pipe.After the glue has dried, trim away the excess filter with scissors or a razor blade. It is a good idea to have several spares of these handy devices available in the lab. Students and professors come up with all sorts of applications for them. The surfaces of the filters should be washed using a distilled water squirt bottle after each use. H. Feeding Pluteus Larvae It is difficult to rear pluteus larvae to advanced stages in the teaching laboratory, particularly in inland institutions, because of the need for relatively large volumes of artificial seawater, the effort that must be expended to grow the marine algae that the larvae require for food, and the need for constant

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slow stirring to keep the cultures aerated. Nevertheless, students often want to try to feed larvae.There actually are some sources of concentrated algal suspensions that can be used for feeding, but there is a less satisfactory but much simpler substitute method that will keep larvae growing for at least a short time. Student hopes for results using this method should not be inflated, but they can achieve some modest short-term success. ( I once raised Lytechinus variegatus plutei to the eight-armed stage with apparently normal urchin rudiment development using the method.) Transfer two-armed pluteus larvae to clean Woods Hole Formula artificial seawater and maintain them in very sparse cultures. Larvae can be supplied food beginning at the two-armed stage.About 100 larvae can be reared in 500 to 700 ml of seawater in each large beaker. I have not yet tested the effects of greater density, but I think that slightly greater densities might be permissible. Aeration should be done twice daily by repeated gentle pipetting of air to the bottom of the cultures. Add Roti-Rich Liquid Invertebrate Food (Aquaculture Supply, address in previous section, or Ward’s Natural Science Est.) three or four times per week. Mix 1 drop of Roti-Rich with 5 drops of seawater, and gently stir 2 drops of this mixture into each culture so that each culture received one-third drop of Roti-Rich at each feeding. Some flagellate organisms may be observed in the culture, and undoubtedly there will be bacterial growth in the cultures as well so the plutei may consume those organisms in addition to the supplied Roti-Rich. Transfer all larvae by pipette to clean seawater twice a week. It might be possible to transfer larvae by filtering and gentle washing, but I have concerns about damaging the larvae and transferring the other organisms in the culture vessels.

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Appendices

B: Preparation of Materials for Experimentation on Frog Embryos


A P P E N D I X

B Preparation of Materials for Experimentation on Frog Embryos

A. Procurement and Storage of Frogs Adult male and female leopard frogs (Rana pipiens) can be obtained from several commercial dealers, one of which is NASCO, 901 Janesville Avenue, Fort Atkinson,WI 53538 ( Tel. 1–800–558–9595; website: www.nascofa.com). Frogs in pre-breeding condition can be shipped from mid-October through late March. Female frogs obtained from dealers in late September or early October can be induced to ovulate, but the percentage of fertilization is not high. Therefore, experimentation on Rana pipiens embryos is best performed during the period between late October and March. Upon delivery, female frogs should be stored in a refrigerator. Male frogs may also be stored at this low temperature, although they can be maintained in frequently replaced cool water ( below 20° C). Place groups of five or six females in large glass bowls covered securely with wire mesh or in large, covered plastic containers with drilled air holes. Add a small volume of spring water or dechlorinated tap water.Tap water should be aerated vigorously or boiled to drive off excess chlorine and chilled to 4° C before the frogs are put in it. Change the water three times per week. It is not necessary to feed frogs during confinement at this low temperature.When females are to be induced to ovulate, they are brought to room temperature and allowed to equilibrate for half an hour before receiving the pituitary injections. Methods for inducing ovulation are described in Part E. B. Glassware, Culture Dishes, and Pipettes Students of amphibian biology traditionally use glass finger bowls for maintaining the frog embryos and larvae. Finger bowls are called “Culture Dishes” by some suppliers. Plastic dishes can be substituted for glass ones. Some workers keep embryos in Syracuse dishes and then transfer the larvae to paraffincovered paper cups (for example, Dixie cups). Clearly, a variety of rearing containers can be used.Tadpole larvae can be fed on canned spinach or boiled, cooled lettuce leaves, but it is important to remove old food material before it fouls the water. The operating dish frequently employed is a Stender dish, 51 mm in diameter and 26 mm in height, but Stender dishes of other sizes, as well as a variety of other small glass containers (or small petri dishes), can be used.As described in a subsequent section, the bottom of the operating dish is covered with agar, since denuded embryos tend to stick to glass surfaces.

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Wide-mouth pipettes are used to transfer embryos.These transfer (“frog”) pipettes are made from glass tubing (outside diameter: 9 mm; inside diameter: 6 mm). A strip of tubing is heated in the middle and drawn out. After cooling, the tapered end is cut at a level that creates an opening having an inside diameter of 1.8 to 2.0 mm.Alternatively, ordinary medicine droppers may be cut off at appropriate levels and fire-polished to produce “frog pipettes.” Other indispensible items are petri dishes, Syracuse watch glasses, and depression slides. Instead of glass depression slides, plastic slides with a central well can be used. Quartz microscope slides can be purchased from McCrone Associates, 850 Pasquinelli Drive,Westmont, IL 60559 ( Tel. 630–887–7100, 1–800–622–8122; website: www.mccrone.com/mac/shopping.html). Sterilization of all glassware in an autoclave is desirable. If an autoclave is not available, the operating dishes may be rinsed with 70 percent ethanol and then placed in a covered container for at least 12 hours to permit all alcohol to evaporate. C. Instruments 1. Glass Needles Very fine needles, made from glass or tungsten, are used for the extirpation and transplantation experiments on the amphibian embryos. During preparation of glass needles, a microburner is employed. The microburner is used to give a small, intense flame and is essentially a modified hypodermic needle. A piece of 1/4-inch tubing is inserted through a hole in a No. 15 cork. On the top side, the rubber tubing should protrude about 1/2 inch.A No. 18 hypodermic needle is inserted into the tubing. Beneath the cork, several inches down the tubing, attach a screw-type pinchcock.This allows very precise control of the size of the flame.Attach the cork to a ring stand with a burette clamp, and connect the free end of the tubing to a gas outlet. Glass needles are prepared from solid glass rods, 5 mm in diameter. A strip of rod (about 20 cm long) is heated in the center over a burner. The ends of the rod are pulled apart, or drawn out, when the central portion becomes soft. The connecting thread is then broken apart, creating two rods, each having a handle approximately 10 cm long and a tapered tip. The tapered tip is then heated over the microburner until the thin piece bends slightly.Then touch the tip of the tapered piece to the back of the hot needle in the microburner. Wait until the glass melts and adheres to the metal needle. At the moment the glass adheres to the needle, pull the glass out in a rapid motion.The tip of the glass should be drawn to a very fine point.The tip should be firm and sharp, not elastic. 2. Tungsten Needles and Cactus-Spine Needles Many workers prefer tungsten needles, since they are more durable. Tungsten wire may be purchased from Alfa AESAR Div., 30 Bond Street, Ward Hill, MA 01835–8042 ( Tel. 1–800–343–0660, 978–521–6300; website: www.alfa.com). Pieces of tungsten wire (for example, catalog # 10408, 0.25-mm dia. tungsten wire from AESAR) approximately 2 to 3 cm long are mounted or sealed on a length of glass rod or tube. Pieces of tungsten wire also may be mounted in ordinary bacteriological loop holders to save time when needles are being “mass-produced.” If an angled needle is desired, the bent portion of the unsharpened wire should be approximately twice as long as the desired final length of the tapered tip. It is essential to wear protective eyewear throughout the procedure described here. The tip of the wire is first flamed with an oxygen-gas burner or ordinary propane torch until the wire tip glows “white.”Avoid looking directly at glowing metal except for an occasional glance to check the flaming process. A reasonably useful needle can be produced directly by this heating method. As heating is continued, periodically check the effects of vaporization of tungsten from the tip by examining the needle under a dissecting microscope.The degree of taper and the length of the tapered portion can be controlled by varying the length of the portion being heated. Because of the problems involved in further treatment of needles, you may want to compromise on needle quality and use needles prepared by heating alone.

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However, a smoother surface can be obtained when the tip of the wire is sharpened to a fine point by dipping it in and out of molten sodium nitrite (or sodium sulfite).The hot bath is prepared by melting anhydrous sodium nitrite in a porcelain crucible on a triangle. The surface of the tungsten needle should be smooth, not uneven and/or pitted, after this treatment. The evenness achieved in the metal when removed by the molten chemical will depend upon the temperature of the bath. The molten sodium nitrite is very corrosive; protective eyewear and heavy, protective gloves are mandatory. Extreme caution must be observed in working with the hot bath of sodium nitrite, and I urgently recommend that students not be allowed to use this dangerous and tricky method. Cactus-spine needles (p. 223) can provide an easy-to-make alternative to glass or tungsten microsurgical needles. 3. Ball-Tipped Glass Rods Another glass instrument is the “ball-tipped” glass rod. Ball tips are used to make depressions in the agar operating surface in which the embryos are held.A strip (20 cm long) of solid glass rod (5 mm in diameter) is heated in the middle, drawn out to a slender rod, and broken in the middle. The thin end of each of the two rods is then held in a vertical position over a Bunsen burner or a microburner.The glass tip will contract into a round ball upon heating. Various sizes of ball tips should be made. 4. Glass Bridges Transplants are held in place during healing by glass bridges.These are rectangular pieces of glass, 3 to 4 mm wide and 10 to 12 mm long, cut from coverslips (thickness No. 2) with a diamond pencil. The rough edges of the glass should be smoothed by passing the four edges slowly through the flame of a microburner.The bridges may be bent at various angles by grasping one end with forceps and heating, from below, an area a short distance from the other end.The glass will bend under its own weight. A glass bridge should stand firmly on its two ends. 5. Hair Loops Hair loops are extremely useful for manipulating embryos and for transferring pieces of excised tissue.A hair-loop holder is prepared using glass tubing (outside diameter: 5 mm; inside diameter: 3 mm).A 20-cm piece of tubing is heated in the middle and drawn out straight.The large end is flamed shut.With watchmaker’s forceps, insert the two ends of a strand of baby’s hair into the narrow end.The loop of hair can be adjusted to the desired size.The insertion of the strand of hair is done under a dissecting scope. The hair is sealed on the glass tube by dipping the hair loop into melted paraffin.The liquid paraffin will be drawn into the opening of the tube by capillary force and will subsequently harden. Paraffin covering the loop itself can be removed by touching the loop to a warm glass slide. Some workers substitute very fine threads obtained by shredding POH-brand dental floss for baby hairs in making hair loops. 6. Watchmaker’s Forceps Ordinary blunt laboratory forceps, which are usually made of nickel-plated forged steel, will not suffice for work on amphibian or chick embryos.Watchmaker’s forceps (also called microdissection forceps in some catalogs) are indispensible tools. These forceps are made of stainless steel and have extremely fine tips. They can be purchased from some general laboratory supply houses, specialized instrument suppliers, or a wholesale jeweler’s supply company. With use, the points of watchmaker’s forceps tend to become bent or otherwise damaged. Sharpening and minor repair can be done with an Arkansas stone that has been soaked in thread-cutting oil. More extensive repairs can be accomplished with a rougher sharpening stone (always use a drop of cutting oil), but fine surface smoothing should be completed with a smooth Arkansas stone. I make students responsible for the condition of their own microdissection forceps from the first day of use. This produces a much greater level of care than does simple replacement of damaged forceps that shifts responsibility to someone other than the users.

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D. Media and Chemicals 1. Ringer’s Solution Fertilization of the leopard frog egg and subsequent development of the embryos can be undertaken in spring water or natural pond water. If local pond water is not available, a satisfactory substitute is diluted (10%) Amphibian Ringer’s solution. Ten parts of stock, or full-strength, Ringer’s solution is added to 90 parts of distilled water.The composition of stock Ringer’s solution (as with all solutions prepared in distilled water using reagent grade chemicals) is as follows: Stock (100%) Amphibian Ringer’s Solution NaCl KCl CaCl2 (2H2O) MgCl2 • 6H2O NaHCO3 (added last) Water (glass-distilled)

6.60 g 0.15 g 0.15 g (0.20 g) 0.20 g 0.10 g (approx.*) 1000 ml

*Adjust final pH to 7.8 with NaHCO3

The anesthetic ethyl m-aminobenzoate methanesulfonate (also called Tricaine or MS-222), may be obtained from various laboratory supply companies or chemical suppliers. The substance is suspected of being mildly carcinogenic, so it should be used with care. 2. Orcein Solution In Laboratory 4, a procedure is described for staining tadpoles’ chromosomes.The stain employed is a 2% solution of aceto-orcein, and a commercial preparation of the orcein stain obtained from Carolina Biological Supply Company (and probably other suppliers as well) works satisfactorily. Alternatively, a staining solution can be prepared from powdered natural orcein, but I strongly urge that students not be allowed to carry out this dangerous procedure. Wear protective eyewear and other protective lab gear. Add 2 g of powdered orcein to a mixture (be careful! ) of 60 ml of glacial acetic acid and 40 ml of distilled water. The mixture must be refluxed slowly for 1 hour ( be sure to use boiling stones). After cooling, the solution is stored in a well-stoppered, dark bottle. Since this highly concentrated stain will tend to crystallize out, it is desirable to filter it just before use. However, the filtering process may be avoided by the simple expedient of drawing the stain for use from the middle of the bottle with a pipette. 3. Barth and Barth’s Operating Medium The operations on the extirpation and transplantation of embryonic tissues in Laboratory 6 may be performed in 100% Amphibian Ringer’s solution, but a preferred operating medium is Barth and Barth’s medium that consists of three components or stock solutions, shown in the following table: Barth and Barth’s Operating Medium Solution A NaCl KCl MgSO4 • 7H2O Ca(NO3)2 • H2O CaCl2 • 2H2O H2O to 500 ml

214

Appendix B

5.150 0.075 0.204 0.062 0.060

Solution B g g g g g

NaHCO3 H2O to 250 ml

0.200 g

Solution C Na2HPO4

0.0300 g

KH2PO4

0.0375 g

H2O to 250 ml


Appendices



The unmixed stock solutions are stored in a refrigerator. When ready to be used, the three components are mixed in the following proportions: 50 ml of A, 25 ml of B, and 25 ml of C. Prior to mixing the three together, heat 25 ml of solution C just to boiling to sterilize it, allow it to cool, and add 100 mg of serum globulin. Solutions A and B are also brought to a boil over a flame and cooled before all three are combined.This sterilization by “light” boiling will suffice. It is advisable to add the NaHCO3 solution ( B) last to avoid precipitation of the calcium and magnesium salts in A. To further reduce the risk of bacterial and fungal infections, add 100 mg each of sulfadiazine ( USP Powder, Lederle) and mycostatin (100,000 units/gm Squibb) per liter of medium. 4. Agar-Bottom Operating Dishes The agar base is prepared as follows: 20 g of charcoal (animal bone black) is added to 900 ml of distilled water in a 1000-ml beaker. Be very careful in handling bone black because it can be explosive. This dangerous procedure should be done while wearing protective eyewear and with other appropriate precautions. I recommend that students not be allowed to carry this out. Keep the dry powder and especially any airborne dust away from flames or sparks. Bring the mixture to a boil and slowly add 20 g of plain, nonnutrient agar. Continue to boil this mixture for a few minutes. Pour this solution directly into the Stender dishes, quickly cover the dishes, and allow the solution to cool. Approximately 15 agar-bottom dishes can be prepared from this one solution.The dishes, when cooled, are stored in a refrigerator until they are ready to be used. 5. Steinberg’s Medium, Disaggregating Solution, Reaggregating Solution Explants of embryonic tissue can also be cultured in Steinberg’s medium.

Steinberg’s Culture Medium Stock Solutions A. B. C. D.

17 g NaCl per 100 ml distilled water 0.5 g KCl per 100 ml distilled water 0.8 g Ca(NO3)2 • H2O per 100 ml distilled water 2.05 g MgSO4 • 7H2O per 100 ml distilled water

In preparing the medium, 10 ml of solution A, plus 5 ml each of solutions B, C, and D, are made up to 500 ml with distilled water.Add Tris buffer (280 mg) and adjust the pH with IN HCl to 7.8 to 8.0. Tris ( hydroxymethyl) aminomethane may be obtained from the Sigma Chemical Company in St. Louis, MO. Steinberg’s complete medium may be modified into a disaggregating medium by eliminating the calcium and magnesium ions ( hence,the solution is made up without solutions C and D) and the addition of 60 mg of EDTA. Lastly, Steinberg’s solution serves well for culturing reaggregating cells if it is supplemented with 0.1% bovine plasma albumen, 0.1% human serum globulin (see Jones and Elsdale 1963, cited on p. 95) or even (as in Laboratory 7 ) albumen from a hen’s egg. Use the “runny” portion of the egg white to make up this solution. Crack the shell into two parts and then grasp the chalaza and pull it and the yolk over the rim of the shell and discard them. Usually the “stringy” portion of the egg white will also be carried along and the “runny” portion will be left behind.The Laboratory suggests a 1% solution of egg white, but you may wish to make up, and compare results obtained with, additional reaggregating solutions that have greater protein concentrations. Mix the egg white in by swirling or shaking the solution in a stoppered flask.

Appendix B

215


Appendices



E. Induction of Ovulation in the Leopard Frog Several suppliers, including NASCO, have prepared “ovulation sets” for use in teaching laboratories. One type of kit consists of a vial of pituitary powder (or several preserved pituitaries) and a set of adult frogs, while a simpler type of kit includes pre-injected female frogs as well as male frogs.“Pituitary kits” are generally available from early November to March. It is also possible to work entirely with living frogs and dissect pituitary glands for use in inducing ovulation, but I recommend against doing so, in light of the problem of declining amphibian populations (see p. 238). An instruction pamphlet is provided with each kit, but general instructions for frog injection written by E. Peter Volpe are provided here, and I recommend that you also read them before proceeding with frog work. When received from a dealer, females should be stored immediately at 4 to 8° C, inasmuch as ovarian eggs undergo degenerative changes at room temperature. Bring the female to room temperature only when she is to be used for the induction of ovulation. Males can be maintained at room temperature at all times. Draw the material to be injected into the barrel of a 2-ml glass syringe and attach a 1-inch, 18-gauge hypodermic needle. Gently tap the barrel to center particulate material over the opening of the syringe and maintain the syringe in a vertical position. Hold the recipient female in your left hand and insert the needle downward (as the glands are heavier than water) through the skin and abdominal muscles in the lower quadrant of the abdomen. Exercise care to avoid injury to the viscera. Quickly inject the glands downward, but withdraw the needle cautiously.As you slowly remove the needle, pinch the skin at the point of needle entry to prevent any loss of fluid or particulate material. Finally, draw more Ringer’s solution into the syringe to ensure that no pituitary material has remained lodged in the barrel of the syringe or in the needle. Reinject the recipient female with all pituitary material that may have been retained. Place the recipient female in a wire-covered bowl with a small amount of water and keep her at a cool temperature. If maintained at a constant 18° C, fertilizable eggs can be obtained within 36 to 48 hours.At 20° C, ovulation can occur in 30 to 36 hours.At room temperature (about 23° C), eggs can be obtained within 24 hours. Experience shows that best results are obtained when the recipient female is maintained at 18 to 20° C for 36 to 48 hours. Recipient females should be initially tested for the presence of eggs by stripping 24 hours after injection.The stripping technique is described in Laboratory 4. If a string of eggs emerges from the cloaca, the sperm suspension can then be prepared. If only jelly or fluid oozes out of the cloaca, retest the female at 12-hour intervals up to 48 hours. If eggs cannot be obtained after 48 hours, a second injection of pituitary is required, comprised of one half the original pituitary dosage. Such reinjected females should ovulate within 24 hours after the second injection.

216

Appendix B


Appendices

C: Preparation of Materials for Experimentation on Xenopus laevis


A P P E N D I X

C Preparation of Materials for Experimentation on Xenopus laevis

The African clawed frog, Xenopus laevis, is fully aquatic, both before and after metamorphosis. Xenopus can be maintained readily in the laboratory, and fertile eggs can be obtained at any time of year by injecting chorionic gonadotropin. Much of the general information about the preparation for work with frog embryos given in Appendix B is applicable to work with Xenopus, but there are some techniques and procedures that are specific to Xenopus. Xenopus adults can be obtained from several suppliers. Some of them are NASCO 901 Janesville Avenue Fort Atkinson, WI 53538 Telephone: 800–558–9595 Website: www.nascofa.com Xenopus I 5654 Merkle Road Dexter, MI 48130 Telephone: 734–426–2083 Website: www.xenopusone.com Carolina Biological Supply Co. 2700 York Road Burlington, NC 27215 Telephone: 800–334–5551 Website: www.carolina.com

Connecticut Valley Biological Supply Co. P.O. Box 326 82 Valley Road Southampton, MA 01073 Telephone: 800–628–7748 Website: www.ctvalleybio.com Ward’s Natural Science Establishment 5100 West Henrietta Road P.O. Box 92912 Rochester, NY 14692 Telephone: 800–962–2660 Website: www.wardsci.com

Xenopus suppliers provide information about maintaining Xenopus, and I recommend that their booklets be consulted. A very useful booklet by Etheridge and Richter, “Xenopus laevis: Rearing and Breeding the African Clawed Frog,” is published and distributed by NASCO. This booklet provides all of the basic information needed to work with Xenopus, and it can be purchased for a modest price. Carolina Biological Supply offers a comparable booklet on Xenopus (also modestly priced) that includes very nice color photographs of developmental stages. I recommend obtaining a copy of this booklet as well.

217


Appendices



There are a few additional comments regarding Xenopus that may prove helpful. Even if four liters (about a gallon) of dechlorinated tap water are provided for each adult Xenopus held in a large (for example, 20-gallon) aquarium, the water should be changed at least every other day.The animals can make it through a weekend but need immediate attention on Monday morning. Tap water usually can be dechlorinated adequately by allowing it to stand in three-quarters-filled, uncovered gallon containers for several days. Cautions about covering tanks in which adult frogs are kept should be observed.These frogs can jump out. In fact, they (especially males) will even attempt escape from the breeding tank. Keep covers on and firmly weighted.The frogs are quite slimy and are adept at slipping through very small cracks. Of course, they don’t fare well out in the air and die by desiccation within a very short time.Adult frogs seem to do well in glass-bottom aquaria, but some rougher aquarium bottoms seem to irritate them (they literally wear off their claws). It is convenient to feed Xenopus the pellet food sold by supply companies according to the directions given. The frogs can also be fed frozen beef liver. Shavings of liver can be sliced off a frozen chunk and cut up into small pieces that thaw quickly enough to be fed almost immediately. Whatever the food used, it is important that the frogs’ water be changed after they have finished feeding to avoid fouling the water by bacterial growth. NASCO suggests a 2-week “conditioning” period before breeding. During this period, there should be daily feedings and water changes. If the frogs are newly arrived, they definitely need at least a week in aquaria before breeding to recover from the stress of shipping, whether or not it is possible to give them a full “conditioning” period. Hormone injection schedules and information on injection techniques are provided in bulletins from suppliers. NASCO’s schedule (nearly the same as Injection Schedule No. 2 suggested by Carolina Biological Supply) produces good results. Please note that chorionic gonadotropin is a human product and should be treated as a biohazard. Wear gloves and handle solutions, syringes, and injected frogs with care. Chorionic gonadotropin comes in vials containing either 2500 I.U. or 5000 I.U.The contents of the vials should be dissolved in either 2.5 ml or 5 ml of amphibian saline, respectively. This gives a concentration of 1000 I.U. per ml of injectable solution. Keep the female and male alone in covered fishbowls or other dishes throughout the injection schedule.The injection schedule suggested by NASCO is 1st Day: 8:00 A.M., inject male with 150 I.U. (0.15 ml of above solution) 2nd Day: 8:00 A.M., inject female with 250 I.U. (0.25 ml of above solution); inject male with 150 I.U. 4:00 P.M., inject female with 500 I.U. (0.5 ml of above solution); place the pair together in the breeding tank. Breeding tanks can be purchased or made, using a plastic dishpan containing half-inch mesh screen held off the bottom with rubber stoppers.The screen should fit snugly, so that the frogs cannot get under it or suffer injury by becoming jammed between the wall of the pan and the screen.The pan should contain enough diluted Amphibian Ringer’s solution or spring water so that the surface is about 5 cm above the screen. Put an escapeproof cover on the pan after adding the two frogs and place the pan in a quiet, dimly lit room where it will be undisturbed overnight. Eggs are released a few at a time during a long period of amplexus, so there should be embryos at a number of stages of development in the pan the next morning.The frogs may stay in amplexus even after shedding ends. When the male and female are removed from the tank, they should be returned to isolation dishes for several days before they are placed with other frogs because they will be excreting chorionic gonadotropin. Change the water if you wish to leave the eggs in the tank, but it is probably better to transfer groups of eggs to finger bowls. A section lifter or some other flat device is very useful for scraping the eggs loose from the pan bottom and the screen because the jelly around them is very sticky.

218

Appendix C


Appendices



If you are used to the timetable of Rana pipiens development, you will be amazed at the speed of development in Xenopus. Some eggs should be put in a cool place so that several stages of development will be available for class use. In fact, if all of the eggs are left at room temperature and your class doesn’t meet until the afternoon, you will find that all of the embryos may be in the yolk plug stage, or beyond, by the time your students see them. It is also possible to adjust the injection schedule and pairing of the frogs to make earlier stages available, but this may require some midnight or early morning work. Diluted (10%) Amphibian Ringer’s solution (p. 214) is an appropriate medium for developing embryos and tadpoles. Spring water from grocery stores works well also, but carefully check what is labeled “spring water” or “drinking water.” Sometimes it is distilled or deionized water. There are good directions for handling and observing embryos and tadpoles in several of the available booklets, but there is one note of caution about feeding tadpoles that must be emphasized. Commercial tadpole powder or powdered nettle can be fed once the hatchlings are swimming actively, but it is very important to make certain that the powder is completely wetted to a paste before putting it into the water.The tadpoles have serious trouble with chunks of dry powder. Tadpoles have translucent bodies that make observations of internal structural details quite easy. The anesthetic ethyl m-aminobenzoate methanesulfonate (MS-222) works well on Xenopus tadpoles when they are immersed in a 1⬊3000 solution of the anesthetic in diluted Ringer’s or spring water.This permits observations of structural details and the eventual dissection of the tadpoles. Swimming tadpoles can also be maintained in aerated tap water, but it might be well to test the water with a few tadpoles before committing an entire batch to tap water. In some cities, aerated tap water seems to be toxic to tadpoles at some times and not at others, a fact that could make one give up drinking tap water! Several different injection schedules are used to prepare Xenopus for use in vitro fertilization. Throughout the injection schedule, keep female and male frogs alone in covered containers in locations where they won’t be disturbed.Three of the injection schedules follow. SCHEDULE 1

Three days before lab, inject dorsal lymph sacs of sexually mature Xenopus females with 30 I.U. (50 I.U. for very large females) Pregnant Mare Serum Gonadotropin (Sigma #G4877). Seven to nine hours* before lab, inject females with 700 I.U. (800 I.U. for very large females) Human Chorionic Gonadotropin (Sigma #CG5 or CG10). SCHEDULE 2

Some workers use and recommend this simpler injection schedule. Seven to nine hours* before projected use, inject female Xenopus with 700 I.U. of chorionic gonadotropin. Male injection on the preceding day is optional in this protocol. SCHEDULE 3

Day preceding lab: Morning: inject one or two male Xenopus with 150 I.U. of chorionic gonadotropin Afternoon: inject several female Xenopus with 200 I.U. Midnight:* inject the females with 600 I.U.

*In order to ensure more precise timing of egg availability, some Xenopus researchers recommend that Xenopus females be brought to 18° C before (final) HCG injection and held at 18° C after injection.This temperature-control protocol should make eggs available 16 hours after injection.The females should be warmed to room temperature an hour before egg stripping.

Appendix C

219


Appendices



This schedule is intended to provide eggs for use in morning labs. For afternoon labs, the two injections of female frogs should be 6 to 7 hours later (midnight and 6 A.M.). To obtain testes, stun a male frog with a blow to the head by firmly gripping the male frog’s legs and swinging it, striking the top of its head against a hard surface. Swiftly decapitate the frog, spinal pith it, and wrap it in wet paper toweling until it is needed for testis dissection. This procedure should be done quickly and carefully and should not be done in a student laboratory. Many people prefer to sacrifice Xenopus males in a chloroform bucket. Some people place Xenopus in a bucket of ice water until they are chill-anesthetized before use. Reference Nieuwkoop, P. D.; Faber, J. Normal table of Xenopus laevis. Hamden, CT: Garland Publishing, 1994. (Reprint of 1956 edition with new forward by J. Gerhart and M. Kirschner.)

220

Appendix C


Appendices

D: Preparation of Materials for Experimentation on Chick Embryos


A P P E N D I X

D Preparation of Materials for Experimentation on Chick Embryos

A. Procurement, Storage, and Incubation of Eggs Fertile eggs are sometimes difficult to obtain, and the supply and quality of eggs is often highly seasonal. A first choice for a source probably would be eggs from agricultural colleges and poultry research stations. However, good, usable eggs may be available from local hatcheries or farmers who keep roosters in their flocks year round. In urban areas, contacts with suppliers of fertile eggs usually can be made through university or hospital virus diagnostic or research laboratories. Several biological supply companies and larger hatcheries (see p. 225) ship fertile eggs, but shipping may be hazardous for the eggs, and supply company prices may be rather high, considering the quantity of eggs required for teaching developmental biology. Eggs may be stored at about 8 to 10° C for up to a week without serious consequences, but longer periods of storage lead to reduced viability. Temperatures below 7° C, and the desiccating conditions that normally exist in laboratory refrigerators, should be avoided. Reset a refrigerator to an appropriately higher operating temperature and place a pan of water in it before using it for egg storage. Before incubation, eggs should be removed from the refrigerator and allowed to come to room temperature. Normally, they should be set out in the laboratory at least 5 or 6 hours before they are incubated. It will do no harm to allow eggs to stand at ordinary room temperature overnight, but the length of such pretreatment might influence the amount of incubation time required to bring embryos to a given stage of development, particularly if the place where they are set is quite warm. Several kinds of incubators are available, but I prefer circulated-air incubators to still-air models. However, almost any type of incubator will be adequate if you take the time to test its performance under your laboratory conditions with the eggs that you have available. Incubator space can be the limiting factor in lab work with chick embryos, but efficient use of space or schedule juggling can make it possible to work with a less than ideal amount of available space. It is wise to have several replacement microswitches on hand if your incubator’s control system includes one, because high humidity levels cause microswitches to stick after a period of use, and a sticking switch can cause damage to incubating eggs through overheating. Once a microswitch begins to stick, replace it. It may seem that working the switch mechanism repeatedly frees it, but it will stick again within a short period of further use. Most people who work with chick embryos agree that incubating eggs should be turned at least once daily. Of course, eggs that have been opened for surgery or grafting cannot be disturbed after they have been returned to the incubator, and they should not be turned.

221


Appendices



It is very helpful to have, and use, a candler to sort out sterile eggs, find dead experimental embryos for removal from the incubator, and to demonstrate structural relationships to students. Candlers sometimes can be purchased from farm-supply companies. Your state university’s department of poultry science may be able to help you locate a local source.A candler can also be constructed as follows. Make a circular hole about 4 cm (11兾2 inches) in diameter in the bottom of a large can such as a threepound coffee can. Mount a light-bulb socket on a base that is small enough so that you can invert the can over it. A 100-watt bulb usually works well for candling, but the can will become quite hot to the touch after it has been in use for a few minutes. It is wise to warm students about this hazard. It is also wise to glue a ring of cork around the hole in your candler to keep eggs from touching its hot surface. See comments on candling in Section D following. The vigor and viability of chick embryos varies from one time of year to another. Spring is the best time for work with chick material, while work done in late summer or very early autumn usually is less satisfactory. If chick embryo work can be delayed until after October 15 in fall courses, you will find the results more satisfactory. Living embryos of the classically described stages are usually obtained only after longer incubation times, but you should test this factor in your own laboratory with your own egg supply.The following are approximate incubation times at 38° C (100.4° F) used in my laboratory in the fall of the year: Classical Stage “24-hour” “33-hour” “48-hour” “72-hour”

chick chick (12–15 somites) chick chick

Incubation Time 30 hours 38–42 hours 55–60 hours 75–82 hours

You may find that embryos will reach these stages after somewhat shorter incubation time at other times of the year, but the only conclusive answer will come when you test eggs from your own supplier under your own laboratory conditions. B. Saline Solution I prefer Howard Ringer’s solution when an isotonic saline solution is required for use with chick embryos.The formula follows: Howard Ringer’s Solution NaCl CaCl2 (2H2O) KCl Distilled water

7.2 g 0.17 g (0.23 g) 0.37 g 1000 ml

For some routine uses, such as brief examination of living embryos that will be discarded after use, 0.9% NaCl is adequate. There are several other chick saline solutions in use in various laboratories, and formulae for several of them can be found in Rugh (1962), Hamburger (1960), and Stern and Holland (1993) (see p. 143). Generally, you will obtain better results if the saline solution is warmed to around incubation temperature before it is used with embryos.

222

Appendix D


Appendices



C. Instruments and Miscellaneous Materials 1. Instruments Cactus-spine microsurgical needles (England, Marjorie, J. Microsc. 123:133–146; 1981) are very useful for a number of applications in the developmental biology lab, and I recommend them for use with chick embryos.To obtain spines, select a cactus plant that has a mix of spine sizes and use a forceps to collect a number of spines of various sizes. Ordinary wooden applicator sticks (for example, Fisher Scientific, catalog # 01–340) make good handles for the needles. Split and slightly sharpen the ends of sticks with a single-edged razor blade, and use clear nail strengthener to glue a spine in place into the split end of each stick. ( We use Revlon Strong Wear, which is also useful for sealing wet-mounted slides and other lab applications. Several other similar products seem to work as well, as does New Skin Liquid Bandage.) A 45° angle of spine to stick makes a good, practical working needle, but students might find that they prefer other arrangements after they have used the needles for a while. Cactus-spine needles can be sterilized by dipping them in alcohol and letting them air dry, but do not flame sterilize! Directions for preparation of the tungsten microsurgical needles and information about watchmaker’s forceps are given in Appendix B. Glass microsurgical needles may also be used, but students sometimes find them difficult to see against the background of a chick embryo, especially when working through a small hole in the shell. Transfer (chick) pipettes are made by scoring, breaking, and fire polishing the tapered portions of ordinary medicine droppers to produce a finished “chick pipette” with an inside diameter of 3.5 to 5.0 mm. 2. Egg Nests Egg nests that are used to hold eggs during grafting or embryonic surgery can be made of the soft Styrofoam used to make pillows. About 22- or 23-cm-long strips of 1-cm-square Styrofoam are bent and stapled into a loop to make simple egg nests that are more practical than elaborate egg holders such as cotton nests in Syracuse dishes. Eggs are left in these Styrofoam nests when they are returned to the incubator following treatment. Sometimes, operated eggs can be handled more easily and more gently if the egg nest and egg are set on half of a petri dish. This also prevents fouling of the incubator by poorly opened or accidentally cracked eggs, but the petri dishes do occupy more incubator space than do eggs in egg nests alone. 3. Egg Saws Hacksaw blades can be sharpened to make very effective “egg saws.” I routinely have used 23 teeth to the inch hacksaw blades, but a variety of other types of blades likely would serve as well.The blades must be ground on a stone wheel until the teeth no longer protrude outward to the sides, but rather are a straight row of thin, sharp teeth.This is very dangerous work that must be done only with proper eye protection and should be undertaken only by someone familiar with the use of power tools. Many people prefer to open eggs with Dremel tools, using the flat cutting wheels. 4. Instrument Jars Instrument jars are prepared by placing a layer of compressed absorbent cotton in the bottom of a small glass jar such as a baby-food jar.The concentration of the ethyl alcohol in the jars must be maintained at higher than 70%, or it will become difficult to ignite the alcohol when instruments are flame sterilized. Empty baby-food jars also make acceptable holders for the handle ends of tungsten, glass, or cactus-spine microsurgical needles if you don’t wish to prepare blocks of wood drilled with holes of the appropriate sizes.

Appendix D

223


Appendices



5. Alcohol Lamps While alcohol lamps can be constructed, relatively inexpensive lamps are available from commercial suppliers. Fifty percent alcohol can be ignited, but, in practice, alcohol lamps are easier to light, and they work much more efficiently when filled with 80 to 90% ethanol. In fact, alcohol concentrations below 70% usually cause problems in alcohol lamps. D. Candling Candling should be done in a dark or very dimly lit room, but sometimes candling may be done inside some sort of “dark box” if it is not convenient to take eggs to a room that can be darkened. There is no substitute for practice in candling eggs, but there are a few points that might be helpful if your experience with candling is limited. These comments describe white-shelled eggs (for example,White Leghorn) but can be applied to breeds with more shell pigmentation. However, the greater opacity due to shell coloration makes it more difficult to pick out details in candling. By the end of the second day of incubation, it is possible to see some evidence of blood vessel development. However, due to the great variability in developmental stage reached by this time, it is wise to be very conservative about discarding eggs that do not appear to be developing. At the end of the third day of incubation, the spidery network of embryonic and extraembryonic (yolk-sac) blood vessels is obvious even in eggs with pigmented shells, although there may be some problems with very mottled shells. In white-shelled eggs, it is usually possible to pick out the embryonic body and to distinguish the head fairly readily. The sinus terminalis (the peripheral boundary of the extraembryonic circulation) is often faintly visible. By candling on succeeding days, you will note the spreading of the vitelline circulation of the yolk sac and the establishment of the chorioallantoic circulation.The embryo becomes more fixed in its position relative to the shell and shell membranes and no longer rotates freely in response to gravity when the egg is turned. By about the fifth or sixth day of incubation, you will notice that the embryo appears to be moving about slowly.This movement actually is due to slow, rhythmic contractions of the amnion that apparently serve to keep the embryo moving about in the amniotic fluid.This prevents adhesions between the embryo and the membrane. Only later will you observe the quick, jerky movements of the embryo itself as neuromuscular coordination begins to become functional. One fairly certain sign that an embryo has died is the presence of a “blood ring.”The blood ring is caused by the settling of blood in the sinus terminalis area at the same time that the dead embryo begins to degenerate. This dense, reddish ring without any visible structure in the center is a reliable index of embryonic death from about the fourth or fifth day of incubation onward. After 10 days of incubation, the embryo is quite prominent and has become opaque. It appears as a dark, moving body when the egg is candled. By this time, it is possible to distinguish vitelline from chorioallantoic vessels.When the egg is rocked, the chorioallantoic vessels which are closely applied to the inner surface of the shell membrane remain in a fixed position. On the other hand, the vitelline vessels of the yolk sac appear to move about as the yolk shifts slightly in response to rocking. After 12 or 13 days, the embryo appears as a very dark object, and following 2 or 3 additional days of incubation, further growth of the embryo makes the egg practically opaque when viewed from some directions.The air space enlarges during incubation, and by later stages, it is prominent and very sharply demarcated in eggs with living embryos. Frequently, it is possible to examine the condition of the extraembryonic blood vessels that normally appear as continuous dark lines if eggs are tipped or rotated over the candler. In the case of embryos that have died at advanced stages, these vessels appear as broken or irregular lines, but this distinction is sometimes hard to make.With luck, it is also possible to detect the jerky movements of the living embryo, but this becomes almost impossible during the final days of incubation.

224

Appendix D


Appendices



Finally, it is well to be very conservative about discarding eggs that contain operated embryos on the basis of candling observations because they are often smaller than control embryos of the same age. As it is possible to be misled by candling, it is best to leave questionable eggs in the incubator for another day or so. Normally, even eggs containing dead embryos decompose only very slowly, and it is unlikely that they will emit enough noxious gases either to be unpleasant to experimenters or to cause damage to nearby embryos in the incubator for at least several days after death. However, the “sniff test” is often the surest way to assess the condition of an embryo late in incubation! Egg Suppliers Murray McMurray Hatchery, 191 Closz Drive, Webster City, IA 50595 (515–832–3280, 1–800–456–3280; website: www.mcmurrayhatchery.com) ships fertile eggs from January to late May. McMurray Hatchery also sells incubators. Information on various regional hatchery companies may be available from state agricultural extension services.

Appendix D

225


Appendices




Appendices

E: Preparation of Materials for Experimentation on Fern Gametophytes


A P P E N D I X

E Preparation of Materials for Experimentation on Fern Gametophytes

A. Procurement and Storage of Spores The Materials sections of Laboratories 14 and 15 specify spores of Pteridium aquilinum, Woodwardia sp., or other fern species whose gametophyte development begins with a linear chain of cells (a protonema). Biological supply companies have been helpful to me in supplying information about the spores that they offer and, in some cases, have been able to supply spores of a particular species, such as Pteridium aquilinum, upon request. Suppliers in the eastern United States probably will supply Woodwardia sp. routinely, but it is important to inquire when ordering spores because this may change depending upon availability. Sometimes a supplier will specify simply that initial growth is linear. Pteridium aquilinum spores remain viable for long periods if they are stored in screw-cap vials and kept in a refrigerator. I have successfully used Pteridium spores in developmental biology labs after 5 or more years of refrigerated storage. However, in my experience, the spores of some other species have not stored nearly so well, and in some cases, have lost viability after a year. Unless you have obtained recently collected Pteridium spores and have stored them carefully, I recommend purchasing a fresh supply of spores each year to avoid problems and disappointments in the lab. Fern spore collecting is an interesting and satisfying experience, and if Pteridium aquilinum (the bracken fern) grows near you, with luck you can easily collect a supply that will last for several years of teaching developmental biology.The bracken fern is a widely distributed species, and in many areas, it should be fairly easy to find fern fronds with ripe sori if the plants are checked periodically throughout the growing season.The spores themselves may be collected in several ways. A commonly used technique for spore collection is to place fern fronds on large sheets of smoothsurface paper in an area free from any drafts that might blow spores away as they fall. If the fronds are left undisturbed overnight (spore drop may continue for several days if the fronds can be left that long), spores will be found on the paper and can be brushed into vials using a camel’s hair brush. Spores can be filtered to remove debris, but this step is not essential. Spores have also been collected by shaking fronds vigorously in plastic bags and passing the materials obtained through a set of soil sieves. Parts of Laboratory 15 involve experiments on heart-shaped prothalli and young sporophytes at the first-leaf stage. It is usually more convenient to order these stages from a supply company than it is to grow them and have them available at the right stage of development on the particular date when they are needed for the lab.

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E: Preparation of Materials for Experimentation on Fern Gametophytes

Several biological supply companies regularly offer Marsilea sporocarps. However, it is important to specify small, hard, smooth, nonhairy sporocarps. There are significant species differences in sporocarp features, and sporocarps from species that produce the larger, hairy type do not yield the same results as the smaller ones.The gelatinous ring is slower to emerge when the sporocarp is cut, and often it does not emerge completely,if at all.Also,timing of subsequent developmental events is less predictable. Ward’s, P. O. Box 92912, Rochester, NY 14692 ( Tel. 1–800–962–2660; website: www.wardsci.com) were supplying the small, hard sporocarps at the time of this writing. B. Media The relatively simple inorganic medium suggested by Davis and Postlethwait has the following formula:

Basic Fern Medium NH4NO3 KH2PO4 MgSO4 • 7H2O CaCl2 • 2H2O Ferric citrate Distilled water

0.5 0.2 0.2 0.1 5 1000

g g g g mg* ml

*0.1 g of ferric citrate should be dissolved in 100 ml of boiling distilled water and, after cooling, 5 ml of the solution should be added to the medium. The ferric citrate stock solution may be stored for several months in the refrigerator.

It is not essential that this medium be sterilized, but autoclave or filtration sterilization might reduce possible microbial contamination in cultures after long periods of growth. Mineral agar medium is prepared as 0.5% (w/v) plain agar (not nutrient agar) in basic fern medium. There are a number of other salt solutions that can be used for growth of fern spores. One of these is Bold’s Basal Medium ( H. C. Bold, A Laboratory Manual for Plant Morphology, 5th ed., Harper & Row; 1986). Formulae for Bold’s medium and another suitable medium are given by Chilton and Graham (1988; see p. 171 for citation). Dyer (1983; see p. 171 for citation) gives another. Even spring water will sometimes suffice. C. Culture Conditions and Comments There are comments on fern gametophyte culturing in the Techniques section of Laboratory 14, but several other general suggestions might be helpful. Generally, cultures are disrupted less if plants are removed with bacteriological loops rather than pipettes. Very bright illumination induces early transition to two-dimensional growth. If plants become twodimensional very quickly under your culture conditions, it is likely that the light intensity is too high. In the photomorphogenesis experiments, red cellophane taped over a fluorescent lamp usually will serve adequately as a “red-light” source if “white-light” leaks are prevented. However, you should pretest your particular growth conditions. Filters that transmit light in a well-defined, narrow band of wavelengths are available commercially, but these experiments should work well even under relatively crude conditions. Make certain that any confined area used for parts of these experiments does not get too warm.

228

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F: Prep of Materials for Exp. on Emerging & Regenerating Developmental Patterns


A P P E N D I X

F Preparation of Materials for Experimentation on Emerging and Regenerating Developmental Patterns

Consult the Materials section at the end of each Laboratory for a list of the materials needed for that lab. This appendix provides information on sources of organisms and formulae and information on acquiring or making listed materials that are not standard stockroom items. A. Spirostomum, Hydra, and Planaria Spirostomum, hydra, and planaria can be obtained from several major supply companies. Some sources indicate that certain types of hydras or planaria are better for regeneration studies than others, but there doesn’t seem to be general agreement on this point. Some of the more definite statements concern planaria (such as Hamburger 1960, cited on p. 94). I have not seen any references to the regenerative capacities of white planaria, so it might be best to plan to work with the brown varieties. When any of these organisms arrive, the jars that they are in should be opened as soon as possible. The medium probably is somewhat oxygen depleted, and it is wise to aerate the cultures immediately by bubbling air through them with a pipette. The organisms usually will remain in better condition if they are kept considerably cooler than room temperature. One way to do this is to set the loosened caps back on culture jars (do not screw on) and place the jars in a shallow pan of water set in a sink. Then, let water drip into the pan from the cold water faucet. Spirostomum can be transferred with the plastic pipettes usually supplied with the cultures, but these huge cells probably will fare better if they are not transferred with Pasteur pipettes that have an inside diameter of 1 mm or less. Hydras can be transferred with fairly small-mouth pipettes, but damage is less likely if pipettes with internal diameters of 2 mm or more are used. Plastic pipettes can be cut at the level necessary to give such an opening or glass pipettes can be cut and fire polished. Sometimes it is necessary to “scrape” a hydra loose from the culture jar side or bottom by nudging it very gently with a pipette.The hydra will respond by contracting its body very strongly and will be easy to transfer in that condition. Usually, after a few minutes in a dish, a hydra will attach to the bottom or side of the dish, relax, and extend its body column and tentacles. Pipettes with an inside diameter of about 3 mm work adequately for transferring planaria, but the animals are squeezed a bit by such openings. Some people prefer to transfer planaria with camel’s hair brushes, but there is some danger that they will be stabbed by the bristles. A planarian can become lodged inside a pipette if it has time to do so. This problem can be avoided by making transfers quickly. Curved scalpel blades are useful for planarian regeneration experiments. No. 10 (smaller) or No. 24 or No. 22 (larger) blades are good choices.

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It is sometimes difficult to determine what you are buying when you set out to buy “spring water.” Some stores offer “drinking water” as a substitute, but careful label reading may reveal that the contents are simply deionized water, which will not sustain living Spirostomum, hydra, or planaria.Tap water in most areas is fatal for all three, and distilled or deionized water will not work. If you are not certain about an appropriate water source, you might wish to make up an “artificial pond water,” which some colleagues recommend. I have not used it myself, but I trust the judgment and experience of the people who have suggested its use.

Artificial Pond Water Solution A:

NaCl 133 g CaCl2 26.6 g In 1 liter distilled water Solution B: NaHCO3 3.8 g In 1 liter distilled water Mix 2.5 ml of each with 995 ml distilled water.

If your students want to offer food to regenerated hydras, you might wish to supply some brine shrimp (Artemia) larvae. Brine shrimp eggs (encysted, dormant larvae) are available from biological supply companies and from local aquarium stores. Brine shrimp larvae begin to emerge between 1 and 2 days after the cysts are scattered on the surface of a saline solution. We have obtained good results using a 70% seawater solution ( 7 parts artificial seawater: 3 parts distilled water). If you shine a light from a source such as a gooseneck lamp on one side of the culture dish, the culture stays warmer, and the larvae tend to congregate on the lighted side. They can be pipetted and transferred to a small volume of spring water from whence they can be transferred to the dishes containing hydras. This washing step reduces the amount of salt that is transferred into the hydra cultures. B. Lumbriculus Charlie Drewes of Iowa State University, who has worked out several teaching applications using Lumbriculus, has collaborated with Carolina Biological Supply (address information, p. 217) who now can supply Lumbriculus in kit form. Here are two other reliable Lumbriculus suppliers: Aquatic Research Organisms P.O. Box 1271 One Lafayette Road Hampton, NH 03842 Phone: 1–800–927–1650 (603) 926–1650 Website: www.holidayjunction.com/aro

Environmental Consulting & Testing 1423 N. 8th Street, # 118 Superior, WI 54880 Phone: 1–800–377–3657 (715) 392–6635 Fax: (715) 394–7414 Website: www.ectesting.com

When ordering from Aquatic Research Organisms, specify “Living Diet Lab” for the best quantity/ price ratio. Lumbriculus is coming into use for fish food in lieu of Tubifex, so they may be available from additional sources. Lumbriculus is sometimes identified as “California blackworms,” but some caution is advised because there is often confusion about precise identification of small aquatic annelids, and the regenerative responses of various species differ considerably. Some suppliers may have minimum order policies that could leave you with a number of extra worms on hand, but it is possible to maintain Lumbriculus cultures with only a small amount of regular care. To begin a Lumbriculus culture, cover the bottom of a large finger bowl or a small fishbowl with finely shredded paper towels and add spring water to a depth of 7 to 8 cm. Supply the bowl with an air line for very slow bubbling and cover it after adding the worms. About once a week, add a couple of sinkable fish food pellets, trout chow pellets, or adult Xenopus food pellets to each bowl. Add distilled or spring water to replace any lost by evaporation. If the water becomes foul, pour most of it 230

Appendix F


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off and replace with spring water. Lumbriculus sometimes reproduces ( by fission) very vigorously under these culture conditions, so it is necessary to subculture occasionally if a culture becomes crowded. C. Cactus-Spine Microsurgical Needles and Iris Scissors Fine needle cactus-spine microsurgical needles work well in the Spirostomum experiments. Information on making these needles is given on p. 223 in Appendix D. Iris scissors can be purchased from biological supply companies and from various other instrument and lab supply companies.They are available in either straight or curved blade models, and either type works for hydra and Lumbriculus regeneration experiments. Students need to be cautioned about the care needed in use of these scissors to avoid damage to the scissor tips. D. Pollen and Seeds Pollen can be obtained from a variety of flowers found in florist shops or gardens. Pollen from tulips or lilies often germinates and grows well in the lab, as does pollen from alstroemaria.Various colleagues have suggested the following plants as sources for pollen that will produce pollen tube growth in culture: Tradescantia, Impatiens, tomatoes, clover (Melilotus), and the bridal veil plant (Gibasis geniculata). I have not tested any of these personally but am confident in the advice of these colleagues. I have obtained excellent results with lupine pollen, which unfortunately is available for only a brief season in nature and in gardens. It is likely that pollen from other flowers will also germinate and grow in the medium suggested here, but pollen from some plants may not. Thus, it is important to test pollen before using it in lab. Seeds are available from many sources, including local garden seed suppliers, as well as from the major biological supply companies. Sweet corn seeds are relatively inexpensive and can usually be purchased in larger quantities than many other seeds. I firmly recommend the “Early Alaska” variety of pea seeds for experiments on root regrowth. If students are to go on to investigate other plants, sunflowers and a number of squash and pumpkin varieties, as well as many bean varieties grow well under these conditions.Students especially enjoy growing peanut seedlings, but if they work with peanuts, they should be strongly cautioned that seeds supplied for planting often have been treated with very toxic substances that repel insects and retard mold growth. It would be very dangerous to eat seed peanuts. ( They are raw, unroasted peanuts anyway.) E. Seed Germination Techniques Seed germination is conveniently accomplished by what Carolina Biological Supply Co. calls the “rag doll method” in their very useful technical brochure called Plants from Seeds. Cut two pieces of waxed paper to the approximate size of ordinary, folded paper towels. Lay two soaked paper towels on one of the pieces of waxed paper. Then place seeds at about 11//2 - to 2-cm intervals in a lengthwise row on the wet towels. Cover the seeds with another wet paper towel. Roll up the stack and set it in a container (for instance, a baby-food jar or a beaker) containing water to a depth of about 1 to 11//2 cm. ( It may be necessary to loosely secure the rolled stack with a rubber band to keep it rolled.) Put the jar in a dark place and daily add water to maintain the level in the jar. Since the timing of root emergence is affected by several factors, it is essential that seed germination be pretested under your particular laboratory conditions. You can accelerate germination of most seeds by soaking the seeds overnight before setting up the “rag dolls.” (But don’t soak bean seeds for more than a couple of hours.) Mold growth on germinating seeds sometimes becomes a problem. Surface sterilization of the seeds before beginning germination helps to prevent mold growth. One convenient method for sterilizing seeds is to place them in a 1:8 (volume) dilution of 5.25% sodium hypochlorite ( household bleach). After the bleach treatment, wash the seeds in at least two changes of water before proceeding. Appendix F

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F. Solutions

Pollen Germination Medium Solution A* Sucrose 10 g H3BO3 0.01 g Ca(NO3)2 • 4H2O 0.03 g MgSO4 • 7H2O) 0.07 g KNO3 0.01 g Distilled water to make 100 ml of solution

Solution B** Sucrose H3BO3 Ca(NO3)2 • 4H2O

10 g 0.01 g 0.03 g

Distilled water to make 100 ml of solution

*Source: Tatina, R.; Hohn, K. Amer. Biol. Teacher, 56:174–175; 1994. **Source: Schimpf, D. J. Amer. Biol. Teacher, 54:168–169; 1992.

Toluidine Blue O (TBO) Staining Solutions 1. For staining tissue squashes, 0.5% TBO in distilled water 2. For staining hand-cut sections, 0.05% TBO in distilled water

G. Preparing and Staining Root Squashes This technique is adapted from work of Joseph J.Nickolas,Northland Community College,Holbrook, Arizona. 1. Bring water to a boil in a 100-ml beaker on a hot plate. Be sure to use boiling stones or chips to prevent popping. 2. Place the fragment of root tissue to be squashed in the center of a slide and put several drops of 1N HCl on it. Set the slide across the top of the beaker of boiling water.This hot acid treatment fixes (preserves) the cells and softens the tissue. Continue this treatment for 11//2 minutes. Use a dissecting needle to roll the root fragment around in the acid in order to consistently ensure complete fixation and softening. Add more acid as needed to prevent drying. 3. Carefully remove the slide from the beaker and use blotting paper or a paper towel to blot away the remaining acid. Immediately add several drops of Toluidine Blue O ( TBO) stain. Make certain that the root tissue is immersed. 4. Return the slide to the top of the beaker and heat it for about 11//2 minutes. Roll the root fragment around with a dissecting needle as in Step 1. Add more stain as needed to prevent drying. 5. Remove the slide and cautiously blot off the excess stain to avoid losing the root tissue, which is hard to see when immersed in the stain. Immediately put a drop of fresh stain on the root tissue. Add a coverslip over the root tissue and stain solution. 6. You should now “squash” the softened, stained tissue to spread and flatten it for examination with the compound microscope. Set the slide on a paper towel and put a folded paper towel on top of it. Press down with your thumb directly over the coverslip. Press down firmly (so that a little “white” shows around the edge of your thumbnail), but try not to twist your thumb. 7. Hold your slide up to the light. A light blue color in the area of the squashed root tissue indicates a wellstained and squashed preparation. 8. Return your slide to the top of the beaker of boiling water and leave it there for 1 minute. This step helps to clear stain that is not bound to root tissue cells. 9. Remove your slide from the beaker. After you have wiped the bottom of the slide, scan over it with the low-power objective of your microscope.When you have located the stained root tissue, switch to high power for your examination of cell characteristics and identification of dividing cells.

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H. Preparing and Staining Hand-Cut Sections With care, patience, and a little practice, usable sections of developing roots can be cut using singleedged razor blades. It is important to be very cautious in handling razor blades in order to avoid injury. Place the root to be sectioned on a flat surface and try to cut the thinnest possible cross sections of the area of the root you wish to examine. Some workers suggest that sections are easier to cut if an oblique section is cut by slicing down and away from the hand that is holding the root, but my students prefer to cut straight downward across the root. Sections can be stained using the following procedure*: 1. Place the section, or sections, in tap water and allow them to soak for 2 to 3 minutes. 2. Transfer sections to 0.05% Toluidine Blue O staining solution and leave them there for 1 minute (the time may need to be adjusted through trial and error). 3. Transfer the stained sections to tap water for a 2-minute rinse. 4. Mount the section on a slide in a drop of water and cover it with a coverslip. 5. Examine the section using the low power of a compound microscope. Be cautious about switching to high power because the preparation may be quite thick and the objective might contact the coverslip.

Alternatives and Additions All the steps of the preparation process can be done on a slide by adding drops and then wicking away solutions with a paper towel. This makes it easier to keep track of the sections, but they can also be accidentally washed or brushed off fairly easily. A preparation that lasts for up to a day or so if refrigerated can be made by sealing the edges of the coverslip. Revlon Strong Wear Nail Strengthener works quite well for this, as do several other similar products.

*Source: Parker, A.J.; Haskins, E. F.; Deyrup-Olsen, I. In Plants in the Laboratory. Nat. Assoc. Biol. Teachers, Monograph Series I; 1984.

Appendix F

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Appendices

G: Notes on Course Scheduling and Use of the Manual


A P P E N D I X

G Notes on Course Scheduling and Use of the Manual

There are many factors to be considered in organizing a course that depends on the availability of living material, especially when some of the organisms involved reproduce seasonally. Also, important biological and ethical questions arise from concerns about the use of animals in the teaching laboratory. The following sections contain suggestions about animal use and conservation, comments on humane sacrifice of animals, suggestions on course scheduling, and some miscellaneous hints about developmental biology labs. A. Availability of Organisms Some seasonal reproductive factors affect the sequence of topics in a developmental biology laboratory course, especially during the fall term.There is a period in the fall when summer-breeding sea urchins are nearing the end of their breeding season and winter breeders cannot yet be induced to shed gametes (see Appendix A). It is important to consult carefully with suppliers regarding urchin usability. During October, and especially early to mid-November, it is wise to be cautious about scheduling sea urchin labs. In planning for sea urchin labs late in the fall term, inquire about possible interruptions in shipping around Christmas. Some suppliers hesitate to provide assurance of prompt delivery during the preholiday period. It might be best to complete spring work on sea urchins by mid-April. Some suppliers are optimistic (without justification) about year-round use of Rana pipiens, but developmental biology labs undertaken with Rana during September and early October generally yield poor results. Xenopus, however, is usable year-round, though some workers report a period of somewhat reduced reproductive vigor during the summer months. There are some problems with vigor and viability of chick embryos during late summer and early fall, but good results usually can be obtained with chick embryo work after October 15. All of the labs on invertebrate animals can be done at any time of year, as can the labs on fern and flowering plant development. However, it is important to keep track of the age of fern spores (see p. 227) and flowering plant seeds. It usually is not worthwhile to use old seeds or old Woodwardia spores, but refrigerated Pteridium spores store well for long periods, and Marsilea sporocarps remain viable for many years.

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B. Using the Various Labs Most of the labs can be completed, or at least initiated, within one weekly 3-hour lab period, but a number of them require that students return one or more times to make further observations. Several of the labs require an essentially “open-lab” approach. However, with careful planning, fairly limited lab access can prove adequate. It is preferable for students to learn the basics of sea urchin methodology and follow the wonderful developmental sequences ( Laboratory 1) during a first week and then use this background to turn to experimentation ( Laboratories 2 and 3) during a second week. Because of the costs involved, however, it may be necessary to combine Laboratory 1 with some elements of Laboratories 2 and 3 for a single week of sea urchin observation and experimentation. With some careful planning and organization, this can be fairly satisfactory. The material in Laboratory 5, Patterns of Frog Development, can be squeezed into one period, but it is sometimes more comfortable to spread this lab over two periods. Laboratory 10, Patterns of Chick Development, requires more than one period for completion. Some instructors split it into three periods, working up through Section D for the first period, doing Section E during a second period, and then finishing the remainder of the lab during a third period. It is possible, however, to complete Laboratory 10 within two 3-hour periods. The fern prothallus development labs ( Laboratories 14 and 15) require many repeated observations after a fairly short set-up period.They actually are best done independently, outside of regular lab periods, because the work consists almost entirely of individual observations. The fern work is best started early in the term so that students will have time to repeat or extend experiments if they wish to do so. Some major events in sexual reproduction in Marsilea occur within 8 to 10 hours. After the first day, development of the young sporophyte plant can be followed by observations made once each day. Students definitely should have a chance to see the emergence of the mucilaginous ring from a Marsilea sporocarp.This is an impressive event! The two labs on Emerging and Regenerating Developmental Patterns ( Laboratories 17 and 18) both are diverse in terms of the time required for significant events to occur. Spirostomum reorganization is completed within a time span of a few minutes to an hour. Hydra regeneration takes several days and requires daily checking. Lumbriculus regeneration requires 10 to 15 days, so it is convenient to check it at each of the next two lab meetings after the lab period in which the segments are cut. Planarian regeneration takes several weeks for completion. Pollen tube outgrowth begins within 1 to 2 hours, if it is going to occur at all, and maximum growth is often achieved by the end of 3 hours. Beginnings of a gravitropic response in sweet corn can be detected within the span of a 3 hour lab, and the response is very evident by the next morning. ( These rapid developmental responses by plants impress even students who are committed animal chauvinists!) Root regeneration requires 5 to 7 days, so it can be followed on an open-lab basis or results can be observed at the next weekly lab meeting. C. Variations and Student Projects This manual contains a mix of descriptive and experimental labs and includes work on some familiar animal systems, some less well-known systems, and several labs devoted to study of plant development.The manual can serve as the sole source of lab work for a developmental biology course, but I presume that many or most instructors will also want to use their own favorite labs along with a mix of labs from this manual. As another alternative, the manual may serve for the basic labs in a course in which students undertake extensive investigative projects. I often follow the latter plan in my own course. I find it useful to begin my course with an informal review lab that is not included in this manual. This lab reviews some aspects of plant reproduction and development and a few basics of animal development. After a flower dissection, students review female gametophyte (embryo sac) development, pollen tube growth and male gametophyte differentiation, fertilization, embryo development (using Capsella slides), and some basics of seed germination and seedling growth. Most of my students need

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a review of these introductory biology topics. Generally, students are better able to recall essentials of animal reproduction, but a brief review of some aspects of development, such as cleavage and gastrulation using prepared slides of sea urchin or starfish embryos, is usually helpful. Such an introductory lab also provides a framework for an initial review of the intriguing basic genetic problem of cell differentiation—nuclear genetic equality but diverse structural and functional differentiation—that students probably have considered in introductory biology. I hope that, in addition to student projects that the instructor might suggest, students will be able to develop projects of their own after consulting the various “Suggestions for Further Investigations” sections and some of the references cited therein. In making time for student projects, there are many ways to economize on lab periods. For example,Laboratory 11 (in vitro culture) can be combined with a second chick descriptive lab,and chorioallantoic grafting and/or surgery can be combined with a third chick descriptive lab. The results of either of these experiments can be obtained after a week, and observing the results will not take a great deal of time away from the lab at hand. Similar selective combining of labs in other parts of the manual can help to make available free lab periods for project consultation sessions or for other in-lab activities, while still drawing on the basic set of lab experiences from this manual. D. Preparation Hints Labs generally require greater quantities of solutions than expected. For example, after making a reasonable estimate of the amount of artificial seawater or chick saline needed, it is often wise to prepare a 50% greater quantity. Things go better when students are made individually responsible for care and maintenance of their own instruments and equipment.The same can be said for lab and glassware cleanup. I think it is important for students to learn how to clean glassware to a standard required for use with cells and embryos and to maintain a clean, functional work space. I have found that a complete laboratory preparation notebook, one that is revised as it is used, is a valuable supplement to the appendixes in this manual and that it saves much time and effort for the instructor and any teaching assistants who might work in the course. E. Humane Sacrifice of Animals An increase in respect and concern for living things is a desired outcome of education in biology. However, personal growth and maturation in this area is difficult for students because they are regularly confronted with extreme viewpoints. On one hand, a callous disregard for nature defines all living things only in terms of their “value” to humans, while on the other hand, there are emotional and often highly publicized assaults on all animal use in biomedical research and teaching. In this context, concern about humane sacrifice of animals is a vital part of teaching, wherever sacrifice proves necessary. Invertebrate animals can be humanely sacrificed by chilling and/or freezing. Adult sea urchins, for example, can be wrapped in aluminum foil and left in a freezer compartment overnight. Up to about 8 days of incubation, chick embryos can be sacrificed by quick decapitation, based on the assumption that sensory function and neural responses are still limited, if developed at all. At later stages of incubation (for example, 16 to 17 days, as with host embryos of chorioallantoic membrane grafts in Laboratory 13), it is wise to chill embryos thoroughly before decapitation. This can be done by holding them under running, cold tap water until they are chilled (and your fingers are numb) before decapitation. Adult male Rana pipiens can be sacrificed by brain and spinal pithing. Xenopus males are usually sacrificed by a sharp blow to the head, decapitation (either complete decapitation or removal of the head above the mouth), and spinal pithing. It is possible to chill-anesthetize Xenopus by adding ice to the water around them or simply by leaving them in chilled water in the refrigerator for several hours. After chilling, the other steps are less traumatic. Many people prefer to induce very deep ether anesthesia or to sacrifice males using chloroform.

Appendix G

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F. Amphibian Conservation During the 1980s and 1990s, many biologists called attention to the problem of declining, and even disappearing, populations of many amphibian species (see references that follow). Supply companies will assure you that there is no problem with regard to Rana pipiens populations, but that may not be completely accurate. Collectors are ranging far and wide to meet the demand for Rana pipiens. Thus, strategies for teaching developmental biology without sacrificing adult Rana pipiens might be attractive to many instructors. One answer is to use Xenopus wherever possible. Xenopus adults, which are reared in great numbers in captivity, are also being substituted for Rana in nerve and muscle physiology laboratory experiments (Bernhart et al 1991). However, some instructors may wish to teach developmental biology without sacrificing any adult vertebrate animals. If you wish to consider these strategies, please proceed to the next section. References Bernhart, D. M. et al. Conservation in the teaching laboratory—substitution of Xenopus for Rana. BioScience 41:578–580; 1991. Blaustein, A. R.; Wake, D. B.The puzzle of declining amphibian populations. Sci. Amer. April: 52–57; 1995. Griffiths, R.; Beebee,T. Decline and fall of the amphibians. New Scientist 27 June: 25–29; 1992. Livermore, B. Amphibian alarm: just where have all the frogs gone? Smithsonian October: 113–120; 1992. Pechmann, J. H. K. et al. Declining amphibian populations: the problem of separating human impacts from natural fluctuations. Science 253:892–895; 1991. Philips, K. Where have all the frogs and toads gone? BioScience 40:422–424; 1990. Wake, D. B. Declining amphibian populations. Science 253:860; 1991.

G. Teaching Developmental Biology Without Sacrificing Adult Vertebrate Animals Working with living, developing organisms brings developmental biology to life for students, but those parts of the courses that require sacrifice of vertebrate animals may be viewed as too biologically or philosophically costly. Chick embryos usually do not bring the issue of vertebrate animal sacrifice into focus because it can usually be agreed that since a chick embryo has already begun development when an egg is laid, the embryo certainly would die if the egg were not incubated (in some cases, embryos would be boiled, fried, or scrambled). Incubation in the teaching or research lab allows development to proceed and may introduce the issue of humane sacrifice (p. 237), but use in a developmental biology course does not fundamentally alter the fate of a chick embryo. In developmental biology laboratories, the sacrifice of adult frogs raises the animal sacrifice issue most directly and dramatically. In one sense, there is no completely consistent philosophical justification for sacrifice of Rana pipiens adults for teaching, although the value of the educational experience does carry some philosophical weight. The situation with Xenopus is quite different. Xenopus is raised for laboratory use and thus might be considered in the same category as animals raised for food.However, this certainly does not obviate all philosophical objections. Here are some specific suggestions regarding teaching developmental biology without adult vertebrate animal sacrifice: 1. Use preserved Rana pipiens embryos for external views of early developmental stages. Despite some shrinkage artifacts, embryo sets supplied in small screw-cap jars are very useful. (Admittedly, an adult male had to die somewhere to provide the embryo set, but one sacrifice probably was adequate to supply embryos usable for a number of years in several biology departments.) 2. Concentrate on experiments that can be done with embryos obtained from Xenopus matings rather than in vitro fertilization. This actually eliminates only three experiments from among all those available in the various labs on frog development. These experiments are (1) UV irradiation of sperm (a part of Laboratory 4);(2) Laboratory 8, which requires UV irradiation at the two-cell stage; and (3) chilling of precleavage zygotes (one of the three experiments in Laboratory 9).

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Appendices



3. You might consider trying a less orthodox technique suggested by McKinnell et al (1976) for obtaining frog sperm without sacrificing male Rana pipiens. ( I don’t know (1) if others have had success using this technique with Rana or (2) if it has been tried with Xenopus.) Basically, the protocol they used is to obtain urine that contains sperm by intraperitoneal injection of male frogs with 100 I.U. human chorionic gonadotropin.The frogs were also given 1 ml of sterile distilled water by intraperitoneal injections, which increased urine volume. One hour after treatment with HCG, urine from treated males was expressed over freshly extruded eggs. McKinnell et al reported that the volume of urine released was great enough to cover 50 to 150 eggs. They reported an overall fertilization rate of 73% using this technique, as compared with a control fertilization rate of 88% using sperm suspensions obtained by standard methods.

Reference McKinnell, R. G.; Picciano, D. J.; Krieg, R. E. Fertilization and development of frog eggs after repeated spermiation induced by human chorionic gonadotropin. Laboratory Science 26(6):932–934; 1976.

Appendix G

239

Patterns and Experiments In Developmental Biology

Key experiments in practical developmental biology

Key Experiments in Practical Developmental Biology

Key experiments in practical developmental biology

Advances in Developmental Biology

Developmental Biology

Developmental Biology

Current Topics Developmental Biology (Current Topics in Developmental Biology)

Current Topics Developmental Biology (Current Topics in Developmental Biology)

Developmental Vascular Biology, Volume 62 (Current Topics in Developmental Biology)

Developmental Vascular Biology (Current Topics in Developmental Biology)

Instant Notes in Developmental Biology

Electroporation and Sonoporation in Developmental Biology

Current Topics in Developmental Biology

Developmental Biology Protocols

Developmental Biology Protocols

Developmental Vascular Biology

Developmental Biology Protocols

developmental biology protocols

Developmental Biology Protocols

Essential Developmental Biology

Developmental Biology Protocols

Developmental Biology (9th Edition)

Cell and Molecular Biology: Concepts and Experiments

Current Topics in Developmental Biology, Volume 64 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 61 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 58 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 52 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 51 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 64 (Current Topics in Developmental Biology)

Patterns and Experiments In Developmental Biology

Key experiments in practical developmental biology

Key Experiments in Practical Developmental Biology

Key experiments in practical developmental biology

Advances in Developmental Biology

Developmental Biology

Developmental Biology

Current Topics Developmental Biology (Current Topics in Developmental Biology)

Current Topics Developmental Biology (Current Topics in Developmental Biology)

Developmental Vascular Biology, Volume 62 (Current Topics in Developmental Biology)

Developmental Vascular Biology (Current Topics in Developmental Biology)

Instant Notes in Developmental Biology

Electroporation and Sonoporation in Developmental Biology

Current Topics in Developmental Biology

Developmental Biology Protocols

Developmental Biology Protocols

Developmental Vascular Biology

Developmental Biology Protocols

developmental biology protocols

Developmental Biology Protocols

Essential Developmental Biology

Developmental Biology Protocols

Developmental Biology (9th Edition)

Cell and Molecular Biology: Concepts and Experiments

Current Topics in Developmental Biology, Volume 64 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 61 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 58 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 52 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 51 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology, Volume 64 (Current Topics in Developmental Biology)

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