Patterns and Experiments In Developmental Biology 3rd ed - L. Johnson (2001) WW.pdf - książki - Bio Med 20 - bibiariel

ISBN: 0-07-237965-0

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

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 to-

gether 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 op-

portunities 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 de-

velopmental 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 frus-

trated with their inability to observe development beyond the earliest stages, and I think that these di-

rections 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 pro-

vides 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

Johnson:Johnson &

Volpe’s Patterns &

Experiments in

Developmental Biology, 3/e

Front Matter

Preface

Companies, 2004

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 pro-

vide 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 particu-

larly 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 sug-

gestions 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 insight-

ful 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 ar-

eas 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 recog-

nition of his original contributions.

Leland G. Johnson

viii Preface

viii

Johnson:Johnson &

Volpe’s Patterns &

Experiments in

Developmental Biology, 3/e

I. Echinoderm Development

1. Fertilization and Early

Development of Sea

Urchins and Sand Dollars

Companies, 2004

LABORATORY

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 proper-

ties 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 as-

sembly 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 ob-

servation 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 individ-

ual 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 ani-

mal 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.

Johnson:Johnson &

Volpe’s Patterns &

Experiments in

Developmental Biology, 3/e

I. Echinoderm Development

1. Fertilization and Early

Development of Sea

Urchins and Sand Dollars

Companies, 2004

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 fine-

gauge needle inserted at a very shallow angle. To enhance effectiveness of the KCl injection, it is advisable to di-

vide 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 in-

jecting 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 gono-

pores 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 refrig-

erated 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 us-

ing 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 arrange-

ment 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 sea-

water 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 sus-

pension 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 ex-

cessively 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.

2 Laboratory 1

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