TESTED STUDIES FOR LABORATORY TEACHING
Proceedings of the Ninth Workshop/Conference of the Association for Biology Laboratory Education (ABLE)
Edited by R. W. Peifer
Associate Education Specialist and Laboratory Coordinator General Biology Program College of Biological Sciences University of Minnesota Copyright © 1988 by the Association for Biology Laboratory Education (ABLE)
All rights resewed. No part of this publication may be reproduced, stored in a retrieval system. or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording, or otherwise, without the prior written permission of the copyright owner.
Printed in the United States of America. 1916-1987
This volume is dedicated to the memory of Stanley Dagley, Regents Professor of Biochemistry, University of Minnesota, for his dedication and many contributions to science laboratory education.
Contents
Foreword ix Preface xi Address xiii Chapter 1 Use of the Rabbit Intestine in Smooth Muscle Pharmacology Experiments: A New Approach 1 Richard L. Walker, Charles C. Scott Key Words: physiology, pharmacology, smooth muscle, neurotransmitters, autonomic control. A technique for demonstrating the pharmacological properties of smooth muscle using the rabbit intestine will be presented. By including a segment of the sympathetic nerve along with a section of the gut, the autonomic control of intestinal smooth muscle activity can be demonstrated through stimulation of the nerve and application of various neurotransmitters. Removal of sections of the intestine along with the sympathetic nerve will be demonstrated as well as a method for recording muscle contraction. This exercise is suitable for general physiology or zoology teaching laboratories.
2 Tracheid Length Measurement in Selected Conifer Species 27 J. Tidswell, W. J. Mullin, K. G. Tidswell Key Words: tracheids, staining, mounting, measuring, light microscopy. Tracheids in conifer wood can be easily separated and their lengths measured with a light microscope. This workshop will illustrate the techniques for macerating the wood and staining, mounting and measuring the tracheids. Mean tracheid length is different for each conifer species, therefore this laboratory can be expanded into a larger scale comparative measurement exercise.
3 Community Ordination Utilizing Winter Stoneflies 37 Vinnedge M. Lawrence Key Words: community ecology, taxonomy, winter stonefly, Taeniopterygidae, Capniidae, computerized key, ordination. This exercise in community ecology can be carried out in mid-winter and introduces participants to useful taxonomic and statistical procedures. Species determinations of winter stoneflies are facilitated by a computerized key featuring color illustrations. Taxonomic data are used to construct a two-dimensional ordination of the communities from which specimens were collected. Correlations are then sought between differences exhibited by communities and gradients of environmental conditions.
4 The Use of Echosounding Equipment in Limnology and Ecology Classes 51 Peter Vaughn Key Words: limnology, echo sounding, ecology, lake morphometry, zooplankton, macrophyte, oxygen, temperature, chlorophyll. This workshop will demonstrate the use of commercially available echo sounding equipment as a tool in ecological studies which are appropriate for class field projects. The type of information that can be obtained is: lake basin morphometry, depth of the mixed-layer, spatial distribution of macrophytes and zooplankton and the location and size of fish. During the workshop, we will do several transects of Lake Minnetonka with the echo sounder, and measure depth profiles for oxygen, temperature and chlorophyll. We will then return to the Gray Institute to discuss the interpretation of recordings.
v 5 A Handbook for Collecting Releve Data in Minnesota 63 John C. Almendinger Key Words: vegetation sampling, releve method, plot method, physiognomic, species composition, multivariate analysis, field methods. The releve method is a semi- quantitative plot method that records both physiognomic (structural) and species composition data. The workshop will demonstrate releve field methods and discuss the multivariate analysis of releve data The application of releve data to problems of habitat evaluation, forest inventory, and research projects in Minnesota will be presented by experts from the Minnesota Natural Heritage Program, the Minnesota County Biological Survey, and the University of Minnesota. All participants will receive a releve handbook that covers the history of the method, field instructions, data-entry forms, and an overview of a menu-driven computer system (SAS Version 5) used to manage a releve database.
6 The Electroretinogram of the Horseshoe Crab, Limulus polyphemus: A Laboratory Exercise in Sensory Physiology 101 Robert A. Linsenmeier, Charles M. Yancey, Wesley W. Ebert Key Words: horseshoe crab, Limulus polyphemus, sensory physiology, electroretinogram. The eye of the horseshoe crab, Limulus polyphemus represents an easily-excised and durable preparation for investigating various parameters of a typical sensory system. One can study the time course of dark adaptation as well as the dependence of response amplitude and latency on stimulus intensity in both the dark- adapted and light-adapted eye. Requirements for specialized, technical equipment are minimal. Suitable for undergraduates in advanced general biology, physiology, and special projects.
7 Methods to Process and Identify Symbiotic Fungi in the Roots of Vascular Plants 131 Iris Charvat Key Words: fungi, symbiotic fungi, vesicular arbuscular mycorrhizae (VAM), processing, staining, clearing, identification. Vesicular arbuscular mycorrhizae (VAM) are present in the roots of almost all vascular plants. VAM play a crucial role in the mineral nutrition of these plants by transferring phosphorus and other minerals from the soil to the plant. Techniques for obtaining VAM samples from natural sources and from inoculated samples will be demonstrated. Methods of processing, staining and clearing root samples, and identification of the fungal structures will be demonstrated 8 Teaching Botany Through Inquiry 131 Gordon E. Uno Key Words: inquiry method, laboratories, plant physiology, photosynthesis, plant anatomy, scientific method, botany. The introductory botany course at the University of Oklahoma is taught using the "inquiry method," integrating the laboratory experience with the lecture and discussion. Workshop attendees will participate in the inquiry method, focusing on first-day activities we use in our class. Inquiry-oriented laboratories on plant physiology, including photosynthesis, and plant anatomy also will be demonstrated.
9 A Laboratory Introduction to DNA Restriction Analysis* 171 David A. Micklos, Greg A. Freyer *Workshop presented by William R. Gette Key Words: restriction enzymes, DNA, agarose gel electrophoresis,lambda virus, molecular biology. This workshop will serve as an introduction to laboratory exercises in molecular biology. DNA from lambda virus will be digested with various restriction enzymes, and the resulting fragments separated using agarose gel electrophoresis. The
vi separation patterns will be visualized, photographed and used to illustrate the relationship between DNA fragment size and electrophoretic mobility. 10 Resource Partitioning in Potentially Competing Insect Taxa 187 John A. Haarstad Key Words: interspecific competition, community structure, resource partitioning, insects, coexisting species, niche. Field exercises in quantifying the niche relationships (e.g. extent of spatial, temporal, and food resource overlap) in coexisting and potentially competing invertebrate species such as ants and carrion beetles will be conducted.
11 Supercooling and Freezing in Winter Dormant Animals 193 William D. Schmid Key Words: winter dormancy, cryoprotectant biochemicals, antifreeze, supercooling, winter ecology, goldenrod gallfly, Eurosta solidagensis, physiology, nucleator chemicals. Winter dormant organisms, both plants and animals, have two general categories of adaptation for survival of exposure to cold climate stress. They can resist the formation of internal ice by supercooling through the production of antifreeze compounds; or, they can tolerate internal ice by addition of cryoprotectant biochemicals to their body fluids. In the latter case, nucleator chemicals may be produced to promote the formation of ice in extracellular fluids. We will use techniques to measure supercooling points of a winter dormant animal, the goldenrod gallfly, and to evaluate its seasonal production of cryoprotectant chemicals. 12 The Use of Yeast for Teaching Microbiological Techniques and Principles 2 05 Robert J. Doyle Key Words: yeast, cell culture, techniques, nutrition, growth, mutation, biological engineering, media. Cell culture methods, so critical in modem biology, may be taught cheaply, safely and simply by using standard brewers yeast or simple mutants derived from it. Media and methods designed to be student proof, and a variety of simple experiments on life cycle, nutrition, growth, and mutation will be described Applications in biological engineering will also be discussed. [Note: Manuscript not received in time to be included.] 13 Ideas to stimulate the Non-Majors Biology Student 207 Roberta B. Williams, Haven Sweet, Barbara Newman 1. Understanding Human Energy Requirements - A laboratory Exercise, Roberta B. Williams 13-209 2. Ideas to Stimulate the Non-Major Biology Student, Haven Sweet 13-223 3. Biology From the Human Perspective, Barbara Newman 13-261 Key Words: undergraduate, non-major, biology laboratory education. techniques, microscopy, nutrition, genetics, evolution, computer programs. This workshop will give the participants a chance to experience several different techniques that have been used by the presenters in biology classes for the non-biology major. Topics such as introduction to microscopy, nutrition, body composition. body systems, genetics and evolution will be presented. The workshop will include a number of stations where participants can do experiments, run computer programs and obtain resource materials. Scientific methodology will be stressed and hopefully the ideas presented can be easily incorporated into the participants existing course.
vii 14 Laboratory Safety Principles 305 Jerry Staiger, Keith Carlson, Jim Lauer Ray Arntson 1. Radioactive Materials, Jerry Staiger 14-307 2. Toxic, Reactive, Carcinogenic, and Teratogenic Chemicals, Keith Carlson 14-337 3. Infectious Agents, Jim Lauer 14-343 4. Fire and Physical Hazards, Ray Arntson 14-355 Key Words: laboratory safety, regulations, hazardous materials, radioactive, toxic, reactive, carcinogenic, teratogenic, infectious agents, fire, physical hazards. This workshop will cover major principles and regulations pertinent to working in laboratories with hazardous materials. It will be divided into 45-minute segments dealing with: A. Radioactive Materials B. Toxic, Reactive, Carcinogenic, and Teratogenic Chemicals C. Infectious Agents D. Fire Safety Concepts and Physical Hazards. 15 Plant Growth Responses to a Nitrogen Gradient 383 Mary Lynn Cowan Key Words: growth response, plant physiology, field experiments, greenhouse experiments, techniques, nutrient gradients, soil sampling. Techniques used in setting up field experiments to study the growth responses of herbaceous plants to a resource gradient will be discussed and demonstrated in the field. Methods of setting up light and nutrient gradients, soil sampling, seed collection and preparation, data collection and analysis will be included Participants will observe on-going studies in the field at the Cedar Creek Natural History Area, and view slides of smaller scale greenhouse experiments.
viii Foreword
Don Igelsrud
The foreword to these proceedings was written in January of 1988. I wrote the forward to the proceedings of the 1982 University of Washington ABLE meetings in January 1984. Those proceedings were the last volume published by Kendall/Hunt and the last volume to appear until now. Since that time the new editor, Dick Eckland of Cornell, has struggled with the problem of producing a scholarly manuscript for a much more limited audience than we hoped to reach by publishing a hardback reference work for libraries. Even though only 1200 copies of the proceedings were published by Kendall/Hunt, we discovered that their acceptance by college and university libraries in the austere economic environment of the 1980's was much more limited than we had anticipated. Consequently, the Executive Board decided to reduce the cost by publishing the proceeding ourselves. The advent of desktop publishing made this seem like a reasonable endeavor. The goal of developing scholarly manuscripts from the materials distributed at the workshop, however, has delayed the publication of subsequent volumes of the proceedings. The proceedings from 1983 to 1986 should appear forthwith. Because Rick Peifer put together a well developed package of materials for the 1987 workshop participants, the ABLE board decided that it would be more useful and expedient to publish the proceedings in a form closer to the way materials were prepared for the meeting. Many of the manuscripts for the meeting were submitted on floppy discs and it soon became clear that the most efficient way to publish materials would be on floppy disc. Consequently, future volumes of the proceedings will be available in both hard copy and electronic forms. Rick Peifer also succeeded in accommodating more participants at the workshop by adding several field exercises. Because the most recent ABLE meetings have been filled to capacity, host institutions will try to present 15 rather than 12 major workshops. This volume of the proceedings is dedicated to the late Stanley Dagley, the first editor of Biochemical Education and Regents' Professor Emeritus of biochemistry at the University of Minnesota. Professor Dagley's eloquent banquet address is included in these proceedings. Readers not familiar with Biochemical Education will be pleased to discover one of the most useful publications concerned with teaching biology. The annual workshop/conference continues to be ABLE's primary activity. The more informal process of distributing the proceedings of these meetings is indicative of the organization's style and is representative of ABLE's main goal: to improve biology laboratory education by developing active communication among teachers. If you would like to become part of the ABLE network write to:
Anna Wilson Biochemistry Department Purdue University West Lafayette, IN 47907
ix 1987 ABLE CONFERENCE
UNIVERSI'I'Y OF MINNESOTA
Row 1: Leigh Callan, Andrew Greller, Gil Gleason, SherylShanholtzer, Marsha Fanning, Anna Wilson, Jim Waddell, Don Igelsrud, Angela Gloss, Joe Nickolas, Les Turoczi. Row 2: Beth Nicholas, Roberta Williams, Jim Perry, Barbara Dcnson, Nancy Goodyear, Sandy Solomon, Lynn Hodgson, Janc Nall, Kathy Tidswcll, Leanne Jablonski, Elizabeth Wurdak, Cynthia Campbcll, Bill Glider, Lorraine Mineo, Sheri Dalton, Karen Morris, Donna Daugherty. Row 3: John Dropp, Neil Grant, Jan Phelps, Lavinia Kumar, Nancy Elliott, Roberta Ellington, Harriet Gray, Barbara Ncwman, Mary Ball, Ruthanne B. Pitkin, Minnie B. Ragland, John Tidswcll. Ron Tempest, Wanda I,. Kcndrick, Eleanor R. Brown, Janc Beiswenger, Barbara Stewart, Paul Snow, Leland Johnson. Row 4: Jon Glase, Pat Harrison, Jean Dickey, Cindy Rcittingcr, Bill Leonard, Vin Lawrcncc, Robert Foulkes, Gcnc Menton, Jerry Opbrock, Rod Sutherland, Lynda Swander, Judy Morgan, Leona Truchan, Mary Jane Turnell, Lucy Dyer. Row 5: James Bittner, Forrest Bent, Bette Nicotri. Bob Kull, Dan Hinrichs, Robert Chase, Rill Werner, Mike Butts, Ned Lyke, Dave Mayette, Elizabeth Godrick, John Wcstbrook, Paul Holeski, Henry Butcher, Bruce Odel, Dick Ecklund. Preface
In an effort to bring the Proceedings volumes more quickly to press, and to cut publishing costs, the Executive Board of ABLE decided at its 1987 annual meeting to abandon the hardcover and typeset format of previous volumes. Those of you familiar with the earlier Proceedings volumes of ABLE will immediately recognize a radical departure in the style and quality of production. With word processing and desktop publishing software becoming commonplace in academic settings, the Board felt that future workshop presenters should submit their manuscripts to the hosting institution on computer floppy disks in camera-ready, ASCII format. This should enable the editors of future proceedings to manipulate the text more easily into a consistent type style throughout the volume, and to produce a volume that has a polished professional look. It should also make other editorial changes much easier, and should shorten the time it takes to publish the proceedings. This volume therefore represents a transition toward this goal. An unavoidable distraction in this volume is the plethora of different type styles and formats in the chapters, but the Executive Board felt that for this volume only, the manuscripts should be printed in camera-ready copy as submitted by the authors. To do otherwise would have necessitated retyping or typesetting the entire volume. In either case the cost would have been much higher, and the time to publication much longer. As in previous volumes, the workshops are organized by chapters. It contains fourteen of the fifteen workshops presented. Because of unavoidable circumstances, the workshop titled The Use of Yeast for Teaching Microbiological Techniques and Principles by Robert J. Doyle was not included in this volume. Five of the chapters in this volume include ecological workshops, four of which were presented in the field. Physiology and Botany each are represented by three chapters, while molecular biology and microbiology are the subject of single chapters. There is a chapter concerning biology for non-majors, which contains three sections by individual authors. Similarly, the chapter on laboratory safety contains four sections, covering the topics of fire and physical hazards, radioactive materials, hazardous chemicals, and infectious agents, each by different authors. The Table of Contents contains, for each chapter, a list of key words and an abstract I wish to thank Geri Grosinger and Bob Jacobson for assisting in the assembly of this volume, and Bruce Fall for his comments and suggestions. I would also like to thank the authors for being extremely cooperative and patient. Because of the camera-ready nature of the manuscripts, the amount of editing in this volume was minimal compared to the task of previous editors Jon Glase. C. Leon Harris, Dick Ecklund and Mel Zimmerman Hopefully future editors will maintain their high standards.
Minneapolis, Minnesota R. W. Peifer
xi
Banquet Address
The following is a transcript of the address that Professor Stanley Dagley gave to the members of ABLE at the closing banquet of the annual Workshop/Conference held at the University of Minnesota, June, 1987. A large number of attendees asked if his address could be included in the proceedings volume. Dr. Dagley graciously agreed, and, true to form, the manuscript was on my desk the Monday following his talk even though he was leaving for England that same day. I have the unpleasant task of informing you that Dr. Dagley passed away suddenly on October 31, 1987. He was a passionate and devoted supporter of laboratory science education. He was the first editor of the journal of Biochemical Education, and active in teaching his entire adult life. Those of us who had the pleasure of interacting with Dr. Dagley over the years have suffered a great personal loss, but not as great as the loss to science education.
Scientists, Artists and Con-artists
Stanley Dagley As Sam Kirkwood has mentioned I'm a scientist by profession interested particularly in the chemistry of living forms, and their evolution and I'm also interested in cathedrals and other buildings, and the way that particular art form evolved in the middle ages. Science and art are both creative activities. Both are the fruit of disciplined imagination. Sometimes the two activities are interlaced in an obvious and direct manner. For example, the glorious tower and spire of Salisbury Cathedral is a permanent tribute to the creative, soaring imagination of mankind. On the other hand my ancestors of the 14 C. did not build a spire reaching to a height of more than 400 ft. without a deep understanding of the properties of the materials they used. You can't raise 6,400 tons of stone to that height without some knowledge of mechanics: you're building a slim spire, not a pyramid. You can't arrange for the whole lot to stand up for 6 centuries unless you have discovered how to distribute its crushing weight, so that the choir below can sing cheerfully and without apprehension. Sometimes when we talk of issues, we say that there are two sides to the coin. A little later I shall say more about how science and art, together, form the design of the upper side of the coin of our culture. I shall also refer to the con-artists who shape the lower side. And since I'm a poor swimmer I'll say right now where the members of ABLE belong. They are absolutely indispensable for the complete education of a life scientist. They are on the upper side of the coin. There is the responsibility for presenting the working materials that can stimulate in students - right at the outset - an abiding interest in biological enquiry. From the experiments that they organize, the young biologists begin to learn how to ask sensible questions of nature, and how to decide whether or not the answers they get are convincing. When this is done successfully it's more basic to biological education than any number of lectures. And it entails an insight which distinction in scientific research does not automatically confer - namely the insight of how laboratory work can be adapted effectively to the appropriate instructional level. Because, after all, a university exists to find and communicate the truth; for research and teaching. It's a poor analogy to describe a university as a ladder that bright young researchers can ascend, until they decide to flit to a taller ladder. Salisbury Cathedral provides a better comparison. Its spire seems to touch the clouds. In these clouds, by analogy - clad I believe in white raiment. are the regents of the U. At the top of the spire of President Keller. As with all portions of great responsibility, this tip is no place to sit down and take it easy.
xiii Moreover, as clouds of retrenchment gather in the State Legislature, he can never be sure that he won't be struck by lightning. Then, as we descent we pass the administrative structure of provost, deans and dept. heads, and at ground level we reach the operation that justifies the whole construction. This is where the students are, and you are part of the company that ministers to their needs. This is the only justification for the 6,400 tons above your heads. I want to say a few words about why these efforts of yours are part of the complete education of the whole man or woman. First, you can't take your full place in modem society without some knowledge of biology. Its benefits to agriculture and medicine in the past are obvious. The use of microorganisms, for commercial production of very expensive biochemicals of value in medicine, encourages the hope that these cures may become generally available for the sick and the poor by the end of this century. Science is sometimes assailed for insults to the environment - an indirect consequence of technological advances; or for contempt of humanity in the form of weapons to wipe out innocent and guilty at one fell swoop. These themes would take us into vast realms of argument. Suffice it to say that we must distinguish between science and technology: between the extension of knowledge and understanding, and the use to which we put it. And in the sphere of environmental pollution, which impinges upon my own area of research, our increasing understanding of biodegradatoin and microbial capabilities encourages the hope that we shall be able to enlist microbes in clearing up the mess we have made of this planet. But having said this, I believe we greatly undervalue the contributions of science to the quality of human life if we see an increase in material comfort as its only justification. Let me put it to you that the universe will always be mysterious, and it is good for our souls that we should admit this. Creation is infinitely vast and complex, and man is fragile. However, the universe makes more sense to us than it once did, thanks to science, and this understanding has liberated us from many of the terrors that beset our ancestors. Let me give you one example of those terrors. The world has been stricken by bubonic plague at intervals throughout history. The activities of the microbe that was responsible for the disease shaped the development of human societies. Its input was as significant as that of the human beings themselves. Thus, epidemics that killed one quarter of the population of Europe in the Middle Ages gave rise to chronic shortages of labor, these in turn by upgrading the value of men's labor, were the death knell of serfdom; and they were, as a consequence, one factor in destroying the feudal system of the Middle Ages. This was the impact of the microbe upon the structure of society. Now - imagine yourself living at this time, before science had bestowed some understanding of the activities of Pasteurella pestis and the way it was spread around. Thought processes were paralyzed by the constant terror of being struck down, apparently at random, by unseen agencies; those who attributed sudden death to divine retribution could not even identify the specific transgressions for which they thought they were being punished. This is just one example of how our everyday lives have been enriched by the scientific conviction that the universe is not beyond the reach of reason: it is ordered; it is therefore comprehensible, within limits; and it's joy, and a privilege, to extend these limits of comprehension by scientific research. For many civilized men and women, life without poetry would lose color; life without the visual arts would lose immediacy; and let us add, that for all of us life without science would lose its structure and coherence. I have to say that the program of your meeting encourages me to think that these broader issues are thoroughly appreciated by all of you. I see a contribution on: ideas to stimulate the non-major biology student, another on the inquiry method, a paper in past published proceedings on the use of fossils to study evolution. And in this connection it was a strange experience for me to leave England where 120 years ago at a famous meeting in my old U., Oxford, some biologists has assailed fundamentalist religion. And then to find the battle still raging in the United states where some religious persuations were telling the young, modem biologist that he was in the dogma dog-house for believing in evolution. However, in Christian charity, the dog-house was air-conditioned Whether it is tax-deductible has not been decided.
xiv You know, well designed practical work, on complex biological systems offers the best hope of getting across the living spirit of science to undergraduate students. The attitude that knowledge is growing: that it is, therefore, incomplete: that our ideas might need revision. And above all, that this admission is not so painful as to spoil the pleasure of getting the story more nearly correct. How would you persuade your peers in Washington to fund your grant application if you - and they - already knew all the answers? Willingness to modify opinions, to receive new ideas, does not indicate spiritual weakness. On the contrary, arrogance combined with ignorance is a menace in public affairs, and a source of excruciating boredom in private life. Barbara Tuchman has given examples of what happens in the public sphere when open-mindedness is banished "Wooden-headedness", she wrote, blindness to any other point of view, the inability to change one's mind - these are the failures that have been at the root of mankind's poor performances in government, and have been the downfall of politicians and soldiers alike. She gives, as examples, the long marches of Napoleon and Hitler into Russia, with no turning back whatever disaster might be encountered; the inability of the German war lords to change their minds concerning unrestricted submarine warfare in 1917, despite the pleadings of the German chancellor who saw clearly that this would bring in the United States, and that Germany would then lose the war. And on a lighter note, the slogan - you can scarcely call it an idea - the slogan of George III and his government that the Americans were simply rebellious colonials, and that was all there was to it: despite the presentation of their case by an intellect as formidable as that of William Pitt, Earl of Chatham. Another English King provides an excellent example of the truth that much learning does not confer wisdom upon the arrogant. James I was one of the few monarchs who might have gained straight A's in written coursework. But his wisdom was something else. He was full of instructions, directed to better people than himself, as to what they ought to believe and how theyought to behave. He himself sincerely believed he had that authority. by Divine Right, as a King. He was wrong. I've observed that even in a Republic, some people hold similar views about themselves. These are examples from the other side of the coin: people who despise, as a sign of weakness, those who seek to test theories by observations or critical experiments. Less serious in their impact are those who seek to make money by pseudo science - the con-artists. I say less serious because they sometimes brighten my day. Careful observation - note the precision - has shown that my teeth will be 57% whiter if I use this toothbrush or that toothbrush. And what a bustling life we lead. That young business lady in the bow tie, full of fibre - at least, I think, that's what she's full of - she sure is busy, literally running around N.Y.C. She chairs a Fortune 500 corporation executive meeting in the morning, and the National Council of Churches in the afternoon, and arrives home with the TV dinners recommended by Robert Morley in the evening; surely she would collapse in a heap were it not for the fact that her stockings massage her legs all day. "What is life if full of care"? "We have no time to stand and stare" wrote W. H. Davies. What indeed. There's that poor singer who hasn't even got time for the pain. He wouldn't be singing but for the advice of 1000 doctors stranded on a desert island. But can you rely on them? Would your advice be reliable if you were stranded on a desert island with 999 doctors? And then there's the affectionate mother whose daughter's calcium requirement are met by chewing indigestion tablets. Perhaps that's what was wrong with the singer: he was on a diet of indigestion tablets. margarine and fibre. But there was a time when calcium was out of fashion and dairy products were simply a source of cholesterol. It was in those days that the Detroit Free Press issued its famous headline:
Milk Drinkers Turn to Powder. As I look around the table at your glasses I see that we have averted this catastrophe.
xv
Chapter 1
Use of the Rabbit Intestine in Smooth Muscle Pharmacology Experiments: A New Approach
Richard L. Walker and Charles C. Scott
Department of Biological Sciences University of Calgary Calgary, Alberta, Canada T2N 1 N4
Richard L. Walker received his B.S. degree in Biology from Alma College, Alma, Michigan in 1969 and his Ph.D. in Physiology from Michigan State University in 1975. He is the coordinator of the animal physiology teaching laboratories at the University of Calgary, a position which he has held since the Fall of 1975. He has lectured in General Biology, Introductory Zoology, Human Physiology and Mammalian and Compartive Physiology. His research interests include the respiratory and acid-based physiology of fish. Charles C. Scott received a B.Sc. degree in Zoology from the University of Calgary in 1968. He has been a laboratory instructor in Introductory Zoology, and Human and Mammalian Physiology. Currently, he is the laboratory technician for the animal physiology teaching laboratories.
1
INTRODUCTION
The isolated intestinal smooth muscle preparation is one of the classical preparations in physiology and pharmacology for bioassays, or the study of drug action and autonomic control of motility. This preparation is included in many "in-house" laboratory manuals of various colleges and universities around North America, and in some commercially prepared manuals deal ing with physiology and pharmacology (e.g. Nicpon-Marieb, 1981).
Basically, the method presented in this report is a modification of the original Finkleman preparation (Finkleman, 1930) for the study of the autonomic control of intestinal motility. What is different about our approach is the method of mounting the preparation and the method of stimulation of the sympathetic nerve. The classic way of mounting the piece of intestine is to suspend it vertically in a muscle bath between an aeration tube and a recording lever. The problems with this technique are (1) stress placed on the intestine when the chamber is emptied during the process of changing solutions, and (2) difficulty in stimulating the sympathetic nerve due to the fact that the preparation is totally submerged in a physiological saline solution. We have overcome these problems by mounting the segments of gut horizontally in a shallow muscle bath. As a result, less stress is placed on the intestine during changeover of solutions, and it is easier to manipulate and to stimulate the sympathetic nerve contained within the mesentery. Also students find it much easier to mount the preparation in the horizontal bath and are less likely to stretch the muscle preparation in the process.
With these modifications we have improved the student success rate from 50-60% to 90-100%. Large recordings of the muscle contraction, such as those shown in Appendix A, are easily obtainable using a kymograph and simple lever system. In fact, another advantage of this exercise is that it does not require expensive recording equipment.
3 INSTRUCTOR'S GUIDELINES
Equipment and supplies:
A list of the equipment, supplies and solutions needed to operate this laboratory exercise is given in Appendix B. Instructions and dimensions for building the muscle bath, swivel assembly and stimulating electrodes are given in Appendix C.
We recommend using a simple isotonic lever system for recording the muscle contraction because of the ease with which students are able to comprehend and manipulate the recording system to obtain excellent tracings. Recordings of comparable quality, however, are also obtainable with an isometric transducer (e. g. a Physiograph F-60 force transducer).
Preparation of isolated segments of the rabbit intestine:
The rabbit should be starved for 24 hours prior to use. Food in the gut will result in a messy dissection and will necessitate flushing to remove the gut contents, a practice which could damage the intestine.
Before sacrificing the rabbit:
1. Prepare Tyrode's Ringer solution (see Appendix B) and place about 250 mL of this solution in a flask on ice.
2. Cut sections of thread (or 4-0 silk) into 5 cm lengths. One piece will be needed for each segment of gut.
3. You will need a pair of fine (iris) scissors, coarse scissors, and curved forceps. Hemostats may also be helpful in retracting the abdominal muscles.
4. A large Petri plate or other container for holding the segments of gut will also be necessary.
5. Wash all instruments and glassware before use. Also make sure your hands are clean before handling the intesting.
Sacrificing the rabbit and removing the intestinal segments:
1. Since anaesthetics can affect motility of the gut, the animal should be sacrificed by cervical dislocation, without use of anaesthetic.
2. Shave the abdomen and carefully vacuum the surface to remove any loose or cut fur.
3. Make a mid-line incision through the skin and abdominal muscle using a pair of coarse scissors.
4. Locate the ileum, a section located near the end of the small intestine. Note the intestinal arteries and veins coursing through the mesentery to and from the small intestine. If a segment of the ileum is picked up with both hands, the intestinal vessels are easily seen spaced about 2 to 3 cm apart (Fig. la). If you look very carefully, a fine white line following the intestinal artery and vein into each segment of the gut is visible. The white line is a branch of the sympathetic nerve arising from the prevertebral ganglion. 4 5 5. Using a pair of fine curved forceps, carefully break through the mesentery on either side of the intestinal vessel-sympathetic nerve complex about 4 cm away from the gut (Fig. lb). Insert a length of thread or silk suture through the hole and tie off the vessels and nerve.
6. Follow the vessels down to the gut. The vessels divide into many branches just above the surface of the intestine. Select a 3-cm segment of the ileum that is supplied by these vessels. Sever the gut and remove this segment along with the mesentery containing the vessels and nerve. Place in a Petri dish containing cold Tyrode's Ringer solution.
N.B. If the rabbit has been starved for 24 hours, the gut segment should be free of chyme. However, if chyme is present, it can be removed by carefully flushing the segment with Tyrode's. A syringe filled with Tyrode's may be placed inside one end of the segment, and using very little pressure, the contents can be carefully forced out into a waste container. Caution must be exercised because high pressure will severely damage the tissue.
7. Repeat this procedure until enough segments have been removed to supply your laboratory needs for the day. If the extra pieces of ileum are kept in ice cold Tyrode's, they will stay viable for several hours. Do not store in warm Tyrode's.
Suggestions:
1. The ileal segments will shorten when placed into the cold Tyrode's solution and motility will cease. However, when mounted in the warm (37°C) Tyrode's solution inside the muscle bath, motility will return within 5-10 min.
2. As the muscle warms in the bath, it will relax and segmental and peristaltic waves will be evident.
3. It is essential that the muscle baths be at 37°C at the start of class. This means starting the circulation of warm water through the baths at least 1/2 hour before class. Also, it is essential that the extra Tyrode's solution for the muscle baths be at 37°C.
4. It is not necessary to warm the drug solutions used in the experiment since they will be added directly to the muscle bath in very small amounts (0.1 to 1.0 mL of drug/100 mL of Tyrode's).
5. One of the advantages in adding drug directly to the bath in concentrated form is that rapid changes in muscle tonus can be clearly recorded. We recommend that the Tyrode's solution be replaced between drug additions so as not to disturb the motility directly before the application of the drug. It may take 1-2 minutes for the rhythm to stabilize after replacing the solution in the muscle bath.
6 Student's Guideline
AUTONOMIC CONTROL OF THE VISCERAL SMOOTH MUSCLE OF THE RABBIT INTESTINE
OBJECT:
To show some of the autonomic properties of smooth muscle as it occurs in the rabbit's small intestine.
EXPLANATION :
All smooth muscle cells have one thing in common: the actin and myosin filaments of the sarcomeres are not organized into bands such as they are in skeletal muscle. Visceral smooth muscle, such as the type found in the digestive tract, is a syncytium of muscle cells where intercellular communication can occur via gap junctions. Electrical current is easily conducted from one cell to the next via the gap junctions; therefore, depolarization of one muscle cell can quickly lead to depolarization of neighboring cells.
Visceral smooth muscle contraction occurs spontaneously, meaning that muscle cell action potentials are generated without input from either the motor or autonomic nervous system. The muscle cells undergo rhythmic oscillations in membrane potential which, occasionally, reach the threshold of an action potential, and, thus, generate a spike. The action potential spreads via gap junctions from muscle cell to muscle cell, initiating a wave of muscle contraction in its wake.
Visceral smooth muscle cells also exhibit muscle tonus, a state of long-term, steady contraction. The tonus is variable, depending on the number of muscle cells that participate.
The rhythmicity and tonus inherent in the intestinal smooth muscle may be enhanced or suppressed by two nerve plexuses found between the layers of muscle and mucosa (Auerbach's and Meissner's plexuses). Known as the enteric nervous system of the gut, the activity of the plexuses can be modified by the autonomic nervous system. When a piece of intestine is removed for studv.. the plexuses remain viable and can be stimulated or inhibited using parasympathomimetic or sympathomimetic drugs (mimicing the action of the parasympathetic and sympathetic nervous systems).
The autonomic control of the gut is very complicated. Figures 2 and 3 illustrate the parasympathetic and sympathetic input to the enteric neurons of Auerbach's and Meissner's plexuses.
7 Figure 2*
(N) NANC
Sympathetic stimuli subdue the inherent rhythmicity of the gut. Note that the sympathetic input is upon alpha-adrenergic receptors located on the axon terminals of the enteric neurons, or upon the axon terminals of the preganglionic parasympathetic fibers. Release of norepinephrine from the sympathetic postganglionic fibers stimulates the alpha receptors, which prevent release of acetylcholine from the enteric neurons or preganglionic parasympathetic fibers. This action is known as presynaptic inhibition.
There are alpha- and beta-adrenergic receptors on the smooth muscle cells which will respond to epinephrine and norepinephrine in the circulation, as shown in Figure 3. Both alpha and beta receptors inhibit muscle tone and rhythmici ty.
Figure 3*.
ADRENAL MEDULLA ---
INTRINSIC GANGLION
The parasympathetic control of the gut is via cholinergic preganglionic fibers which synapse with enteric neurons. When activated, the parasympathetic fibers release acetylcholine at the enteric neurons, thus stimulating the enteric neurons to release acetylcholine at their neuro-muscular junctions. Enhanced smooth muscle tone and rhythmicity result.
*Reprinted from "Innervation of the Gastrointestinal Tract" in A Guide to Gastrointestinal Motility, Christensen and Wingate (eds.), with permission from John Wright Medical Publishers.
8 The receptors of the neuromuscular junction are muscarinic type cholinergic receptors. They can be blocked by the addition of atropine to the muscle preparation.
In the following laboratory exercise you will study the action of sympathe- tic nerve stimulation, and the effects of various pharmacological agents on the rhythmicity and tonus of the rabbit gut. The agents will include sympatho- or parasympathomimetic drugs, along with adrenergic and choliner- gic blocking agents.
PROCEDURES:
The rhythmical activity of the piece of rabbit gut will be recorded using a lever system and kymograph. To maintain its activity, the gut will be sus- pended in a bath of Tyrode's solution at 37ºC with adequate oxygen supply. (A mixture of 95% 02 - 5% CO2 will be used).
Wash your hands thoroughly with soap and water before handling the gut. Body salts, oil and dirt will kill the gut tissue.
FIGURE 4. Arrangement of Intestine in Smooth Muscle Bath
9
Preparation:
The muscle chamber is set up as in Figure 4. Warm water is circulated through the outer bath to keep the muscle chamber at 37°C. One end of a section of intestine is attached to the anchor pin in the chamber using an S-hook. The other end is connected via an S-hook to a swivel which transposes horizontal contractions and relaxations of the gut into vertical movement of a recording lever. The movement of the recording lever are traced upon a kymograph drum, or chart recorder.
1. Attach two S-hooks to the gut, on opposite sides, at the opposing ends. During this operation, keep the gut in a Petri plate containing Tyrode's solution. Do not allow the tissue to dry.
2. Attach one hook to the anchor pin in the muscle chamber. Fasten the other hook to the swivel.
3. Fill the chamber with 100 mL of warm Tyrode's solution and bubble 95% 02-5% CO2 gas into the chamber fluid. There should be a fine stream of bubbles, rather than large bubbles which will disrupt your recordings.
4. Attach a thread from the muscle lever to the recording lever using plasticine. Remove slack from the thread by adding plasticene to the recording lever to act as a counter-balance. Keep the recording lever horizontal.
Experiment:
During the following experiment, check that your tracings are labelled clearly. Be sure that all drug concentrations, washes and solution changes are labelled so as not to confuse a drug response with an artifact. The "normal" muscle rhythmicity and tonus (i.e. prior to drug application) will vary throughout the experiment. Compare the drug effect to the "normal" muscle contraction in Tyrode's just preceding the addition of the drug.
10 A. Acetyl chol ine and Norepinephrine:
These two compounds act as postgangl ionic neurotransmitters of the parasympathetic and sympathetic nervous systems, respectively. Apply the drugs in the order listed below. After you have noted the effect of the drug, drein the muscle bath, and refill with 100 mL of fresh, warm Tyrode's solution. After the muscle contractions have returned to the normal rhythmicity and tonus, you may add the next drug. Add drops to the bath away from the gut; mixing occurs quickly by diffusion. Check bath temperature frequently.
1. 10-7 M acetylcholine: add 0.1 mL of 10-4 M acetylcholine to the bath.
2. 10-6 M acetylcholine: add 1.0 mL of 10-4 M acetylcholine to the bath.
3. 10-7 M norepinephrine: add 0.1 mL of 10-4 M norepinephrine to the bath.
4. 10-6 M norepinephrine: add 1.0 mL of 10-4 M norepinephrine to the bath.
B. Alpha- and Beta-Adrenergic Agonists:
1. Drain and refill the chamber with fresh, warm Tyrode's. Apply 1.0
mL of 10-4 M phenylephrine and note the response. Phenylephrine is an alpha-adrenergic agonist (stimulant).
2. Drain the chamber and wash the gut with Tyrode's. Refill the - 4 chamber and add 1.0 mL of 10 M isoproterenol, a beta-adrenergic agonist.
What can you conclude about the type of adrenergic receptors present in the smooth muscle of the gut?
C. Sympathetic Nerve Stimulation:
1. Mount the stimulating electrode on the side of the muscle chamber. Carefully slide the mesentery containing the sympathetic nerve into the electrode sleeve via the thread attached to the mesentery.
2. Stimulate the nerve with 30V (0.5 msec duration, 15 pps) for 10-20 seconds. If you do not see a response try 40V. Note the change in tonus and rhythmicity.
D. Alpha and Beta Adrenergic Blockade
1. Place 1.0 mL of 10-4 M phentolamine in the chamber. Wait one or two minutes, then try stimu lating the sympathetic nerve.
11
2. Add 1.0 mL of 10-4 M phenylephrine.
If the alpha receptors are blocked, phenylephrine should have no effect.
3. Drain and refill the chamber with warm, fresh Tyrode's solution. Add 1.0 mL of 10-4 M propranolo l. Wait two to three minutes, then try stimulating the sympathetic nerve.
4. Add 1.0 mL of 10-4 M isoproterenol to the chamber.
If the beta receptors are blocked, isoproterenol should have no effect.
E. Atropine - a cholinergic blocking agent
Drain and refill the bath with warm, fresh Tyrode's solution. Add 1.0
mL of 10-4 M atropine. Atropine blocks the action of acetylcholine and is, therefore, a parasympatholytic drug. Wait two to three minutes and
then add 1.0 mL of 10-4 M acetylcholine to the bath. Does the atropine successfully block the action of ,acetylcholine?
12 APPENDIX A
The chart recordings of the motility of an isolated segment of rabbit ileum, shown below and on the following pages, were collected by a pair of students using a kymograph and ink-recording lever system. The intestine was mounted horizontally in a muscle bath containing Tyrode's solution at 37°C. The preparation was aerated with a mixture of 5% C02-95% 02. Small volumes of concentrated drug solutions were added directly to the muscle bath at the points indicated in each record. The Tyrode's solution was replaced between drug additions and the motility was allowed to stabilize before the next addition of drug. Stimulation of the sympathetic nerve was performed at 15 pulses/sec, with a pulse duration of 0.5 msec and strength of 30-40 volts (A.C.), for 20 seconds. See text of the exercise for other details.
t Tyrode's Solution Added 10-7 M Acetylcholine
paper speed = 1 mm/sec
Tyrode's solution t Added 10-6 M Acetylchol ine
paper speed = 1 mm/sec
13 14 15 APPENDIX B
List of equipment/group of students: muscle bath and swivel assembly clamp for muscle chamber drain tube thermometer spool of thread 2-"S" hooks wax pencil 7 30-mL beakers 250-mL beaker Petri dish 100-mL graduated cylinder pipet bulb or Pi-Pump 0.1-mL pipet 1-mL pipet Pasteur pipet and bulb platinum wire electrodes stimulator (capable of delivering 10-100 volts A.C.) recording device (e.g. kymograph or Physiograph ) stand for holding transducer or recording lever plasticene
Materials at supply bench: extra plasticene recording paper for chart mover or kymograph extra Pasteur pipets and bulbs recording ink water bath at 37ºC containing 2-L bottles of Tyrode's Ringer solution carboy with extra Tyrode's solution (make up 2 L/group of students; we run through about 100 L/50 groups of students). the following drugs made up in Tyrode's solution: (make fresh prior to each lab*, and protect from light)
10-4 M norepinephrine 10-4 M atropine 10-4 M acetylcholine 10-4 M propranolol -4 -4 10 4 M phenylephrine 10 M phentolamine 10- M isoproterenol
* see p.18 concerning preparation of these dilutions.
16 Common Apparatus :
Warm water must be recirculated to each muscle bath. We use one water bath to supply two muscle baths. Water is pumped from the central water bath to each unit and back. Because of the temperature drop en route, we set the central bath thermostat to 40-45ºC, depending on the distance the water must travel and the velocity of flow.
If you wish to construct such a system, a sturdy submersible pump and Tygon tubing are necessary.
There are commercial organ baths available that contain built-in heating elements and a vertical muscle chamber.
A system for delivering a 5% C02-95% 0, gas mixture is necessary. We use one 200 cu. ft. tank of gas to supply 10 stations. This provides enough gas for five 3-hr laboratory sessions (with a 1/4 tank reserve). The gas stream to each muscle bath need only be enough to provide a steady stream of small bubbles.
Tygon tubing is used to deliver the gas to each station.
The gut segments survive much longer and the contractions are more regular using the 5% C02-95% 02 mixture than with air or pure 02.
Tyrode's Ringer Solution:
NaCl 8.0 g KCl 0.2 g
NaHCO3 1.0 g . MgCl2 6 H20 0.1 g . CaCl2 2 H20* 0.1 g . NaH2PO4 H2O 0.05 g Dextrose 1.0 g
* dissolve the calcium chloride separately and add last
Bring up to 1 L with distilled water.
17 Adrenergic and Chol inergic Drugs:
All the drugs mentioned in "Materials at supply bench" are available from Sigma Chemical Co., with the exception of phentolamine, which usually can be purchased through a local pharmacy. The names and Sigma catalogue numbers are shown below: drug name Sigma cat. No.
(-) norepinephrine (arterenol hydrochloride) A 7381
L-phenylephrine hydrochloride P 6126
( ± ) isoproterenol hydrochloride I 5627 acetylcholine chloride A 6625
DL-propranolol hydrochloride P 0884 atropine A 0132
Preparation of 10-4 M dilutions of each drug
To facilitate ease and accuracy in preparation, 100 mL of a 10-3 M stock solution of each drug is prepared at the beginning of the week and stored in a refrigerator. Acidify with 2-3 drops of 2.0 M HCl. Just prior to each laboratory session, a 1:10 dilution of the stock solution is made, using Tyrode's, to obtain 10-4 M.
18 APPENDIX C
Constructing the Muscle Bath
Figure 5 illustrates the muscle bath used in the measurement of the motility of the rabbit intestine. As shown in the diagram, the bath is constructed of 1/4-inch and 1/8-inch acrylic plastic. A water jacket surrounds the muscle chamber and allows warm water to be circulated on three sides of the chamber to keep the gut at 37°C. An aeration tube, made from P.E. 90 polyethylene tubing, is glued around the bottom of the chamber. The tube is perforated with small holes over its entire length to provide maximum aeration and mixing of the Tyrode's solution with a fine stream of bubbles. The shank of a 20 gauge needle is attached to the end of the aeration tube as a means of connection with an air line from a tank of compressed 5% C02-95% 02 gas. A small platform attached to one end of the chamber contains a pin hole. The platform serves as a holder for a push pin, which is used to anchor one end of the preparation in position. The gut segment is mounted horizontally and attached to a swivel and the push pin via "S" hooks (see Fig. 4). When attached to a writing stylus or a transducer lever with thread, the swivel serves to convert horizontal movements of the gut into verticle movements of the stylus or transducer lever. A drain is attached near the bottom of the muscle chamber. Except for the aeration tube, which is epoxyed to the bottom of the chamber, the entire unit is glued together with dichloromethane.
Constructing the Swivel Assembly
Figures 6a and 6b show the dimensions and placement of the swivel assembly which attaches to one side of the water jacket. The assembly is constructed from pieces of 3/4, 1/2, 1/4 and 1/8-inch acrylic plastic. There are three parts to the assembly; the swivel, the part that holds the swivel, and a track which is glued to the side of the water jacket.
The swivel is made from 1/8-inch plastic. The arms, which form a right angle, should be 5/16-inch wide and 2 inches on a side. A small hole is drilled at the end of each arm for attachment of the hooks or thread to the muscle and recording lever. Alternatively, the ends can be notched. A 3/32-inch hole is drilled at the intersection of the two arms to act as a pivot. When placed in the holder, the swivel is held in place by a 1/16-inch rod which acts as an axle, as shown in Figure 6b. Refer to Figure 4 for a side view of the swivel.
As shown in detail in Figure 6a, the holder for the swivel is constructed from pieces of 1/4 and 3/4-inch plastic. The swivel is mounted in a 5/8 x 1/4-inch groove as shown, with the axle extending through holes which hold it snugly in place. A bolt two inches long secures the holder onto a track (Fig. 6b) which is glued to the side of the water jacket.
The track is made from a piece of 1/2-inch plastic which extends the full length of the water jacket. A 3/16-inch slot is cut into the plastic as shown in Figure 6b. When held in the slot by the bolt, the holder for the swivel can be positioned to accommodate the length of the intestinal segment.
19 Constructing the Electrode
The stimulating electrode consists of a pair of platinum wires which are encased in an epoxy cuff (see Fig. 7). Two 2-cm pieces of 28-30 gauge platinum wire are soldered to two I-meter lengths of 20 gauge steel wire, which have been braided together. The solder joints are insulated with a layer of epoxy to keep them from making a short circuit. The platinum wires are then looped to form a circle about 3 mm in diameter. The inside curvature of the loop is filled with wax and then the terminal 2-3 cm of the electrode is dipped in epoxy several times to form a protective layer around the patinum wires. When the epoxy has hardened, the wax layer on the inside curvature of the electrodes is scraped or dissolved away to expose the bare platinum wire.
20 21 22 23 ACKNOWLEDGEMENTS
The authors wish to extend their gratitude to Dr. J. S. Davison and B. Greenwood of the Department of Medical Physiology, University of Calgary for their invaluable assistance in developing this exercise. Thanks are also extended to Mrs. Florence Robertson for her assistance in proofreading the manuscript.
24 REFERENCES
Davison, J. S. 1983. Innervation of the gastrointestinal tract. In: A Guide to Gastrointestinal Motility. Christensen and Wingate, (eds). Wright and Sons. Bristol, U.K.
Finkleman, B. 1930. On the nature of inhibition in the intestine. J. Physiol. 70: 145-157.
Goodman, L. S. and A. Gilman. 1975. The Pharmacological Basis of Therapeutics. 5th ed. MacMillan and Co., New York.
Nicpon-Marieb, E. 1981. Human Anatomy and Physiology. Benjamin/Cummings Publishing Co., Menlo Park, Ca.
Staff of Department of Pharmacology, University of Edinburgh. 1970. Pharmacological Experiments on Isolated Preparations. E. And S. Livingstone, Edinburgh, U.K.
25
Chapter 2
Tracheid Length Measurement In Selected Conifer Species
J. Tidswell, W. J. Mullin and K. G. Tidswell
Department of Biology University of New Brunswick Fredericton, N.B., Canada E3B 6E1
John Tidswell received his B.Sc.F. in Forest Management and his M.Sc.F. in Wildlife from the University of New Brunswick. He is currently a scientific technician in the Department of Biology at the University of New Brunswick. His research interests are in the area of upgrading science teachers. W. J. Mullin received his B.Sc. from McGill University and his Masters in Education from the University of New Brunswick. He is currently a senior teaching associate in the Department of Biology at the University of New Brunswick. His research interests are in developing individualized instruction. He is a board member of ISETA. K. G. Tidswell received her B.Sc. (biology) from the University of New Brunswick. At the time of this workshop she worked for the Maritime Forest Research Center, Government of Canada. Her area of interest is in the area of forest genetics.
27
INTRODUCTION
Most of the secondary xylem of conifers consists of longitudinally-arranged tracheids (sometimes called fibres). White pine wood, for example, is 95% by volume longitudinal tracheids (Panshin and de Zeeuw, 1980). Tracheids are long cells with rounded or tapered ends which overlap each other. They allow conduction of fluids and provide support (see Meylan and Butterfield, 1972, and Papermaking Fibres. A Photomicrographic Atlas, 1980).
The length of tracheids in wood is an important determinant of the use to which that wood is put (Hocker, 1979). The quality of both lumber (Daniel, Helms, and Baker, 1979) and pulp (Clark, 1978) is determined, in part, by the length of tracheids in the wood. Long fibres give lumber greater strength and paper produced from pulp more strength and fold-resistance. Tracheid length is also being investigated as a predictive factor in tree breeding programs (Daniel, Helms and Baker, 1979) because of observations that tracheid length is correlated with tree height (Echols, 1958) and appears to be an inherited characteristic (Zobel, 1961).
Tracheids are easily separated and can be measured using compound light microscopes or stereomicroscopes. They are therefore well suited for a laboratory practical teaching measurement using calibrated microscopes or for a project intended to draw students into an area of active research.
This paper describes an undergraduate project based on these concepts. Some of the factors which influence tracheid length are also discussed because each factor is potentially a source of new laboratory exercises. Though the project described in this paper used three conifers which are abundant in Canada similar techniques have been used with other conifer species (Echols, 1958, and Jagels, Gardner and Brann, 1982).
METHOD
The method described here is easy to manage in an open laboratory setting, but many refinements and alternatives are available. Jagels, Gardner and Brann (1982) is a good source of current information.
Small blocks of wood were cut from similar positions on similarly-aged local trees of three conifer species: Pinus strobus L. (Eastern White Pine), Picea glauca (Moench) Voss. (White spruce), and Abies balsamea (L.) MIll. (Balsam Fir). Each student cut two or three thin sections of approximately 20 mm x 1 mm from a block. Students were instructed to confine the cuts to one quadrant and within a few growth rings and this was supervised whenever possible.
29 Each thin section was placed in a small, brown bottle filled with a macerating solution of 1:1 (v:v) 30% hydrogen peroxide: glacial acetic acid. The bottle cap, lined with aluminum foil, was placed loosely on the bottle to allow gases to escape. (Students were warned that the solution is corrosive and taught appropriate safety procedures). The section was left in macerating solution at 60ºC for 48-72 hours, with adequate ventilation to disperse the escaping gases. During this time the section began to break into fragments and the tracheids became silvery-white. After cooling to room temperature, the macerating solution was removed and the tracheids were gently rinsed with several changes of distilled water. With the last change of distilled water, the container was covered and shaken 5-15 minutes with a regular motion. This motion which causes the tracheids to separate is a critical step in this method. The distilled water was replaced with 1% (w:v) active formaldehyde. Appropriate safety precautions with this chemical fixative were stressed. At this stage, the tracheids could be stored for periods of at least one month, a definite asset for laboratory management.
For microscopic examination, the tracheids were placed on a slide in either distilled water or 1% (w:v) basic fuchsin. After the technique of using and calibrating compound and stereomicroscopes was learned by the students, each student measured the length of fifty tracheids.
THE CLASS PROJECT
The project was conducted during the second month of an 'Experimental Laboratory Techniques' course for 60 undergraduate students, chiefly second-year biology majors. The advance preparation in light microscopy, including calibration, was given in this course in a series of mastery-based modules of laboratory work. For reasons related to our course objectives, this project was preceded by two additional exercises. A series of computer drills allowed students to become able to use the computer to handle data. An advance, individualized, mini-project gave each student the opportunity to apply information received in lectures about experimental design and execution and about data analysis.
With this advanced preparation, the project was introduced to the students. While they received all details of the method, all that was said about the project was that it was intended to determine whether the average tracheid length differed among Pinus strobus, Picea glauca and Abies balsamea. Students measured fifty tracheids each at their convenience over two weeks. The measurements were shown to the instructor who verified correct calibration technique then they were entered by the student into the class computer workspace. The importance of accurate data entry into the computer workspace was stressed. After all the data was entered, each student obtained a print-out of the computerized t-test analysis of the class project data. Provided with some information about factors which are known to influence tracheid length, each student prepared a scientific report for evaluation by the instructor.
30 PROJECT RESULTS AND DISCUSSION
Table 1 summarizes the class project data. Even with no direct supervision of the data entry into the computer workspace and no editing of the data once entered, there are no apparent data entry mistakes such as double digit entry or forgotten decimals. Since the project lasted only two weeks and involved sixty students, a few students measured less than the requested fifty lengths. The total number of observations for the three species was 2920. Though using the computer for this purpose was one of our course objectives, data can be analyzed by any available means.
We elected to manage this project in an open laboratory format and encountered no difficulties. We were limited to one fumehood so the students were "paced" by the availability of that facility. We feel that availability of fumehoods or other means of ventilation and microscopes are most likely to determine the format. Provided the facilities are available, this method and class projects based on it should also fit easily into more traditional laboratory time slots.
For this particular project, the students were asked to determine whether average tracheid length is characteristic of the species. The first step was to analyze the samples obtained by the class using a t-test analysis. Table 2 shows the result of the t-test analyses. The differences between sample means are all highly significant.
How this information should be interpreted was addressed in the discussion section of each student's scientific report of this project. With appropriate advance information about data analysis and evaluation, it should be clear to them that, while the sample means appear different, the differences are not necessarily attributable to species. In this particular project, some care was taken to control for factors known to influence tracheid length (Table 3), however not all of these factors could be controlled.
The information provided in-Table 3 can also serve as the starting point for other projects of this type. Among many factors lumped under growth conditions are latitude, altitude, longitude, and amount of rain and sun (Panshin and deZeeuw, 1980). The fact that enhanced growth rate often results in shorter tracheids has implications for woodlot managers who thin and fertilize their lots (Daniel, Helm and Baker, 1979). The assumption often made in tree breeding programs that longer tracheid length is advantageous still needs more testing in properly controlled experiments (Keith and Kellogg, 1981).
There is also the possibility of relating results like these to the pulp and paper industry. Clark (1978) describes in detail the type of measurement and reporting procedures used in the industry and provides formulae for calculating the weighted average length values which are used.
31 CONCLUSION
Tracheids in conifers are easy to separate and measure. The method is flexible enough to be used in fixed-hour laboratory periods or in open-laboratory project formats.
Conifer wood is an inexpensive source of material for laboratory practicals or microscope use. Both chemicals and equipment used in this exercise are readily available in most laboratories. Since there is a fair amount of information available about factors influencing tracheid length, a class can select and design their own project. The information, however, is not available for all conifer species so the project will be current. Tracheid lengths characteristic of the species make these measurements particularly suitable for classroom discussions of data analysis and evaluation.
32 Table 1. Lengths of tracheids in three conifer species. Class projec: data.
Species Number of Observations Numerical Average Length (%)
Pinus strobus Picea glauca
Abies balsamea 789 1.97 ± 0.81
Key: *, mean ± standard deviation
Table 2. Statistical comparison of numerical average tracheid lengths.
Comparison T in TTest* Probability T
Pinus vs Picea 3. 91** < 0.0001
Pinus vs Abies 29.50*** < 0.0001
Picea vs Abies 30.90**
Key: * Using "PROC TTEST", Statistical Analysis System **The hypothesis that variances are equal was rejected in a F' test (p < 0.001) so the T test assumes unequal variances. ***The hypothesis that variances are equal was accepted in a F' test (p < 0.5) so the T test asumes equal variances.
33 Table 3. Some factors influencing conifer tracheid length.
A. Within a single tree
maturity of the Layer of wood age of wood across a single layer vertical position within a single Layer growth rate
B. Within trees of the same species
growth conditions genetic make-up
C. Between species
genetic (?)
References:
Keith and Kellogg (1981), Daniel, Helms and Baker (1979), and Panshin and de Zeeuw (1 980 ) .
34 REFERENCES
Clark, J. D'A. (1978). Pulp Technologv and Treatment for Paper.
San Francisco: Miller Freeman Publications, Inc.
Daniel, T.W., Helms, J.A., and Baker, F.S. (1979). Principles of
Silviculture. 2nd edn. New York: McGraw Hill Bock Company.
Echols, R.M. (1958). Variation in tracheid length and wood density in
geographic races of scotch pine. Yale Universitv School of Forestry
Technical Bulletin No. 64.
Hocker, H.W. Jr. (1979). Introduction to Forest Biology. New Ycrk: John
Wiley and Sons.
Jagels, R., Gardner, D.J., and Brann, T.B. (1982). Improved techniques
for handling and staining wood fibres for digitizer assisted
measurement. Wood Science 14: 165-167.
Keith, C.T. and Kellogg, R.M. (1981). The structure of wood. In Canadian
Woods Their Properties and Uses. 3rd edn., ed. Mullins, E.J. and
McKnight, T.S. Toronto: University of Toronto Press.
Meylan, B.A. and Butterfield, B.G. (1972). Three-dimensions1 Structure
of Wood. A Scanning Electron Microscope Study. Syracuse: Syracuse
University Press.
Panshin, A.J. and deZeeuw, C. (1980). Textbook of Wood Technology. 4th
edn. New York: McGraw-Hill Book Company.
Papermaking Fibres. A Photomicrographic Atlas (1980) ed. Cote, W.A.
Syracuse: Syracuse University Press.
Zobel, B. (1961). Inheritance of wood properties in conifers. Slivae
Genetica 10: 65-70.
35
Chapter 3
Community Ordination Utilizing Winter Stoneflies
Vinnedge M. Lawrence
Department of Biology Washington and Jefferson College Washington, PA 15301
Vinnedge M. Lawrence is a Professor of Biology at Washington and Jefferson College, Washington, Pennsylvania. He received his B.S. (education) and M.A. (zoology) degrees from Miami University and his Ph.D. (entomology) from Purdue Univeristy. In addition to introductory biology, he teaches courses in ecology, entomology, invertebrate zoology, developmental biology, and aquatic biology. His research interests include population dynamics of odonate naids, distribution of winter stoneflies, and breeding behavior of Henslow's sparrow.
37
COMMUNITY ORDINATION UTILIZING WINTER STONEFLIES
Vi nnedge M. Lawrence Washington and Jefferson Col lege
Objectives. This exercise in community ecology can be conducted in the field in mid-winter when a course offered during the spring term may otherwise be restricted to indoor laboratory exercises. It encourages students to make systematic observations in the field. It combines field and laboratory activity with library research, thereby introducing students to various aspects of scientific endeavor. It involves students in original research without a predictably correct outcome. It uses a computerized key to overcome the resistance exhibited by many students to taxonomic work.
Background. An instructor may be reluctant to take a class into the field for investigations that require interpretation of environmental conditions. It may be impossible to determine those factors that most directly influence the outcome of the investigation. This exercise enables students to obtain data with which they can calculate and graph differences between communities. Each student may then apply observations in accounting for these di f ferences.
Although lack of knowledge of critical environmental factors is not a problem in this exercise, the instructor must locate suitable sites for collecting winter stoneflies. These are insects of the order Plecoptera that are members of the families Taeniopterygidae and Capniidae, although these taxa sometimes are treated as subfamilies of the family Nemouridae. Sites may be located during the winter months by examining bridges that cross streams of good water quality. Peak emergence periods vary with latitude, as Taeniopteryx maura, for example, emerges from mid-December through January in Alabama, from January through March in Virginia, and through mid-April in Nova Scotia. Since bridges are favored emergence sites, locating winter stoneflies should not be difficult in areas with at least a few streams with reasonably good water quality.
Developing a computerized key requires familiarity with the winter stonefly fauna of an area. The key should include every taxon that has been collected, with the possible addition of previously uncol lec ted t axa whose distributions over lap the region of the study. A good way to accomplish this is to engage one or more students in independent studies to catalog the local fauna. My experience suggests that an area limited by easy access for field trips, which may be a circle with a 15-mile radius, will include a manageable number of species that will not over whel m students unfamiliar with stonefly taxonomy. The number probably will be small enough to allow you to include several additional taxa in your key. The key used in this workshop includes four species that definitely occur at my study sites and four others that have not been found or confirmed in the area. Stoneflies neither bite nor sting, and have no characteristics that most students would find objectionable. Many of my students are fascinated to find adult insects in winter in places so accessible, but which they have
39 overlooked. Procedure. Depending on the instructor's preference, this exercise may require from two to five three-hour laboratory periods. Assuming that five periods are devoted to the project, the first session would occur in the laboratory, followed by two periods of gathering data in the field, after which a fourth period is alloted for identification, and the fifth is used to explain statistical procedures and other details of format to be followed in writing the report of the exercise.
Period 1. I introduce the recording barometer in the laboratory, the sling psychrometer, and Hach Chemical Co. kits for determining pH, dissolved oxygen, ni trate, nitrite, and orthophosphate. After orientation in the correct use of this equipment, each student obtains physical and chemical data in a "dry run" of the procedures each will follow in the field. The barograph is read, relative humidity is determined outside, and prepared water samples arc analyzed. Over the next two weeks, each student will be required to carry out some or all of these procedures at one or more sites in the field.
Period 2. The class travels to an area where a stream system affords at least four bridges that will serve as collecting sites which can be sampled during the period. The sites should be at different tributaries or streams of different sizes. At each bridge, one student obtains physical and chemical data while all other students collect stoneflies from the structure and its immediate vicinity during a 10-minute period. Stoneflies are collected with forceps and placed in homeopathic lip vials that are half filled with 70% ethanol and contain labels indicating the collector's name, date, and collection site. At the end of the collecting interval , a green neoprene stopper is inserted in the vial with the aid of a bent paper clip to prevent pressure buildup from trapped ai r that could subsequentl y loosen the stopper and permit evaporation of contents. Screwcap vials, although more convenient in the field, introduce a high probability of evaporation and resultant destruction of specimens.
Following the collecting interval, time is allotted for taking notes on characteristics of the site. These may include estimates of cloud cover, extent of canopy cover when leaves would be present abundance of leaf packs in the water, substrate type in terms of particle size, extent and proximity of riffles upstream of the site, presence and location of tributaries, size of stream above the site, time of collection, and other pertinent information. By now the student gathering physical and chemical data has completed this activity. This includes obtaining water and air temper atures, relative humidity, pH, a water sample fixed for subsequent dissolved oxygen t itrat ion, and a separate water sample for subsequent determination of nitrate, nitrite, and orthophosphate. When we return to the laboratory, we determine the chemical characteristics of the water samples and read the barometric pressure that has been recorded for each of the four collecting intervals.
Period 3. We carry out the same procedures as in the second 40 period, but on a different stream system. The addition of these four rites brings to eight the number of sites we will compare on the basis of their winter stonef l y communi t ies.
Period 4. Although students may begin identifying their specimens as soon as they have collected them, I usually devote a laboratory period to assisting them in this endeavor. Students who have already completed their determinations nay wish to reconsider some of them. Those who haven't finished are now imbued with a sense of urgency to do so. Each student is required to submit all stoneflies in their proper vials along with a determination sheet on which the sex and species of each specimen are indicated. I make spot checks for accuracy and examine the entire contents of a collection if I detect errors. Students receive credit for their collections based upon the total number of specimens and the number properly identified. There may be 50 points for the collection, with the number of misidentifications subtracted from the number of spec i mens submitted, t tie number proper ly i dent i f i ed , or some other formula that weighs both collecting effort and accuracy of identification.
Since each workshop participant will identify the stoneflies in a sample vial, here are some characteristics to watch for in making your deter mi nat ions.
1. The last segment of the leg of an insect is the tarsus. In stoneflies, ttie tarsus is divided into three subsegments known as tarsomeres. Carefully compare the relative lengths of the tarsomeres on a leg. In the family Taeniopterygidae, the 2nd tarsomere is subequal to (about as long as) the 1st or the 3rd. In the Capniidae, the 2nd tarsomere is much shorter than the 1st or 3rd. Since capniids are called small winter stoneflies, you know two characteristics that distinguish them from taeniopterygids.
2. The last abdominal segment of many insects bears a pair of appendages called cerci. The cerci of stoneflies tend to be long and filamentous, but are reduced in Taeniopterygidae where they also exhibit sexual dimorphism. Taeniopteryx females have short, six-segmented cerci, whereas the cerci are represented by a single, inconspicuous segment in males.
3. The cerci of Capniidae are long, but sexual dimorphism is evident in the recurved, supra-anal process of the male that is absent in the female. The supra-anal process recurves anteriorly over the 9th and 10th abdominal segments. It consists of a single element in the genera Nemocapnia and Paracapnia and in Capnia vernalis. In other species of Capnia and in the genus Aliocapnia, it is comprised of two distinct elements, one dorsal and one ventral.
4. The hind wings of capniid stoneflies exhibit a marked dichotomy with respect to the extent of the posterior, flexible lobe, known as the vannal lobe or vannus. In the getier a Capn ia, Nemocapnia, and Paracapn ia, the vannus is of normal size (Fig. 1), whereas it is exceptionally
41 large in members of the genus Allocapnia in which it
vannus
Paracapnia A llocapnia Figure 1: Hind wings of Paracapnia and Allocapnia.
With these characteristics in mind, obtain a vial of stoneflies and a Syracuse watch glass or culture dish partially filled with water. Locate your stonefly identification sheet. Record the date and site from the label in the vial onto your identification sheet (the site is recorded by circling the appropriate number). Remove the stoneflies from the vial with forceps and place them in the water. Place the watch glass or culture dish on the stage of a stereomicroscope and adjust the i lluminator. Insert a minidisk labeled "Stoneflies" into the disk drive and plug in the microcomputer and monitor. When asked for the program you want, type in "Stoneflies" and press the "RETURN" key. When the program menu display appears, select the dichotomous key by typing in "K". The program will begin illustrating characteristics and requesting yes (Y) or no (N) responses. Type in only the appropriate letter. Use a pair of teasing needles to position a specimen so as to determine the characteristic being requested. Continue until the program indicates the species and sex of the specimen. Record the species name on the identification sheet and place a hash mark in the space corresponding to the proper site and sex. Return this specimen to the vial and proceed to determine another specimen. Continue this procedure until you have identified each specimen. When al l specimens are back in the vial,, insert the straightened end of a paper clip along with the neoprene stopper into the mouth of the vial. When the stopper has been twisted in securely, hold it in place with your thumb and withdraw the paper clip. Follow the same procedure with any other vial for which you are responsible for determining the contents. Now transfer your data to another identification form on which the totals of the hash marks are entered as numbers. Submit this identification form so that your data are included with those obtained by other members of the group. Pour the water from your otherwise empty watch glass or culture dish and dry it with a paper towel. You may wish to attempt the quiz or examine other parts of the STONEFLIES program while the group results are being tabulated.
Period 5. We now proceed to analyze our stonefly data by constructing a community ordination to compare collection sites. Each bridge is treated as a community and analyzed for its degree of similarity or dissimilarity to every other bridge on the basis of their respective stonef l y faunas.
For the sites whose fauna you helped determine, find the frequency for each species. The frequency, in this context, is the percentage of samples obtained in which a particular species
42 43 occurred. Now find the number of individuals of this species in all samples combined, and record this as the densi ty of the species. Calculate a prominence value for each species by multiplying its density by the square root of its frequency. Sum the prominence values for all species found at the bridge. This information must be determined for every site.
Construct a matrix of similarity by filling in the portion of the chart on Form 1 above and to the right of the diagonal. The value to be placed in each block is the coefficient of similarity between the two sites that intersect at the block. The coefficient of similarity (C) is calculated by the formula:
where a = the sum of the prominence values for one bridge? b = the sum of the prominence values at the other bridge, and w = the sum of the lower prominence values of species common to both bridges. Assume, for example, that species X, Y, and Z occur at bridges A and B. Their prominence values at Bridge A are X = 10, Y = 63, and Z = 1. At Bridge B, X = 1, Y = 33, and Z = 36. To find w, the lowest values for each species common to the two bridges are summed; thus w = 1+33+1, or 35. The coefficient of similarity for these bridges would be:
or .486, a value near the middle of the similarity index which ranges from 0.000, if no species is common to both bridges, to 1.000 when every species occurs at both bridges with the same numbers of individuals present at each. Place your results in the proper spaces of the similarity matrix of Form 1.
It is necessary to convert similarity coefficients to values expressing dissimilarity because the distances between communities, as they will appear on the ordination graph (Form 2), represent degrees of difference rather than similarity. Dissimilarity values are obtained by subtracting the calculated coefficient of similarity for each community from its theoretical maximum similarity value of 1.000. Place these results in the proper spaces of the matrix of dissimilarity of Form 1.
To determine the two most dissimi lar communities for placement on the x-axis of the ordination graph, sum the dissimilarity values associated with each bridge. List each community and the sum of its dissimilarity values in the chart at the bottom of Form 1. The bridge with the greatest dissimilarity sum is assigned a value of zero and placed at the left end of the x-axis. Actual placement of points on the graph must await determination of values for the y-axis. Do not begin your graph until the values of all coordinates have been determined. Identify the bridge located at zero on the x-axis as bridge "a", and refer to the matrix of dissimilarity for the bridge exhibiting the greatest dissimilarity to it. This bridge, designated "b", will be placed at the opposite end of the x-axis, and the length of the x-axis will be made equal to the dissimilarity of the two reference bridges. The distance (x) of
44 each of the remaining bridges from bridge "a" along the x-axis is obtained from the formula:
where L = the distance between the two reference bridges (or dissimilarity value between bridges "a" and "b"), Da = dissimilarity value between bridge "a" and the bridge being plotted, and Db = di ssimi lar it y value between br idge "b" and the bridge being plotted. Record these values in the appropriate column in the chart.
Even after the dissimilarities between bridges have been plotted on the x-axis, some communities that may be quite dissimilar could still be placed close together. Their placement on the y-axis coordinate should be accomplished in such a manner that the maximum component of the remaining dissimilarity is spread along the y-axis. To achieve this, the bridge with the poorest fit on the x-axis must first be determined. A poorness of fit value (e) is calculated for each bridge by the formula:
Record your calculated e values in the appropriate column of the chart. The bridge exhibiting the greatest e value is designated a' and assigned a value of zero on the y-axis. The bridge with the greatest dissimilarity to a' and located within 1/10 L of a' on the x-axis is designated b' and placed at the opposite end of the y-axis. The 1/10 L restriction may necessitate a compromise in that any bridge located so near a' on the x-axis may not exhibit much dissimilarity to it. Even though this may be the case, disregarding the 1/10 L restriction could result in the x-axis merely being tilted on the graph, thus negating any true 2nd dimension that the y-axis may otherwise have conferred. When the two reference bridges, a' and b', have been selected, the length of the y-axis is made equal to t hei r di ssi mi l ar it y.
The distance (y) of each of the remaining bridges from bridge a' on the y-axis is obtained from the formula:
where L' = the dissimilar it y val ue between br idges a' and b' , etc. Record these values in the appropriate column in the chart. The positions of the communities may now be plotted on Form 2 which should consist of a two-dimensional graph constructed on regular graph paper. The graph should be entitled "Two-dimensional ordination of communities on the basis of prominence values of winter stoneflies."
The ordination interval, or direct distance between any two communities in the ordination graph, may be calculated by the formula: ordination interval = , where dx = distance between communities on the x-axis and dy = distance between communities on the y-axis. The extent to which the spacing of the communities on the two-axis ordination accounts for
45 46 the calculated dissimilarities in community composition may be estimated by correlating ordination intervals with dissimilarity values for at least 20 random1 y selected community pairs. If the ordination interval is designated X and the dissimilarity value Y, the correlation coefficient (r) is determined by the formula: -
where N = number of community pairs used in making analysis. For this analysis, 20 sets of values should be chosen with the use of a table of random numbers and entered in Form 3. The higher the correlation coefficient, the more satisfactorily the two-axis ordinat ion accounts for the calculated dissimilar i tes between communities. A correlation coefficient above 0.90 indicates a very satisfactory accounting.
Discussion. One of the values of the ordination graph is that it may help indicate those environmental factors that are most important in influencing community composition. This is illustrated by Finni (1973) for winter stonefly communities in Little Pine Creek during the winters of 1967-68 and 1968-69. After the ordination has been constructed, environmental gradients corresponding to the x- and y-axis variables should be sought and plotted along these axes. The number of stoneflies collected at a site may be influenced by air temperature and amount of insolation at the site during and prior to the collecting interval. Stonefly abundance at a site may be influenced by the number and size of leaf packs upstream. Prevalence of leaf packs may, in turn, reflect extent of riffle areas to catch leaves and extent of canopy cover in summer to provide leaves. The occurrence of diapause in the life history of a species may influence its presence at a site. Permanent streams can support species that are absent from streams that dry up periodically. Species whose nymphs undergo diapause in the hyporheic zone may be particularly abundant in intermittent streams where competition from non-diapausing species is lac king. Small , spring-fed tributaries may support populations of small, non-diapausing species that require very little flowing water, but must be in water throughout their development.
This exercise should include the requirement of a laboratory report written in a style reflecting proper format for scientific writing. Helpful suggestions are provided by Brower and Zar (1984), who have produced a very useful ancillary text for ecology courses that i ncludes a table of random numbers, common statistical procedures and tables, and detai led methodologies for various ecol ogi cal investigations.
Literature Cited
Brower, J. E., and J. H. Zar. 1984. Field and laboratory methods for general ecology. 2nd ed. Wm. C. Brown: Dubuque, Iowa.
Finni, G. R. 1973. Biology of winter stoneflies in a central Indiana stream (Plecoptera). Ann. Entomol. Soc. Am. 66: 1243-1248
47 48 Sources of Materials
Hach Company P. 0. Box 389 (800) 525-5940 Loveland, CO 80539
Cat. No. Item 1802-02 Water Ecology Test Kit , Model AL-36B 1475-00 Orthophosphate Test Kit, Model PO-14 14081-00 Nitrate-Nitrite Test Kit, Model NI-12
VWR Scienti fic P.O. Box 8188 (800) 257-8407 Philadelphia, PA 19101
Cat. No. Item 59589- 110 Green Neoprene Stoppers, Size 0 66010-129 Homeopathic Lip Vials,3 dram 66010-140 Homeopathic Lip Vials, 4 dram
Weat her Measur e Cor por at ion P.O. Box 41257 (916) 481-7565 Sacramento. CA 95841
Model No. Item HM 10 Sling Psychrometer B201-S Barograph with spr ing-wound clock
Ac knowledgement
I greatly appreciate the contributions of Dr. Thomas Hart (Department of Biology, Washington and Jefferson College) and Dr. Rick Pei fer to the preparation of this workshop. Tom wrote the computer program STONEFLIES to provide my students with an alternative to the other taxonomic keys I provide in my course. Rick was most helpful in gathering the equipment and supplies to run the wor kshop.
49
Chapter 4
The Use of Echosounding Equipment in Limnology and Ecology Classes
Peter W. Vaughan
Department of Ecology and Behavorial Biology University of Minnesota 31 8 Church St. S. E. Minneapolis, MN 55455
Peter Vaughan received his B.A. degree from Dartmouth College in 1978, and is a Ph.D. candidate in the Department of Ecology and Behavorial Biology at the University of Minnesota. He has held a Bush Fellowship and an instructorship in General Biology at the University of Minnesota. He currently holds a Dissertation fellowship and expects to defend his thesis in June of 1988. The focus of his reserch is the role that light plays in mediating between zooplanktivorous fish and their prey.
51
THE USE OF ECHOSOUNDING EQUIPMENT
IN LIMNOLOGY AND ECOLOGY CLASSES
INTRODUCTION: Chart recording echosounders of the kind now widely used for sportfishing in lakes are valuable for describing the spatial and temporal distribution of fish, zooplankton and rooted aquatic plants. The equipment employs high-frequency sound, typically near 200 kHz, for use in shallow, freshwater lakes. Sound of this frequency is reflected by objects in the size range of cladocera and copepoda (0.5 - 5mm diameter), which are typically the predominant freshwater zooplankton (McNaught 1968, 1969). Of course, larger objects, such as fish, also reflect these frequencies.
The instrument we will demonstrate is a Lowrance X-16 computer sonar. It features a high resolution printout (125 lines per inch), variable pulse length
(30 to 2,000 us) and dual transducers ( 8 or 20 degree beam angle). It operates off of a 12 volt battery and may be purchased through the manufacturer, at an educational institution discount, for about $575.00, Depending on the accessories you choose.
The shortest sonic pulses (30 us) emitted by the the Lowrance X-16 echosounder can discriminate between objects that are separated from each other
by a vertical distance of 2.5 cm. Echotraces can be recorded from objects in all depths beneath the water surface, or a range of intermediate depths can be selected and traces from objects only in this increment recorded. The operating characteristics are controlled by a microprocessor so that different
53 combinations of depth range, chart speed, acoustic pulse duration, and other
characteristics can be selected rapidly with a keyboard. These features enable
the equipment to provide much information that is not detected by conventional
sampling with nets. The technique can be used quantitatively and obviates many
of the problems associated with traditional sampling, such as: active net
avoidance by and patchy distribution of zooplankton, working with hazardous
preservatives, and tedious and time consuming counting of samples. The
echograms obtained are concise graphic records that show instantaneously how
populations of zooplankton and pelagic fishes are distributed with respect to each other, to features of underwater topography, and to areas occupied by
rooted aquatic plants. Depths for conventional sampling can be selected judiciously based on this information . Zooplankton are typically distributed heterogeneously, and the density of echotraces varies correspondingly along the depth-scale of an echogram.
Scattering layers, depths where zooplankton are abundant, often are conspicuous features which appear as bands of high density tracings. The depth limits of scattering layers sometimes coincide with layers of water that can be
distinguished from each other on the basis of temperature and concentrations of
dissolved substances, but some scattering layers are not closely related to water strata. Depths of scattering layers often change somewhat during a day as
zooplankton migrate vertically (Northcote, 1964). Therefore, boundaries of scattering layers generally represent instantaneous depth distributions of
different zooplankters. Environmental conditions at these boundaries thus appear to be of critical importance for the physiology and behavior of
zooplankton.
54 WORKSHOP DEMONSTRATION: We will use this workshop to demonstrate the use of echosounders in obtaining information about a number of physical and biological parameters in Lake Minnetonka, and to discuss several other possible applications for class or individual student projects.
Lake Minnetonka is a large lake consisting of a group of interconnected
"kettle hole" basins situated in glacial drift in eastern Hinnesota, about 20 miles west of Minneapolis. The lake became an important vacation and tourist area in the late 19th century, and subsequently became a commuter suburb of
Minneapolis in the 1950's. Lake Minnetonka has been the subject of extensive limnological study, largely because planktonic algae became very abundant due to increased sewage effluent to the lake that diminished the lake's recreational value (Megard, 1977). Sewage diversion in the early 1970's resulted in an amelioration of this concern, and in much of the lake algae are now much less abundant. It is the home of the Gray Freshwater Institute, which is associated with the University of Minnesota.
The demonstration will be done in Crystal Bay (see map), which has an area of 6 2 3.4 * 10 m , a mean depth of 8.5 m and a maximum depth of about 30 m. The procedure we will follow is given below:
1) Two complete transects of the bay at constant boat speed and echosounder settings will be made. This will:
a) provide a sketch of the morphometry of the basin.
b) demonstrate how this equipment can be used to survey rooted aquatic
macrophytes.
c) determine the depth of the mixed layer and any horizontal
irregularities that might be present due to internal seiches.
55 d) indicate that this technique can not only locate individual fish, but
also be used to survey fish populations and their distribution.
e) indicate depths at which scattering layers exist, and which are
therefore of interest to sample more extensively with traditional
techniques.
f) indicate a location in the basin in which to do the more extensive
sampling.
g) If meteorological conditions permit, these transects may also allow the
demonstration of Langmuir circulation and the concentration of zooplankton
in regions of current convergence.
2) A sampling location will be chosen (traditionally, the deepest spot in the lake is used) and we will double anchor to provide a stationary platform on which to work. The following measurements will then be made:
a) a remote sensing thermistor (available from Fisher for $400.00) will be
used to construct a vertical temperature profile.
b) An oxygen meter (available from Fisher for about $500.00) will be used
to construct a vertical oxygen profile. Oxygen may be measured less
expensively and easily by the Winkler titration method (American Public
Health Assoc. 1985)
c) A three-liter Van Dorn water sampler (Wildco, $300.00) will be used to
collect water samples at designated depths. The samples will be filtered
on board using a hand-held vacuum pump/filtration system (Fisher, $45.00)
and 0.45um glass-fiber filters. The chlorophyll samples thus obtained
will be visually examined, but it is a simple procedure to measure the
chlorophyll quantitatively (Am. Pub. Health Assoc. 1985).
56 d) A Schindler trap (Wildco, $300.00) will be used to sample zooplankton
at designated depths. Zooplankton nets could be substituted; they are
more laborious but much less expensive. The samples will be inspected
visually and if time permits, microscopically.
3) We will return to the Gray Institute to examine and discuss our results. Correlations between the echotracings, temperature, oxygen, chlorophyll and
zooplankton profiles will be examined, and a summary figure constructed.
OTHER APPLICATIONS SUITABLE FOR CLASS PROJECTS: The echosounder is a versatile
tool, and may be easily adapted to a variety of ecological and limnological
studies. A few are mentioned below:
a) Surveys and estimates of fish, zooplankton, and macrophyte distribution
and abundance.
b) Monitoring of the amplitude and timing of zooplankton diel vertical
migration.
c) Demonstration of correlations between zooplankton and other
biologically significant parameters, such as: phosphorus (which
zooplankton are thought to recycle and secrete), chlorophyll (which
zooplankton eat), and oxygen and carbon dioxide (which zooplankton consume
and generate respectively).
57 REFERENCES
American Public Health Association. 1985. Standard Methods for the
Examination of Water and Wastewater. 14th Ed. Amer. Pub. Health Ass.
Washington, D.C
McNaught, D.C. 1968. Acoustical determination of zooplankton distributions.
In Proceedings of the 11th conference on Great Lakes Research. p. 76-84.
McNaught, D.C. 1969. Developments in acoustic plankton sampling. In
Proceedings of the 12th Conference on Great Lakes Research. p. 61-68.
Megard, R.O. 1977. Phytoplankton, phosphorus, and sewage effluents in Lake
Minnetonka. EPA-600/3-77-086. North American Project- A Study of U.S.
Water Bodies. E.P.A. Corvallis, Oregon.
Northcote, T.G. 1964. Use of high frequency echo sounder to record
distribution and migration of Chaoborus larvae. Limnol. and Ocean.
9:87-91.
58 Figure 1. Acoustic scattering layers in Elk Lake during mid-morning, 21
July 1986, recorded along a transect marked with dashed line on the lake map. Top echogram recorded during a traverse from shallow to deep water, and bottom echogram recorded during the return to shallow water. Note 0 scales for depths on the right and scales for temperature ( C) and - 1 dissolved oxygen (mg liter ) on the left. Echosounder operating 0 characteristics: Top echogram, 8 transducer, 200 usec sonic pulse 0 duration, sensitivity 2, grayline 1; bottom echogram, 8 transducer, 50 usec sonic pulse duration, sensitivity 7, grayline off.
59 60 61 APPENDIX
EQUIPMENT LIST FOR WORKSHOP
1) Lowrance X-16 computer sonar. $575.00 with educational discount. Talk to Charley Ramsey 1-800-331-3889. Lowrance Electronics Inc., 12000 E. Skelly Dr., Tulsa, OK 74128.
2) Remote sensing thermister. $500.00. Many offices, we use Fisher Scientific, 10230 West 70th St., Eden Prairie, MN 55344. (612) 941-5460. 3) Oxygen meter. $500.00. Fisher Scientific. See #2 above.
4) Van Dorn water sampler. $300.00. Wildco, 301 Cass St., Saginaw, MI 48602. (517) 799-8100.
5) Schindler trap. $300.00. Wildco, see #4 above.
6) Nalgene vacuum pump (hand-held). Nalgene Labware Department, Nalge Co., Box 365, Rochester, NY 14602. (716) 586-8800.
62 Chapter 5
A Handbook for Collecting Releve Data in Minnesota 1
John C. Almendinger
Minnesota Natural Heritage Program Department of Natural Resources Box 7, 500 Lafayette Road St. Paul, MN 55146
John C. Almendinger received his B.A. degree from Ohio Wesleyan University in 1976 (botany) and his Ph.D. degree from the University of Minnesota in 1985 (ecology). He is a plant ecologist for the Minnesota County Biological Survey, Minnesota Department of Natural Resources. His research interests include paleoecology, phtyosociology, and the ecology of rare species.
1 This is one of several versions of the Releve Handbook. Omitted from this version are several length appendices that cover topics specific to Minnesota. A complete version of the Handbook and information on data management systems for releves are available from J. C. Almendinger at the above address.
63
INTRODUCTION -- HANDBOOK FOR COLLECTING RELEVE DATA IN MINNESOTA
INTRODUCTION
The procedure for collecting and summarizing vegetation data at almost any scale involves four "essential steps" (Mueller-Dombois and Ellenberg 1974). First, the researcher must define the area to which the study applies and then segment the study area into preliminary vegetation units. Second, for each preliminary vegetation unit. the researcher must select the stands to be sampled. Third, a sampling method must be chosen. And fourth, the researcher must decide what properties of the vegetation to measure. Several approaches might be adopted to complete these steps, and choosing the best or most practical approach requires (1)a clear definition of the project purpose and (2)f i eld reconnaissance.
The purpose of this handbook is to explain how the Minnesota Natural Heritage Program (MNHP) has approached these four essential steps to accomplish specific goals and al so to promote general interest in studying the vegetation of Minnesota. The first section of the handbook -- Using Releves to Study the Vegetation of Minnesota -- describes in general whet the goals of the MNHP are and how the releve method of vegetation analysis (Mueller-Dombois and Ellenberg 1974, Westhoff and Van Der Maarel 1978) is used to meet those goals. This first section also describes the initial classification of Minnesota's vegetation and how sample stands are Located in those units (steps 1 & 2 above). The second section of this handbook -- The Minnesota Natural Heritage Program Releve Methods -- describes in detai l the technical aspects of the releve sampling method as practiced by the MNHP staff [steps 3 & 4 above]. This section includes both releve field methods, and the transcription of releve site information from maps and other scientific literature.
The Minnesota Natural Heritage Program encourages researchers working in Minnesota to contribute their releve data to the MNHP database. By doing so, contributors gain access to the entire releve database and to the MNHP staff for assistance in any phase of col lecting or analyzing releve data. By fol lowing the guidelines set forth in this handbook, researchers are assured that releve data of comparable quality can be obtained to extend thei r datasets or to construct datasets for hypothesis testing. Individuals or institutions needing vegetational data or distributional data on particular plant species should contact the MNHP for more information.
65 SECTION ONE -- USING RELEVES TO STUDY MINNESOTA'S VEGETATION
General Description of a Releve Sample
Releve is a French word with one connotation that translates to the English word "abstract" (see Mueller-Dombois and Ellenberg 1974). As a Literary abstract is a concise summary of a body of text, a releve is a concise summary of a unit of vegetation. Note that both the releve and the li terary abstract are used to characterize a predetermined unit. Thus, releves are used to characterize larger vegetation units that have already been classified according to some criteria. These criteria might be related to geography, physiognomy, canopy dominants, or environmental gradients. The criteria selected depend enti rel y upon the purpose of a particular study. Releve data can be used to express the variability of the vegetation within each predetermined unit and also to express the hierarchal relationship among those units.
Physically, releves are small plots, within which the vegetation is described both structural ly and compositionally. Releve plots are typicall y square and may cover 25 to 1000 square meters. The vegetation structure is recorded by estimating the collective cover of taxa in life-form groups that form the height strata characteristic of the stand. For example, the strata recorded in a deciduous forest would typically include three strata of woody, deciduous species (trees, shrubs, and seedlings), as well as separate stratum Layers for graminoids, mosses, and herbs. Then, for each stratum. the cover of each component species is recorded. Thus, plant occurrences are recorded by Life-form group, then by height stratum, and finally, by species -- making it possible fur a species to be doubly recorded if it occurs in different height strata or in different Life-form groups. Releve data, therefore, are mul tidimensional and may be numerical ly ordi nated or classified on a physiognomic/life-form basis or a floristic basis.
General Description of the Releve Method of Analysis
The releve method has come to represent not only a method of vegetation sampling, but also a method of data analysis. At the heart of the releve method of analysis is the phytosociological table. The rows of a phytosociological table record the occurrence of a particular species in a set of releves (columns). Thus, the cel ls of the species by releve matrix contain information about a species (row designate) in a particular releve (column designate). Cel l values are usual l y cover estimates, which are often accompanied by an indication of how that species is distributed within the releve plot (sociability). The rows and columns of a phytosociological table are arranged so that spacies hav ing simi lar distibutions among the rel eves are placed in contiguous rows, and releves having similar sets of species are placed in contiguous columns. A table thus arranged (Table 1.1) is an effective way to visual ly summarize releve data, and it provides a basis for a hierarchal classification of the releves.
66 SECTION ONE -- USING RELEVES TO STUDY MINNESOTA'S VEGETATION
Table 1.1 A sample phytosociological table showing how releves with simi lar species occurrences are placed in contiguous columns and how species with similar distributions among the releves are placed in contiguous rows. The table elements a re Braun-Blanquet cover values. (The table data were selected from Almendinger, 1 985)
ReleveNumber 253345 235 1 444 21221 55124 980267 46556 78901 34045 23565 34608
Hieraci um scabrum 12221. Equisetum hyemale 2.1.2. Epigea repens ...2. Polygala paucifolia .22222 ...... Picea glauca .211.1 ..2...... Py rola virens .212.. ..2......
Linnaea borealis .....2 ...22 2.22. .21.2 ...... 2 Betula papyrifera ....2. ...32 .22.1 2..2. 2 ...... Acer rubrum ...... 3. .23.2 2.221 ...... Monotropa uniflora .....2 ..2.2 ..... 22.1...... Py rola rotundifolia ...... 2...... 1222 ......
Smilax herbacea ...... 1...... 21.21 2.11. Populus t remuloides ...... 3...... 3 ...... 3.2. 2...2 Puercus borealis ...... 2.. 1.1.2.21.. Quercus macrocarpa ...... 2.2 -52.. Amorpha canescens ...... 2...... 2.... .21..
Prunus serotina ...... 1.1...... 22122 Cornus racamosa ...... 2.2 Viburnum rafinesquianum ...... l. ... .1 ,2212 Pyrola asarifolia .2...... 2...... 21 Viola sororia ...... 2. 2....
The procedures for arranging the rows and columns of a phytosociological table were developed in Europe by Braun-Blanquet (1928, 1932) and his associates at Zurich and Montpel lier (see Westhoff and Van Der Maarel 1978 for European references). These procedures were the subject of consi derab le debate in Europe, which inevitably led to the formation of vegetational ideology and terminology. The ideology of the followers of Braun-Blanquet (the Zurich-Montpel lier School, see Becking 1957) dictated how tables were to be rearranged or compared to best illustrate the "natural" grouping of species into associations -- the fundamental vegetation unit. The three major premises of the Zurich-Montpelier School ideology are: (1) plant communities are best recognized by their f loral composition with equal weighting of the component species, (2) a subset of all species in a table have wil l have "diagnostic" properties that wil l consistently al low for the identification of a particular plant community, and (3) "diagnostic species" (character species, 67 SECTION ONE -- USING RELEVES TO STUDY MINNESOTA'S VEGETATION
differential species, or constant companions] can be used to create a hierarchal classification of the plant communities (Westhoff and Van Der Maarel 1978). The terminology of the releve method as it pertains to species, spacies groups, or groups of releves is linked to a particular table. That is, terms like character species, differential species, and constant companions are applied to species by calculating table-dependent statistics like presence, fidelity, and constancy. The hierarchal classes of releve groups are named according to formal rules of nomenclature and a structured taxonomy beginning with divisions as the largest units, fol lowed by classes, orders, allliances, and f inallly associations.
Today, most plant ecologists perceive plant communities as varying continuous1 y along complex environmental gradients (Gauch 1982, Whi ttaker 1967). This perception of plant communities is reflected in the common1 y used methods of analysis. This is particularly true of ordination and direct- gradient analysis, where the continuity of sample variation is obvious in the graphical ly displayed results. In spite of a shift away from the Zurich- Montpellier School's ideology and terminology, the phytosociological tab le -- the backbone of their method -- remains as particularl y useful way to display vegetation data. Grouping releves of similar species composition and grouping species with simi lar occurrence patterns is particular1y conduci ve to constructing hypotheses about the factors control ling plant distributions. Unlike ordination results, phytosociological tables retain al l of the sample information. It is good practice to create a phytosociological table to help interpret a particular ordination or classification by al lowing the ordination scores or classification groups to determine the order of the rows and columns of the table. Phytosociologi cal tables may be constructed di rectl y from programs where reciprocal averaging (correspondence analysis) is the primary algorithm for determining spacies or sample scores. Of such programs, TWINSPAN (Hill 1979) is the most consistent with the Braun-Blanquet method of producing a phytosociological table.
The Need For Community Data in a Natural Heritage Program
A major function of the Minnesota Natural Heritage Program is to maintain a database of "element occurrences" according to a set of guidelines prepared by The Nature Conservancy (1986). The term "element" may apply to any component of a natural ecosystem -- particularly plant and animal species, natural landscape features, and plant communities. The element concept is fairl y straightforward when applied to species because a physical specimen may serve as an element occurrence record and that specimen may be identified as a particular species (element) accordi ng to a documented taxonomic classification system. In contrast, the occurrence of a plant community element is difficult to record objectively because there is no agreement as to what constitutes an individual specimen (record), nor is there a universal ly accepted classification system. Plant communities lack a mechanism for inherent redundancy such as reproduction within the relative1y narrow confines of a species genome. In spite of the obvious problems of dealing with community elements, the MNHP is charged with the responsibility of (1) identifying community element occurrences, (2) classifying those occurrences, (3) determining the rel ative rarity of community classes, and (4) recommending management or protection policies for the rarer community types.
68 SECTION ONE -- USING RELEVES TO STUDY MINNESOTA'S VEGETATION
A state Natural Heritage Program may meet the above responsibilities on a short-term basis by constructing a vegetation classification system from the avai lable literature. One problem in doing so is that different researchers use different vegetation sampl ing methods, resulting in classification sy stems that are not general ly comparable. That is, it is often impossible to merge several datasets and reclassify the samples in an attempt to determine which community types are equivalent or different among the studies. A second problem is that researchers tend to focus on smal l areas, and the resulting classifications are not applicable on a state-wide scale, i.e. the community distinctions are too fine. Thus, a state-wide classification pieced together from several local studies results in too many community types, most of which are artificial ly rare.
An alternative to a literature-based classification is for Natural Heritage Programs to col lect their own vegetation data and then produce a community classification appropriate for their own needs. In most cases, this is prohibitively time consuming and expensive. In Minnesota, however, this is not the case. The consistency of instruction at the University of Minnesota has led several researchers to adopt the releve method of vegetation sampling. There are nearly 1,500 releves of comparable quality available to the MNHP (Almendinger 1985, Glaser 1983, Glaser et al. 1981, Glaser and Wheeler 1977, Janssen 1967; end unpubl ished data from the Minnesota Natural Heritage Program, Coper-Nickel Project. Kawi shiwi Map ping Project, The Nature Conservancy, Chippewa National Forest, Nerstrand Woods State Park, and other miscel laneous sources). Thus, much of the data necessary for a preliminary, statewide communi ty-classification has been collected -- but until now, there has been no serious attempt to compi le alll of this information.
Natural Communities as Pre-Defined Vegetation Units
A primary goal of the Minnesota Natural Heritage Program (MNHP) is to identify and conserve ecological communities that are rare on a state-wide scale. The first step in meeting the above goal was to devise a community classification system based upon ecological criteria that vary continuously across Minnesota. The Natural Community classification [Fig. 1.1, Wendt 1984) was the result of this first step, and the definitive ecological criteria include several habitat features -- vegetation, floristic province, topography, hydrology, land form, substrate, and soi ls. These criteria are by no means equally weighted or evaluated in a specified order. A natural community is subjectivel y named by its most characteristic feature -- biotic or abiotic -- such as Sioux Quartzite Outcrop, Calcareous Fen, Jack Pine Forest, or Gravel Prairie. Because vegetation integrates much of the variability of abiotic habitat features, Natural Community distinctions are strongl y correlated with vegetation.
The Natural Community classes were initial ly identified from the existing literature on the vegetation and natural areas of Minnesota. The classes are also intentional ly broad, so that the classification of any particular site is relatively unambiguous. The literature base and the broad categories make it easy for anyone fami liar with Minnesota's natural areas to envision the appearance of a site based upon its class designation. Thus, the classi f ication serves welll the need for communication among researchers. 69 SECTION ONE -- USING RELEVES TO STUDY MINNESOTA'S VEGETATION
teachers, students, naturalists, legislators, and others interested in the conservation of natural areas. The classification is less useful for other purposes and interests. For example, management plans for particular sites need more detailed information on the vegetation and abiotic features. The classification is also too coarse to demonstrate meaningful relationships between vegetation end environmental gradients. The lack of a true hierarchical structure makes classification revisions at any level (clustering or splitting) equal ly subjective.
Recognizing the need for more detailed descriptions of the vegetation and the need to classify the vegetation more objectively, the MNHP initiated the RELEVE project in 1986. With regard to the Natural Community classification, the MNHP plans to use releve data to (1) describe the variability of the vegetation within Natural Communities and (2) to establish a hierarchical relationship among Natural Community classes where the vegetation is the primary factor distinguishing the classes. To serve this need, releves col lected by the MNHP staff have been subjectively located within areas representative of a particular Natural Community class -- i.e. the Natural Community classes are the pre-determined units (see Step 1, Introduction) control ling the location of releves. The MNHP releve database also contains releves located by site (management projects) and by plant community (University of Minnesota studies). For any one using the MNHP releve database, it is critical that they understand the criteria used to initial ly locate the releves that make up the subset to be ordinated or classified. Extensive site information is kept on each releve so that researchers can evaluate the degree to which a subset of releves from the MNHP database meets their own criteria for analysis.
70 SECTION ONE -- USING RELNES TO STUDY MINNESOTA'S VEGETATION
NATURAL COMMUNITIES OF MINNESOTA Minnesota Natural Heritage Program
Format Communities Wetland Communities
Aspen - Birch Forast Calcareous Fan Black Spruca - Feather Moss Forast Conifer Swamp Floodplai n Forast Emergent Marsh Great Lakea Pine Forest Floating-Leaved Marsh Maple - Basswood Forast (Big Woods Section) Forested Bog Maple - Basswood Forast (Drifltless Section) Ha rdwood Swamp Maple - Basswood Forest (East Central Section) Opan Bog Maple - Basswood Forest (West Central Section) Patterned Peatland Complex Mixed Oak Forest Poor Fen Northern Hardwood - Conifer Forest Rich Fen Spruca - Fir Forest Sedge Meadow Upland White Cedar Forest Shrub Swamp Submergent Marsh
Savanna end Parkland Communities Primary Corunitiaa
Aspen Parkland Complex Algific Talus Slope Dry Sand Savanna, Jack Pine Subtype Bedrock Beach Dry Send Savanna, Oak Subtype Cobble Beech Mesi c Blacksoi l Sevenna Dry Cliff (Driftless Section) Dry Cliff (NE) Granite Outcrop (MNR Valley) Prairie Corunitiaa Moist Cliff (Driftless Section] Moist Cliff (NE) Bluff Prairie Sand Beach Dolomite Prairie Sioux Quartzite Outcrop Dry Send Prairie Glacial Till Hill Prairie Gravel Prairie Mesic Blacksoi l Prairie (EC) Mesi c Blacksoi l Prairie (NW) Mesic Blacksoil Prairie (NW) Calcareous Seepage Subtype Mesic Blacksoil Prairie (NW) Saline Subtype Mesic Blacksoil Preiria (SE) Mesic Blacksoil Prairie (SW) Mesic Blacksoil Prsi ris (SW) Sioux Quartzite Subtype Wet Blecksoil Prairie (EC) Wet Blecksoil Prairie (MI) Wet Blacksoil Prairie (SE) Wet Blecksoil Prairie (SW)
Figure 1.1 The Natural Communities of Minnesota (Wendt 1984), which serve as the pre-determined units controlling the distribution of releves taken by the Minnesota Natural Heritage Program staff. Releves are generally located within stands characterf stic of the above communities. Abstracts dascri bing the general appearance and composition of some of the above communities are available at the MNHP office. 71 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
LOCATING AND CONSTRUCTING THE RELEVE PLOT
Releve Location
Locating releves within sample stands is probably the most critical field decision to influence the results of a vegetation study. In a well-designed study, the landscape is initial ly segmented into study units by some predetermined criteria, eg. management units, forest cover-types, physiographic units, soil units. etc. Sample stands from within each study unit should be chosen by design (random, stratified-random, systematic, etc.) and then sampled by releve. If releve plots are not placed within areas of the sample stand that are representative of the study unit, then the purpose wi ll not be met. More specifically, a researcher would not be able to use the segmentation criteria or sampling design to draw inferences about the vegetation as it is perceived from the releve samples. Thus, the very first consideration that one should have when surveying a sample stand is to identify the portions of the stand that meet the criteria that define the study unit to which it belongs.
After acceptable portions of the sample stand are located, the surveyor should then consider the vegetation itself. Because the segmentation criteria (management units, cover types, soi ls. etc.) vary at a coarser scale than the vegetation, the vegetation wil l appear to vary within acceptable sampling areas. (If it did not, then a single releve would be sufficient to sample a stand.) One of the major purposes of sampling the stand is to express the variability of the vegetation within the study unit and the releves should be placed in a way to record that variability.
In the releve method, the process by which one surveys the vegetation of acceptable sample stand areas is cal led entitetion, i.e. visual ly dividing the vegetation into apparently homogeneous entities. The requirement of homogeneity applies to both the canopy and the understory vegetation. Obv iousl y, entitation invol ves the recognition of vegetational boundaries, and the releve plot should not be placed near boundaries. It it especially appropriate to record on a releve data sheet some impressions about the vegetational entities within a stand and the nature of the boundaries between them (diffuse, sharp, etc.). It is also appropriate to hypothesize as to what environmental factors are causing the apparent vegetational pattern. The convention for recording these observations on the MNHP Releve Data Sheet (see next subsection, Figures 2.1 and 2.3) is to record observations about the stand in general under the "Notes or Other Data Collected" section, and to record the position of the actual releve plot ralative to the entities and boundaries in the "Remarks" data-entry f ield.
The importance of entitation should not be underestimated. Remarks concerning entitation are field data, and as such, a review of thesa remarks is a valid means of rejecting sample stands or even reassessing the criteria by which the predetermined study units were defined. Most vegetation studies would be improved if the cycle of defining segmentation criteria and entitation was repeated several times before the first releve plot was described.
72 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELNE METHODS
Releve Size and Shape
A releve should be sufficiently large to include most species having a regular distribution within the sample stand. The appropriate releve size for a particular vegetation unit can be determined empirically by sampling nested plots and then plotting the number of species recorded versus plot size (Fig 2.1). For vegetation of temperate regions, the species/releve-area curve assymptoticall y approaches some maximum number of species as the releve size increases, i.e. the derivative approaches zero with increasing sample size. There are formal rules for using such curves to determine the "minimal sample area" for a vegetation unit (see Mueller-Dombois and Ellenberg 1974). In traditional European releve work, the "minimal sample area" is often considered a property of vegetation units.
In practice, the "minimal sample area" is sufficiently correlated with the structure and life-form of the vegetation so that it is not necessary to determine minimal releve sizes for each new study. Guidelines for releve sizes in communities of different physiognomy can be found in Mueller-Dombois and Ellenberg (1974), Westhoff and Van Der Maarel (1978), Benninghoff (1966), and Table 2.1 For the vegetation of Minnesota, a 400 square meter plot in forested communities and a 100 square meter plot in treeless communities safely exceed the minimal sample areas for those physiognomic units.
Table 2.1 Minimal area values for physiognomical ly distinct plant communities. Releve plot sizes should exceed the minimal area values. Compi led from Westhoff and Van Der Maarel (1978) and from Mueller-Dombois and Ellenberg (1974).
2 PLANT COMMUNITY SIMILAR MINNESOTA AREA m COMMUNITY
Tropical rain forest 200 - 1000 Temperate deciduous forest Maple-basswood forest 100 - 500 Dry grassland Upland prai rie 50 - 100 Scrub communities Oak barrens 25 - 100 Weed communities Old-field 25 - 100
The MNHP uses square releve plots--20 x 20 meters in forest and 10 x 10 meters in prairie or open wetlands. The shape of the plot and its orientation (if not a square or circle) are important only if it interacts with a regular or periodic vegetation pattern. For example, the "string bogs" of Minnesota consist of al ternating peat ridges and inundated troughs, topographically resembl ing the surface of a washboard. In some regions these ridges and troughs are sufficiently narrow so that a 10 x 10 meter square releve plot would always contain a portion of both trough and ridge. Under such ci rcumstances it migh be adv isable to use long-rectangular releve plots,
73 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
placing them entirely on a ridge or trough if the vegetation of those two features is to be contrasted or, alternatively, placing them transverse1 y across the ridges and troughs if the vegetation of the area as a whole is to be compared with some other type of bog vegetation. Releve shape might also be altered when sampling vegetation that varies with regard to slope or aspect. The specific purpose of a vegetation study should dictate releve shape.
SPECIES/A REA CUR VE
6 7 8 No. of plots
Figure 2.1 A nested-plot system used to determine minimal area, and a hypothetical species/area curve for the system. In general, a releve plot is considered to be sufficiently Large when doubling the sample area results in en increase of less then 10% in the number of species -- 32 m 2 om the above example. (After Muel ler-Dombois end Ellenberg 1974)
74 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
Mapping the Releve Site
After deciding upon the location, size, and shape of a releve plot. the location of the plot should be mapped while in the field. The Location should be plotted on e 7.5 minute U.S.G.S. topographic map and when possible, one should also plot the releve on the appropriate air photograph and township plat. A photocopy of the appropriate map sections should later be made and attached to the field form. This much effort in the field greatly expediates the transcription process end aids in relocating permanent releeve plots.
It is appropriate to sketch on the topographic map or on a separate piece of paper, unmapped features that would aid in relocating a releve site. Sketch maps showing trails, fencelines, buildings, clearcuts, etc. should be photocopied and attached to the Releve Data Sheet.
Constructing the Releve Plot
The final step to be completed before filling out the Releve Data Sheet is to lay out the actual releve plot. For most work, the plot boundaries and corners can be established by measuring along the perimeter of the plot and turning 90º at each corner. The 90º angles may be turned by using a field compass or by measur ing-in smal l, right t riang l es (30-60-90 or 45-45-90 triangles) in the releve corners. Releve plots (10 x 10 m) established in this manner general ly close to within 0.5 m of the point of origin (i.e. within 3 m2 of 100 m2).
THE WWHP RELEVE DATA SHEET
After the releve plot has been properly located and constructed, the actual releve data are recorded. This is accomplished by filling out both sides of the MNHP Reieve Data Sheet (Figures 2.2 and 2.3). Copies of the MNHP releve date sheet, suitable for photocopying. are avai lable from the MNHP office.
Site information is recorded on the "SITE INFORMATION" side of the data sheet, by fil ling in blenk date-entry fields (Figure 2.2) Some of these data- entry fields ere filled in while in the field end others are filled in while -in- the -office by transcribing site information from maps, lists, or tables.
The structure and species-composi tion data are entered on the "VEGETATION" side of the Releve Data Sheet (Fig. 2.3) whi le in the field. Structure data lines and species occurrence records are entered in the rows of a blank table with the appropriate. labeled columns. Some site information fields are repeated on this side of the Releve Data Sheet so that single-sided photocopy would contain sufficient site information to find the computer record of that releve in the MNHP database.
75 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
Recording Site Information
General information about the site, locational information, and specific information about the releve plot are recorded on the SITE INFORMATION side of MNHP Releve Data Sheet (Fig. 2.2). All of thee data-entry fields are of a specific length and of a specific type (character or numeric). The reason for this is that the data entered in these fields is eventually entered into the MNHP computer database as the values for variables of specific length and type. Much of the information entered in these fields is coded so that it can be more efficiently stored and retrieved on the computer. Code dictionaries end coding schemes are available from the MNHP office.
SITE INFORMATION VARIABLES
DNR -Releve Number, Transcribed variable. Each releve is assigned a four- -character number as it is entered into the MNHP releve database. The numbers are assigned by the HNHP data-manager. Numbers less than 1000 are to be prefixed with "0"s to fil l out the entry field, e.g. releve number 35 is entered as "0035."
Surveyor's Releve Number. Field variable. The surveyor should identify the releve plot with a number or code. This code should be created by the individual surveyors to distinguish the releve from others that they have col lected. The code can be any combination of alphabetic or numeric characters, in an eight-character field.
Surveyor's I.D. Code. Field variable. Surveyors should enter their identification codes in the three-character field. The code is general ly the surveyor's initials, however the surveyor should first check with the MNHP data manager to avoid duplicate coding of surveyors with the same initials. Surveyors without an assigned I.D. code should fil l in the parenthetical blank with their full name so that the MNHP data manager can make the code assignment.
Date: Field variable. The date of the month is entered in a two-character field. Because it is a character field, single-digit dates dates should be prefixed with a "0," e.g. 04 for the fourth day of the month. Otherwise "04" and "_4" and "4_" wil l not be interpreted by the computer as the same date.
Month. Field variable. The first three letters of the month are entered in the three-character field.
Year. Field variable. The full four-digit AD year is entered in the four- character field.
Site Name Code. Transcribed variable. The four-character code for site name is entered by the MNHP data manager.
76 77 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
DNR Ownership Code. Transcribed variable. The two-character code is entered -by the MNHP data manager. The dictionary of ownership codes is found in the MNHP Operations Manual. If the surveyor knows the land ownership, the owner's name should be entered in the parenthetical blank. When possible a township plat map showing the location of the releve should be attached to the Releve Data Sheet to aid in transcribing this variable.
Natural Community Code, Transcribed variable. The surveyor should enter the two-character code. The dictionary of codes for Minnesota Natural Communities is available from the MNHP office. If the surveyor is not certain as to which natural community to assign to the releve, a list of the possible community codes should be included in the parenthetical blank.
Natural Community Section, Transcribed variable. The surveyor should enter the two-character code. The natural community section codes and maps ere available from the MNHP office.
Natural Community Subtype Transcribed variable. The surveyor should enter the two-character coda. The natural community subtype codes are available from the MNHP office.
Element Occurrence Rankinq. Transcribed variable. The two-character element occurrence ranking is assigned by the MNHP data manager. The rules for determining the rank are described in detai l in the MNHP Operations Manual.
Element Occurrence Size. Field variable. The number of acres characterized by the releve are entered in the four-numeric field. The surveyor should review the entitation process in the "Releve Location" section above in order to make this estimate.
Site Size. Transcribed variable. The number of acres of the site is entered in the five-numeric field by the MNHP data manager. For small sites the acreage is often known with some precision (e.g. TNC properties, Scientific and Natural Areas, etc.). For very large sites like the Red Lake Peatlands or the Superior National Forest, site size should roughly correspond to the size of the forest cover-type within which the releve occurs.
State Field or Transcribed variable. The surveyor should enter the standard, two- letter, U.S. Postal Serv ice state coda in the two-character field.
County Code. Field or Transcribed variable. The surveyor should enter the number code for the Minnesota county in the two-character field. The dictionary of county codes is avai lable from the MNHP office.
DNR Quadrangle Code. Transcribed vari ab le. The four-character code is entered by the DNR data manager. A dictionary of DNR codes for USGS quadrangles in Minnesota is kept in the MNHP offices. To avoid transcription errors, it is good practice to enter the quadrangle name in the parenthetical blank while in the field.
78 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
Universal Quadrangle Code. Transcribed variable. The seven character code is entered by the MNHP data manager. A dictionary of Universal codes for quadrangles in Minnesota is kept in the MNHP offices.
Township. Field variable. The surveyor should enter the township number and the direction relative to a standard parallel in the four-character field. The number of the township is entered in the first three field blanks. with two-digit township numbers being prefixed with a "0," e.g. township 34 north would be entered as "034N." In Minnesota, all townships are north of the reference parallel, therefore, "N" will always be the fourth character of this field in Minnesota.
Range. Field variable. The surveyor should enter the renge number and the direction relative to a principal meridian in the three-character field. The number of the range is entered in the first two field blanks, with single- digit range numbers being prefixed with a "0," e.g. range 2 west would be entered as "02W."
Principal Meridian, Transcribed variable. The surveyor should enter the number of the principal meridian that references the range given above. It is a one-character field. In Minnesota, ranges are west of the 5th principal meridian, and they are either west or east of the 4th principal meridian. A map showing the regions of Minnesota that ere relative to the 4th and 5th principal meridians is available from the MNHP office.
Section. Field variable. The surveyor should locate the releve to quarter- quarter-section accuracy (approximately 40 acre cells). The two-character quarter-quarter-section f ield precedes that of the quarter-section f ield, and they are both filled in with the following codes: NE, northeast; NW northwest; SW, southwest; SE, southeast. If the releve Location is near a boundary and its quadrant location questionable, half-sections and half-quarter-sections may be indicated by the following codes: N_, north; S_, south; E_, eest; W_, west. The section number is entered in a two-character field, with single- digit section numbers prefixed with a "0," e.g. section 3 is entered as "03."
Latitude and Longitude, Transcribed variable. The surveyor should record the location of the releve by latitude and longitude by entering the degrees, minutes, and seconds in their respective two-character fields. A speci al map overlay for determining the latitude end longitude of a point on a 7.5 minute USGS topographic map is kept at the MNHP office.
-Releve -Size. Field variable. The releve size in square meters is entered in a three-numeric field. If a releve is larger then 999 square meters, 999 should be entered and the actual size of the releve indicated in the "Remarks" data-entry field. The convention is to right-justify areas Less than 100 square meters within the data-entry field.
Elevation, Field variable. The elevation of the releve site in feet is entered in the four-numeric field. The convention is to right-justify elevations Lass than 1000 feet within the data-entry field. The elevation may be estimated to the nearest contour elevation or the midpoint between two contour elevations.
79 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
Slope, Field variable. The degrees of slope end the downslope direction ere recorded in the four-character field. The degrees of slope are entered in the first two field blanks. Slopes of less then 10 degrees are prefixed by e "0," e.g. slope of 8 degrees southwest is recorded as "08SW." The two-character direction abbreviations are the same as the quarter-section end half- section abbrev iations (NE, NW, SW, SE, N_, S_, E_, W_ ).
Soi l Landscape Unit. Transcribed variable. The four-letter code for the Soil Landscape Unit in which the releve occurs is determined from the Minnesota Soil Atlas maps (1969, 1971, 1973, 1977, 1979, 1980 a,b,c, 1981 a,b). The codes as they appear on the map are entered in the four-character field. The Convention for codes having less than four characters is to left- justify them within the data-entry field.
Geomorphic Unit, Transcribed variab le. The alphanumeric codes for the Geomorphic- Unit in which the releve occurs is determined from the Minnesota Soil Atlas maps. The codes as they appear on the map are entered in the three character field. The convention for entering a code is to left-justify the two numbers and to use the rightmost column for letters when needed (e.g. 03_, 13_, 10A).
Remark. Field variable. Remarks pertaining specifically to the releve plot are recorded in the 200-character field. Any environmental factor that has possibly effected or that might effect the vegetation within the plot should be mentioned. A list of such factors would include some of the items listed below:
Soils: texture, color, aeration, parent material Geology: bedrock outcrops, landform Topography: hill crest, swale, foot of slope, midslope Disturbance: fire, Logging, burrows, disease, erosion Management: prescribed burn, selective cutting
The remarks field should also include deviations from standard releve procedures, for example, releves larger then 999 square meters. A brief description of other data collected might also be included.
Because the data-entry field is just 200 characters long, it is good practice to record potential remarks in the "Notes of Other Data Collected" section of the Releve Data Sheet. After some review and much condensation, the set of concise remarks can be fitted into the "Remarks" data entry field.
Other Data Collected. Field and transcribed variables. If soi ls, forestry, or water-chemistry-- date have been collected within the releve plot or sufficient1 y nearby as to be considered a characteristic of the releve plot, then that should be indicated by entering a "Y" in those one-character fields. Otherwise, enter an "N" in those fields. If such additional data exist for the releve, then the location of those data should be e priority comment in the "Remark" data-entry field .
The Plant Elements date-entry field should be filled with a "Y" if a plant designated as a MNHP element occurs in the releve. A list of plants with the element distinction is availabel from the MNHP office. 80 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
The Publication data-entry field cannot be entered in the field. It is to be filled with a "Y" if the releve has served as a sample in a published paper.
Recording Structure and Species Composition
The structure and species composition data-lines are entered on the rows of a blank table on the "VEGETATION" side of the Releve Data Sheet (Figure 2.3). A structure data line is a coded record of the col lective cover of a group of plants having the same life-form and also in a particular physiognomic height stratum. A species composition data l ine is a coded record of the cover (and some other information) for a plant species that is a component of a particular stratum. Every structural data line has associated with it a set of species composition data lines describing the occurrences of its component species in the releve plot.
Because releves record both stratum cover and species cover, and because species data other than cover are being recorded, it is impossible to design a simple fill-in-the-celltype of table for data entry. The data are multidimensional and only partly compatable, i.e. they cannot be merged into a si ngl e n-dimensional dataset. Entering mul tidimensional and non-compatable data in an essentially two-dimensional fashion on the Releve Data Sheet is very confusing to someone who has never col lected a releve. In practice. however, entering these data is not difficult -- if it is understood that the multidimensionality of the releve data is maintained by entering the releve data concurrently and in a specified order. The releve data entered concurrently in that data lines containing structural information are intermixed with data lines for species occurrences. The specific order of entering releve data is first by life-form, then by physiognomic height stratum, and finally by species. This confusing concept is best explained by example.
81 82 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELNE METHODS
Example--Entering Releve Data Blocks. Releve data are recorded in blocks of data lines belonging to a structural entity cal Led a physiognomic/life-form group. The data block consists of a single structure data-line end one to several species-composition data lines. The first data line of a block defines the physiognomic/life-form group by recording a code for the life- form, followed by a code for the physiognomic height class, followed by an estimate of the col lective cover of that group of plants. An example of a physiognomic/life-form data line is as follows:
E 4-6 c
In this example, "E" is the life-form code for needleleaf evergreen plants, "4-6" refers to the range of height classes [classes 4, 5, end 6) of the "E" plants, and "c" is the cover class code (75% to 100% cover] for for the "E 4-6" stratum. (See below and Appendix A for code descriptions.)
Obv iousl y, physiognomic/ life-form groups are composed of species. The data Lines for species occurrences immediately follow the data line for the physiognomic/life-form group of which they ere a part. Let's say that for the above example, the component species are white pine (Pinus strobus], balsam fir (Abies balsamea), and white spruce (Picea glauca). The entire entry for the "E 4-6 c" physiognomic/life-form group would take the following general form on a Releve Data Sheet:
SPECIES NAME OR CODE ** C.S REMARKS
E 4-6 c Pinus strobus 0 4.1 FR Abies balsamea 0 1.2 DF BR Picea cf. glauce 4 +.1 01
In this example, the codes following the species names (see below end Appendix A) describe the occurrence of a species in the releve plot as follows:
"**" refers to the column in which the codas for reliabi lity of identification of the species are entered. These codes (0,0, and 4 above) indicate the level of taxonomic certainty of the plant identification (family, genus, species, variety, etc.).
"C" refers to the column in which cover/abundance codes are entered. These codes (4,1, and + above) ere estimates of the species' absolute coverage or abundance within the releve plot.
"S" refers to the column in which sociability codes are entered. These codes (1,2, and 1 above] describe how the species is distributed within the releve plot (evenly, clumped, etc.)
83 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
"Remarks" refer to the column in which two-character remark codes are entered. These codes (FR, DF, BR, and 01 ) describe the vitality and condition of the species and also refer to miscel laneous remarks. Several remark codes may be entered on the Releve Data Sheet, but only two are entered into the RELEVE database. Thus, the codes should be entered in order of importance.
As illustrated above, the column headings of the Releve Data Sheet refer only to species-composition variables, which were entered by recording the appropriate code values for these variables in their respective columns. The structural data codes -- life-form, height stratum, and stratum cover -- are entered in the "SPECIES NAME OR CODE" column, and it is understood that the record of these structural variables applies to all of the species in the rows immediately following that entry. Blank rows are Left to separate the blocks data Lines for physiognomic/life-form groups. Refer to the sample records given in Figure 2.3 to clarify the entry of structural and species composition data by physiognomic/life-form groups.
Organization by Physioqnomic/life-Form Groups. From the above discussion and example, it should be clear that releve data are col lected and organized as blocks of records describi ng physiognomic/life-form groups. This structure is obvious in a completed releve form (Figure 2.3). Whi le organization by physiognomic/life-form group assures that the multidimensionality of the releve data is preserved, experience has shown that it causes the novice surveyor to be confused on two issues: (1) the factors that determine the number of physiognomic/life-form groups and (2) the number of data lines that can be entered for a single species.
The number of physiognomic/life-form groups recorded for a releve is determined by both the number of Life-forms and the number of distinct strata recognized for each Life form--in that order. For example, a releve with three distinct height strata and plants of four life-forms would NOT have three physiognomic/ Life-form groups composed of plants with different life forms. Rather, the releve would have minimum of four physiognomic/life-form groups because there are four Life-forms. The releve could have as many as twelve groups if representatives of all four life-form groups occurred in all three height strata. The height strata need not correspond between life-form groups as they do in this simple example. The estimates of height pertain to a particular l ife-form group only.
It is possible for a species to be listed several times if it occurs in different height strata, or if it exhibits different life-forms. For instance, releves in which sugar maple occurs almost always have three records of sugar maple -- one record in each of the height classes corresponding to tree, sapling, and seedling classes. Occasional ly, species exhibiting some plasticity with regard to life-form also may be doubly entered. An example would be woody species like poison ivy or woodbine which can grow either as lianas or as erect, deciduous shrubs.
84 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
STRUCTURE DATA LINE VARIABLES
Life-Form Codes. Structure Data Line. The life-form groups are represented by one-character codes (Table 2.2 and Appendix A). The codes are capitalized to distinguish them from the Lower-case codes for classes of stratum coverage (see below). The definitions and codes for life-forms follow Kuchler (1967). The MNHP uses Rosendahl (1975) as a guide to the life-form of woody plants in Minnesota.
Structure data-line example: B 1-2 b
In the above example the "B" is the Life-form code for woody plants with broad-leaf form and the evergreen habit. Note its distinction from the lower case "b," which is a cover code for the "B 1-2" stratum.
Table 2.2 Code definitions for life-forms of Minnesota plants. See Appendix A for detailed descriptions of the codes and examples of plants with particular life-forms.
Woody Plants Herbaceous Plants Speci al Life-Forms
B broedleef evergreen 6 graminoids C climbers D broadleaf deciduous H forbs K stem succulents E needleleaf evergreen L lichens, mosses X epiphytes N needleleaf deciduous
Height Class Codes. Structure Data Line. Height class codes ere numbers (I to 8) that refer to the heights of plants in a particular Life-form category. Each code refers to a range of heights (Table 2.3). These codes should be entered singly if the plants of that Life form fall within the range of heights defined by a code. Alternatively, a range of contiguous height classes may be given when plants of a certain life-form are not strongly stratified. The height class codes and definitions are follow Kuchler (1967).
Structure data-line example: B 1-2 b
In the above example the "1-2" defines the range of heights for broadleaf evergreen plants. When a renge is given, it is implied that the heights of the individual broadleaf evergreen plants ere evenly distributed between 0.0 cm (the Low range value for height class 1) and 50.0 cm (the high range value for height class 2). If, for example, there were just a few species of broadleaf evergreen plants in this releve, some with a creeping growth form (height class 1) and the others mostly 40 to 50 cm tall, then it would be appropriate to record the species as occurring in two, relatively distinct physiognomic/life-form classes, B 1 and B 2.
85 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
Table 2.3 Code definitions for physiognomic height classes. (After Kuch l e r 1967)
CLASS HEIGHT CODE
Coverage Classes. Structure Date Line. Coverage class codes are one- -character- codes that record the estimated cover of the entire physiognomic/ life-form stratum (Table 2.4). Cover is defined as the percent of the releve area covered by the vertical projection of leaf surface-areas. The cover class codes and definitions follow Kuchler (1967).
Structure data-line example: B 1-2 b
In the above example the "b" is the coverage class code for the entire "B 1-2" stratum. Note that coverage class codes ere entered in lower case as a means of distinguishing them f rom the upper case l ife-form codes.
Table 2.4 Code definitions for coverage classes of physiognomic/ life-form groups. See Appendix A for detailed descriptions of the classes. (After Kuch ler 1967)
CLASS COVERAGE DESCRIPTION
c = > 75% Continuous i = 50 - 75% Interrupted p = 25 - 50% Parklike r = 5 -25% Rare b = 1 - 5% Barely present a = < 1% Almost absent
86 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
SPECIES OCCURRENCE DATA-LINE VARIABLES
Species Names and Codes. Species Occurrence Data Line. The surveyor has the option to enter in the "SPECIES NAME OR CODE" column, either the Latin binomial for the plant or the MNHP eight-character code for the plant. The MNHP uses the Minnesota Checkl ist written by G. B. Ownbey, curator of the University of Minnesota Herbarium, as the authoritative guide to Minnesota's vascular plant names. The mnemonic codes were constructed directly from the Latin binomials in e way that minimizes the use of a code dictionary (Appendix B) and allows for construction of the codes at the date-entry step. The MNHP manages the Minnesota checklist, plant codes, and other species specific data in both manual and computer files. When recording the occurrence of nonvascular plants, the surveyor should enter the entire binomial and also cite the authoritative reference work that was used to identify those plants.
Species occurrence data-line: Pinus strobus 0 4.1 00 BR PINUSTRO 0 4.1 00 BR
In the above examples, the alternatives for referencing an occurrence of white pine are shown -- by its binomial "Pinus strobus," and by its mnemonic code "PINUSTRO." It is important to reference the variety or subspecies if it is known, because the mnemonic codes are constructed differently for taxa identified below the level of species.
Reliability of Identification Code. Species Occurrence Data Line. The codes for the reliability of identification are numbers that indicate the Level of taxonomic certainty at which plants have been identified (Table 2.5). It is important to recognize that plants cannot always be identified to the species level. When col lecting plot data, the surveyor must identify particular specimens, and particular specimens often lack key taxonomic characters as they ere influenced by f lowering-phenology, physical condition, life-cycle stages, and herbi vory. It is the surveyor's responsibi lity to express this uncertainty so thet others can evaluate the work. The MNHP uses these codes to make decisions about including species or releves into large datasets comprising the releves of several surveyors with varying degrees of taxonomic expertise. The codes were constructed by E.J. Cushing, University of Minn esota (personal communication, 1986).
Note that the computer MNHP system requires the entry of valid genus or valid binomial codes for plants-even if a plant has not been identified to that level of certainty. The MNHP system uses the codes for reliabi lity of identification to correctly express the uncertainy (by inserting "cf.") when writing the binomial names in reports.
Species occurrence date line: Pinus strobus 0 4.1 00 BR
In the above example, the "0" following the species binomial is the code for reliability of identification. The "0" indicates that the binomial is assigned without qualification, i.e. the surveyor is certain thet the species is white pine and there are no varieties or subspecies recognized in Minnesota.
87 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
Table 2.5 Code definitions for reliability of identification at the lowest taxonomic level. The identifications at higher taxonomic levels are presumed to be certain. See Appendix B for a detailed description of the codes. (After Cushing, unpublished)
CODE RELIABILITY OF IDENTIFICATION
variety or subspecies identified species identified, but variety or subspecies unknown species identified species complex identified species identification is uncertain genus identified genus identification is uncertain unknown taxon
Cover/Abundance Codes, Species Occurrence Data Line. The cove r/abundance codes are both numbers (1 to 5) and the characters "+" and "r" [Table 2.6 and Fig. 2.4). The numbers 2, 3, 4, and 5 refer to the cover classes for species with 5% to 100% cover in the releve plot. Species cover is defined as the percent of the releve covered by the vertical projection of the Leaf-area of that species as it occurs in a particular physiognomic/ life-form stratum. Codes 1, +, and r refer to qualitative estimates of the abundance [number) of plants of a species with less than 5% cover in that physiognomic/life-form stratum. See Mueller-Dombois and Ellenberg (1974) for discussion of the Braun-Blanquet cover/abundance scal e and simi lar scal es.
Species occurrence data Line: Pinus strobus 0 4.1 00 BR
In the above example, the "4" is the cover/abundance code for white pine as it accounts for 75% to 100% of the cover in a particular physiognomic/ life- form stratum. Note that the cover/abundance code is always separated from the sociability code, "1," (see below) by a period, ".".
88 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
Table 2.8 Code definitions for the Braun-Blanquet cove r/abundance scale. See Appendix B for a detailed description of the codes and scale (After Mueller-Dombois and EL Lenbarg 1974)
CODE CWER TYPE
5 = > 75% Cover 4 = 50 - 75% Cover 3 = 25 - 50% Cover 2 = 5 - 25% Cover COVER ABUNDANCE 1 = < 5% Cover + = < 5% Abundance r = < 5% Abundance
Figure 2.4 Example of cover/abundance values as described in Table 2.6.
Sociabi l i ty Code. Species Occurrence Data Line, Soci abi l ity codes describe how a species is distributed within the releve plot (Table 2.7). These numeric codes refer only to the distribution of a species as it occurrs in a particular physiognomic/life-form stratum. For example, it often happens that the distribution of a tree species is uniform within the tree stratum and clumped within the seedling stratum.
Species occurrence data Line: Pinus strobus 0 4.1 00 BR
In the above example, the "1" is the sociabl i ty code, which indicates that the white pines are distributed uniformly across the releve for a particular physiognomic / l ife-form stratum.
Table 2.7 Code definitions for sociability. See Appendix B for a detai led description of the sociability codes. [After Mueller- Dombois and Ellenberg 1974)
CODE DESCRIPTION
5 = Extensive mat 4 = small colonies, broken mat 3 = large group. many plants 2 = grouped, few plants 1 = growing singly
89 SECTION TWO -- THE MINNESOTA NATURAL HERITAGE PROGRAM RELEVE METHODS
Remark Codes, Species Occurrence Data Line. Remark codes are two-charecter codes that indicate some special attribute of the species as it occurs in a particular physiognomic/life-form stratum (see Releve Code Sheet in back pocket). Some remark codes refer to the species' vitality, i.e. they are qualitative estimates of the ability of the species to perpetuate itself. Other remark codes refer to the condition of the species, which varies as a result of both inherent forces (seasonal phenology, life-cycle, etc.) and external forces [herbi vory, windthrow, fire, etc.). Mi scellaneous remarks, which do not refer to vitality or condition, may be created to suit particular vegetation studies.
Species occurrence data Line: Pinus strobus 0 4.1 00 DF
In the above example, the "00" and the "BR" are remark codes. "00" is a vitality code that indicates that the white pines are of poor vitality. "BR" is a condition code that indicates that the white pines have been defoliated. When vitality codes are given, it is good practice to express a condition code that helps explain the vitality condition or, alternatively, to include an explanatory note in the "REMARK" or "NOTES OR OTHER DATA COLLECTED" fields on the SITE INFORMATION side of the releve data sheet. For this example it would be appropriate to explain that the white pines are of poor vitality (00) because they have been defoliated (DF) by white pine blister rust.
90 REFERENCES
Almendingar, J.C. 1985. The Late-Holocene development of jack pine forests on outwash plains, north-central Minnesota. Ph.D. Dissertation. University of Minnestoa, Minneapolis, Minnesota, USA.
Becking, R.W. 1957. The Zur ich-Montpel lier School of phytosocio l ogy. Botanical Rev i ew 23:411-488.
Benninghoff, W.S. 1966. The releve method for describing vegetation. The Mi chi gan Botanist 5:109-114.
Braun-Blanquet, J. 1928. Pf Lanzensoz iol ogi a, Grundzuge der Vegetationskunde. Springer-Verlag. Berlin.
Braun-Blanquet, J. 1932. Plant sociology: the study of plant communities. [Translation of Pf Lanzensoziologie, C.D. Ful ler and H.S. Conrad, editors] McGraw-Hill Book Company, New York, New York, USA,
Gauch, H.G. 1982. Multi variate analysis in community ecology. Cambridge University Press. 298 pp.
Glaser, P.H. 1983. Vegetation patterns in the North Black River peatland, northern Minnesota. Canadian Journal of Botany 61:2085-2104.
Glaser, P.H. and G.A. Wheeler. 1977. Appendix B. Vegetation. In Terrestrial Vegetation and Wildli fe Supplement, Envi ronmental Impact Statement, Minnesota Power and Light Company, Proposed Unit 4, Clay Boswel l Steam Electric Station, Minnesota Pol lution Control Agency, St. Paul, Minnesota, USA.
Glaser, P.H.. G.A. Wheeler. E. Gorham. and H.E. Wright, Jr. 1981. The patterned mires of the Red Lake peetland, northern Minnesota: Vegetation, water chemistry and landf orms. Journal of Ecology 69:575-599.
Hi l l, M.O. 1979. TWINSPAN - A FORTRAN program for arranging multi vari ate data in an ordered two-way table by classification of the individuals and attributes. Cornel l University, Ithaca, New York, USA.
Janssen, C.R. 1967. A floristic study of forests and bog vegetation, northwestern Minnesota. Ecology 48:751-765.
Kuchler, A.W. 1967. Vegetation mapping. The Ronald Press Company, New York, New York, USA.
Mueller-Dombois, D. and H. Ellenberg. 1974. Aims and methods of vegetation ecology. John Wiley and Sons, New York, New York, USA. 547 pp.
Minnesota Soi l Atlas. 1969. 1971, 1973. 1977, 1979. 1980 a,b,c, and 1981 a,b. A series of maps [scale 1:250,000) and Miscellaneous Reports, published by the Agricultural Experiment Station, University of Minnesota, St. Paul, Minnesota. USA.
91 REFERENCES
Rosendahl, C.O. 1975. Trees and shrubs of the upper Midwest. University of Minnesota Press, Mi nneapol i s, Minnesota. USA.
Mueller-Dombois, D. and H. Ellenberg. 1974. Aims and methods of vegetation ecology. John Wiley and Sons, New York, New York. USA 547 pp.
The Nature Conservancy. 1988. Natural heritage program: model operations manual. The Nature Conservancy, Arl ington, Virginia, USA
Wendt, K.M. 1984. A preliminary classification and description of natural communities in Minnesota. The Minnesota Natural Heritage Program, Department of Natural Resources, St. Paul, Minnesota, USA. 36 pp.
Westhoff, V. and E. van der Maarel. 1978. The Braun-Blanquet approach. pp. 287-399 Classification of plant communities, R.H. Whittaker. editor. The Hague: Junk.
Whittaker, R.H. 1967. Gradient analysis of vegetation. Biological Reviews 42:207-264.
92 APPENDIX A - STRUCTURE DATA LINES
CODE DEFINITIONS FOR LIFE-FORM After Kuchler (1967)
WOODY PLANTS
B: Broadleaf Everqreen. This group of woody plants has broad leaves (as distinguished from needle-li ke leaves) that persist for two to severa l years. In Minnesota, this group is most often represented by members of the Eri caceae (Andromeda, Arctostaphylos, Chamaedaphne, Epi gea, Gaul theria, and the cranberries of Vaccinium). The MNHP also includes suffruticose evergreen plants in this group (e.g. Py rola, Chimaphi la, and some of the above Ericeceous genera).
D: Broadleaf Deciduous, This group of woody plants have broad leaves as in "B" above, but these leaves are either shed or dead (nonphotosynthetic) during some part of the year. In Minnesota, this group encompases e very large number of tree and shrub genera (e.g. Acar, Betula, Corylus, Fraxinus. Quercus, Ulmus, etc.).
E: Needleleaf Evergreen. This group of woody plants includes both needle-leaved and scale-leaved evergreens. In Minnesota these plants are gymnosperms. Need le-leaved genera are: Abies, Picea, Pinus, Taxus, Tsuga. Juniperus and Thurja are examples of scale-leaved evergreens.
F: Needleleaf Deciduous. This group of woody plants has needle-like leaves as in "E" above, however these leaves are shed every year. The only Minnesota species having this life-form is Larix Laricina.
HERBACEOUS PLANTS
G: Graminoids. This group of herbaceous plants includes alll plants that appear grasslike because of their long, linear Leaves and unbranched form. In Minnesota alll members of the fol lowing fami lies are considered to be graminoids: Cypereceae, Gramineae, Juncaceae, and Typhaceae.
H: Forbs. This group of herbaceous plants have broad Leaves as opposed to the graminoids [above] and are not lichens or mosses [below]. In Minnesota, this group is represented by a very large number of angiosperm families as well as the ferns and fern allies.
L: Lichens and Mosses. This group includes all lichens and mosses that grow on the ground (rock or soil). In Minnesota the Cladonia lichens and the moss genera. Brachythecium, Hylocomnium, Mnium, Pleurozium, Polytrichum, and Pti l ium are examples of ground-covering taxa. Epiphytic mosses and Lichens are included within the epiphyte special life form.
93 APPENDIX A - STRUCTURE DATA LINES
LIFE-FORM CODES cont...
SPECIAL LIFE FORMS
C: Climbers. This group includes al l woody plants that are rooted in the ground and climb objects or other plants. In Minnesota, this group is most often represented by Rhus, Parthenocissus, and Vitis, Herbaceous climbers Like Convolvulus, Cuscuta, Oioscorea, Lathyrus, Vicia, etc. are included in the "H" group.
K: Stem Succulents. In Minnesota, this group includes only the native cacti, Coryphantha, and Opuntia. Plants with fleshy eaves, Like Sedum, are included within the "H" group.
X: Epiphytes, Epiphytes includes a wide variety of plants with different growth forms, therefore plants in this group are not all similar in appearance. The HNHP convention is to include all plants that live on the above-ground parts of other plants in this group. This group includes all epiphytic mosses, lichens, and higher plants Like Arcauthobium. Parasitic plants that are apparently rooted in the soil ( Monotropa, Orobanche, etc.) are included within the "H" group.
CODE DEFINITIONS FOR HEIGHT CLASSES After Kuchler (1967)
CLASS HEIGHT CODE
94 APPENDIX A - STRUCTURE DATA LINES
CODE DEFINITIONS FOR COVERAGE CLASSES After Kuchler (1967)
CLASS COVERAGE EXPLANATION AND EXAMPLES
c = > 75% Continuous. Implies that the cover is distributed evenly across the releve. In general, plant canopies wi ll touch and the coverage will exceed 75%. For some sparsely- leaved species (needleleaf evergreens, graminoids, etc.), the plant canopies wi ll touch, but not account for 75% coverage--these are still designated as having continuous cover.
i = 50 - 75% Interrupted. General ly assigned (1) to strata with a "hole" in an otherwise continuous coverage, (2) to strata in which plant canopies do not touch. and (3) to strongly clumped herbaceous or grami noid species when the clump canopies don't touch.
p = 25 - 50% Parklike or in Patches. This code is most often assigned to shrub strata where the shrobs occur in patches or to patchy colonies of herbaceous plants.
r = 5 - 25% Rare. Applies to strata in which the plants are more widely scattered than in "p." Often, the distinction between "p" and "r" is to separate strata composed primarily of plants with vegetatively reproducing col onies f rom strata composed of plants reproducing by Long rhizomes or seeds.
b = 1 - 5% Barren or Barely Present. Generally assigned to strata with widely scattered plants that have fai rly large leaf-areas (eg. bracken fern).
a = < 1% Almost Absent. General ly assigned to strata with widely scattered plants that have small leaf-areas (eg. grami noids, conifer seedli ngs, etc.)
95 APPENDIX B - SPECIES-OCCURENCE DATA LINES
MNEMONIC CODES FOR MINNESOTA PLANTS Codes assigned by J.C. Almendi nger
Mnemonic codes were assigned to all of the vascular plants listed in Ownbey's (in press) Checklist of the Vascular Plants of Minnesota. All of the appropriate codes can be assigned by using Ownbey's checklist and the rules given below, with only a few exceptions. The exceptions are cases of duplicate codes for different plants, and a listing of these plants is given below. The MNHP maintains a computer dataset of Ownbey's checklist of species along with their codes, authors, hybrid crosses, end other species specific data. A Listing of this datafile is available upon request. The codes are eight characters long and were assigned according to the fol lowing rules:
GENERA
The codes for genera are the first eight characters of the genus name.
Example: MUHLENBE for MUHLENBErqia
SPECIES WITH N0 RECOGNIZED VARIETIES OR SUBSPECIES IN MINNESOTA
The code is constructed by concatenating the first four letters of the genus name with the first four letters of the species name.
Example: LITHCANE for LITHospermum CANEscens
SPECIES WITH RECOGNIZED VARIETIES IN MINNESOTA
The code is constructed by concatenating the first four letters of the genus name, with "VA" for variety, end with the first two letters of the variety name.
Example: LATHVAIN for LATHy rus venosus VAr. INtonsus
SPECIES WITH RECOGNIZED SUBSPECIES IN MINNESOTA
The code is constructed by concatenating the first four letters of the genus name, with "SS" for subspecies, and with the first two letters of the variety name.
Example: LITHSSCR for LITHospermum ceroliniense SSp, CRoceum
96 APPENDIX B - SPECIES-OCCURRENCE DATA LINES
HYBRID SPECIES
The code is constructed by concatenating the first four letters of the genus name, with "XX" for hybrid, and the fi rst two letters of the species name for the hybrid.
Example: LYCOXXBU for LYCOpodium X BUttersii (L. Lucidulum X L. selago)
SPECIES WITH SHORT NAMES
Occasionally a species or genus name wil l be too short to form the code according to the above rules. In such cases, the code is formed by leav ing blanks within the concatenation.
Example: POA WOLF for Poa wolfii ACER for
DUPLICATE CODES
Constructing the codes according to the above rules resulted in some duplicate codes for different plants. The codes were made different by assigning numbers as the last character.
97 APPENDIX B - SPECIES-OCCURRENCE DATA LINES
CODE DEFINITIONS FOR RELIABILITY OF IDENTIFICATION Codes assigned by E.J. Cushing
CODE RELIABILITY OF IDENTIFICATION
0 = Binomial assigned without qualification. This includes plants identified to species level when (1) varieties or subspecies are not distinguished in Minnesota collections and (2) when the typical variety is understood.
Example: Baptisia bracteata var. glabrescens Aster cordifolius ssp. sagittifolius -Pinus - strobus [var. tvpica understood] 1 = The species identification is confident, but the variety or subspecies identification is in doubt.
Example: Baptisia bracteata cf. var. glabrescens
2 = The species identification is confident, but the variety or subspecies is not distinguished--even though varieties or subspecies occur in Minnesota.
Example: Baptisia bracteata
3 =Species identification is trivial because of hybridization among several recognized species, but hybrid complexes are recognized within the genus.
Example: Amelanchier interior complex
4 = The genus identification is confident, but the species identification is in doubt.
Example: Baptisia cf. bracteata
5 = The genus identification is confident, but the species is not distinguished.
Example: Bapti sia
6 = The genus identification is in doubt.
Example: cf. Baptisia
7 = The plant is unknown. but only one species is probably included. Plants recorded as unknown should be col lected and the col lection number given in one of the remark fields in the species occurrence data line. 98 APPENDIX B - SPECIES-OCCURRENCE DATA LINES
BRAUN-BLANQUET COVER/ABUNDANCE SCALE After Muelle r-Domboi s and Ellenberg (1974)
CODE COVER EXPLANATION
5 = > 75% A cover code. Assigned to a species occurrence within a particular stratum where that species' cover is more then 75% of the releve area.
4 = 50 - 75% A cover code. Assigned to a species occurrence within a particular stratum where that species' cover is between 50 and 75% of the releve area.
3 = 25 - 50% A cover code. Assigned to a species occurrence within a particular stratum where that species's cover is between 25 and 50% of the releve area.
2 = 5 - 25% A cover code. Assigned to a species occurrence within a particular stratum where that species' cover is between 5 and 25% of the releve area.
1 = < 5% An abundance coda. Assigned to a species occurrence within a particular stratum where there ere numerous individuals of a species, but those individuals collecti vely cover less than 5% of the releve area.
+ = < 5% An abundance code. Assigned to a species occurrence within a particular stratum where only a few (appx. 2 - 20) individuals occur and col lectively cover Less than 5% of the releve area.
r = < 5% An abundance code. Assigned to a species with only a single individual occurring within a particular stratum and also to species occurring just out of the plot (Remark "OP").
99 APPENDIX B - SPECIES-OCCURRENCE DATA LINES
CODE DEFINITIONS FOR SOCIABILITY After Mueller-Dombois and Ellenberg (1974)
CODE EXPLANATION
5 = Assigned to species occurrences where the plants ere growing in large, essential ly monotypic stands that form an extensive mat. Typically this is applied to non-woody plants, eg. moss or Lichen carpets, graminoid sods, etc.
4 = Assigned to species occurrences where the plants are growing in small colonies or broken mats. Typically this is applied to non-woody plants, eg. broken carpets as described above, and also colonies of herbaceous plants that have enlarged to the point that they are beginning to coalesce.
3 = Assigned to species occurrences where the plants are growing in small patches or as cushions. Typically the small patches include small, isolated clones of herbeceous plants as well as patches of shrubs. Cushions generally refer to moss or Lichen colonies.
2 = Assigned to species occurrences where the plants form small, often dense, clumps. These small clumps may be rather evenly dispersed within the releve. This often applies to woody or herbeceous plants where several aerial stems originate from the rootstock of a single genet.
1 = Assigned to species growing solitarily. This applies to both woody and herbaceous plants where single stems appear to be evenly dispersed within the releve.
100 Chapter 6
The Electroretinogram of the Horseshoe Crab, Limulus polyphemus: A Laboratory Exercise in Sensory Physiology
Robert A. Linsenmeier 1 , Charles M. Yancey 2 , and Wesley W. Ebert 3
1 Department of Neurobiology and Physiology, and Department of Biomedical Engineering, 2 Interdepartmental Graduate Program in Neuroscience, and 3 Section of Biological Sciences Northwestern University O.T. Hogan Hall 2153 Sheridan Road Evanston, IL 60208
Robert Linsenmeier is an Assistant Professor in the departments of Neurobiology and Physiology, and Biomedical Engineering at Northwestern University. He received his B.S. in Chemical Engineering from Carnegie-Mellon and his M.S. and Ph.D. (1978) from Northwestern. After spending four years as an Assistant Research Physiologist at the University of California-San Francisco, he returned to Northwestern, where he teaches an animal physiology course and a biomedical engineering laboratory course. His research interests are in the neural circuitry of the retina and the microenvironment of the retina.
Charles M. Yancey received his B.A. and M.S. degrees from Northwestern University in 1983 in biology with an emphasis in sensory physiology. He is currently a fellow in Northwestern's combined M.D./Ph. D. program and has recently completed his Ph.D. dissertation. He has been a laboratory teaching assistant in animal physiology and medical neuroscience. His research interests are focused on the role of altered retinal oxygenation and metabolism in ocular disease. Wesley W. Ebert received his B. S. degree from the University of California-Davis and his M.S. degree from the University of Wisconsin-Madison. He is currently a Lecturer at Northwestern University in the Section of Biological Sciences, with responsibility for coordination of the Ecology and Evolutionary Biology Department's teaching labs. His earlier responsibilities included operation of the teaching labs for the Neurobiology and Physiology Department.
101
THE ELECTRORETINOGRAM OF THE HORSESHOE CRAB, Limulus polyphemus:
A LABORATORY EXERCISE IN SENSORY PHYSIOLOGY
INTRODUCTION
Over the past several years, we have developed two versions of a laboratory exercise in which we record the electroretinogram (ERG) from the large, lateral, compound eye of Limulus polyphemus, the horseshoe crab. The simpler version is for undergraduates (mostly juniors) taking their first course in Neurobiology and Physiology. It is designed to be completed in one four-hour laboratory period. Students in this course are also expected to read relevant sections of the textbook, Animal Physiology (Second Edition) by R. Eckert and D. Randall (W. H. Freeman and Company; San Francisco, CA 1983). The advanced version is for juniors, seniors, and graduate students in Biomedical Engineering, and takes two four-hour laboratory periods. The simpler version, as given to the students, is presented here as an Appendix. Ideas from the advanced version that may be of interest to instructors and preparators are included in the following sections.
The objectives of both exercises are 1) to give the students some technical experience in working with an animal preparation and in making electrophys- iological recordings; 2) to enable the students to leave the laboratory with a set of data which they can analyze; and, hopefully, 3) to teach some principles of sensory physiology. While the system under study is vision, many of the concepts apply to other sensory systems as well.
One of the great advantages of this experiment is the variety of investigations that can be done after a relatively simple preparation. All students are expected to study the dependence of response amplitude on the intensity and duration of the stimulus, the variation of response latency with stimulus intensity, and the time course of dark adaptation following a prolonged period of illumination. Many students also investigate the shift in visual sensitivity that occurs during steady illumination and the effect on response amplitude of trading off stimulus duration for intensity. The data lend themselves to quantitative analysis, particularly linear regression and determination of time constants, but these are not stressed in our simpler version.
We are aware of only two undergraduate laboratory exercises that deal with visual physiology in an animal preparation. These are an exercise on the frog electroretinogram in Experimental Neurobiology by Oakley and Schafer (1978). and one on the visual system of the horseshoe crab in Twenty-six Afternoons of Biology by Wald et al. (1968). A slightly modified version of the Wald experiment has also been published by Packer (1967) in Experiments in Cell Physiology.
We feel that it is advantageous to expose students to a wide range of animal preparations, particularly when one can use an animal as interesting as the horseshoe crab. Direct conclusions about the mammalian visual system have been drawn from research on horseshoe crabs despite the enormous evolutionary gap, and students should appreciate the similarities. We also suspect that the horseshoe crab preparation is hardier than that of the frog. Furthermore, the
103 100 msec
Figure 1. Limulus ERG in response to 100 msec flashes. Log attenuation of the stimulus (number of 1.0 log unit neutral density filters in the light path) is shown next to each trace. (For this experiment, 0 log attenuation was approximately 4 mW/cm2.) These responses were recorded on FM tape, digitized by a PDP 11/23 computer, and plotted.
Studies of the ERG of the horseshoe crab apparently began with the work of H. K. Hartline (1927). who eventually pursued work on this visual system at the level of single-cell recordings. In 1964, Hartline won the Nobel prize in Physiology and Medicine for his work on visual systems, an important part of which involved Limulus polyphemus. Hartline showed that the ERG could be recorded from many kinds of arthropods, but that the ERG of the horseshoe crab was one of the simplest - considerably simpler, in fact, than the insect ERG, which may have several components. Little quantitative work has been done on the ERG of the horseshoe crab, since one can address questions of visual physiology more directly in this animal by recording from the optic nerve or from photoreceptors. Two studies on the ERG are of interest, however, and these will lead to other literature.
Chapman and Lall (1967) measured the spectral sensitivity of the ERG recorded from the lateral eye. They suggested that only a single type of receptor, with a peak sensitivity at 525 nm, as present. Their article includes some response waveforms and intensity-response functions. More recently, Barlow and coworkers have used the ERG to monitor several circadian changes in the structure and function of the lateral eye (e. g., Barlow 1984). They show that the sensitivity of the eye is at least a factor of ten greater at night than during the day, and that the response is graded over four to five log units of illumination. If the optic nerve is cut, the rhythms disappear, indicating that efferent fibers to the eye carry the circadian information.
A great deal of information relevant to this exercise can also be obtained from studies of action potentials recorded from individual axons of the optic
104 ERG of the horseshoe crab is a very simple wave, not a complicated set of potentials like the frog ERG. Thus, there is no question about what to measure as the response amplitude. In addition, the horseshoe crab ERG is generated primarily by the photoreceptor cells themselves (Barlow, 1984). It is much more difficult to explain the origin of the various components of the vertebrate ERG, some of which are probably generated by glial cells and retinal pigment epithelial cells (e.g., Armington, 1974).
Wald's exercise on the horseshoe crab gives a good starting point, but most faculty would find it difficult to implement since the write-up is really only an outline. Our intent is to give a more complete description than Wald et al. provide so that one can teach the laboratory without being a specialist in visual physiology.
BACKGROUND
Limulus polyphemus has impressive visual capabilities. Under dark-adapted conditions, one photon of light can lead to an action potential in an optic nerve fiber, so that its sensitivity is at least capable of matching that of humans (Kaplan and Barlow, 1976). The horseshoe crab has ultraviolet receptors in the median ocelli (Chapman and Lall, 1967), and it has at least six differ- ent photosensitive systems: 1) the large, lateral, compound eyes, 2) the pair of median ocelli at the front of the carapace, 3) ventral photoreceptors near the front legs which are not organized into an eye, 4) photoreceptors in the telson (tail) that also act as mechanoreceptors and can shift the phase of the circadian clock (Hanna, Horne, Renninger, Kaplan, and Barlow, 1985). 5) rudimentary lateral eyes that can be seen under the cuticle just behind and medial to the lateral compound eyes as masses of white connective tissue containing photoreceptors, and 6) an endoparietal eye, similar to the rudimen- tary lateral eyes in structure and just rostral to the median ocelli (Millec- chia, Bradbury, and Mauro. 1966). What the horseshoe crab uses all of these eyes for is not entirely clear. They are probably not used for feeding, but vision does seem important in mating (Barlow, Powers, and Kass, 1987).
--The ERG The electroretinogram is a potential that results from currents generated by retinal cells and conducted to the surface of the eye. It bears the same relation to the neurons of the retina that the electrocardiogram has to the electrical activity of cardiac muscle. In humans, the ERG is recorded clinically between an electrode embedded in a contact lens and a reference electrode at another point on the head. In animals, it can be recorded in the same way, but one electrode is often placed inside the eye in order to have recordings that are larger in amplitude and more stable. In the experiment presented here, one electrode is placed in contact with the cornea and a reference electrode is placed in sea water underneath the isolated eye. As shown in Figure 1, the horseshoe crab has a corneal negative ERG. This is because depolarization of the Limulus photoreceptors during illumination causes a sodium current to flow inward at the distal (top) portion of the cell. Extracellular current then flows from the proximal to the distal end of the cell, and the extracellular voltage at the distal end of the cell, which is nearest the corneal electrode, is therefore more negative than the voltage at the reference electrode.
105 nerve of the horseshoe crab. An introductory article on this subject is by Miller, Ratliff, and Hartline (1961). Hartline and McDonald (1947) wrote a very useful article on the time course of dark adaptation following different conditions of light adaptation and on sensitivity adjustments during light adaptation. In the 1970s and 1980s. the horseshoe crab has continued to be an important animal in vision research. Most of the very recent articles are cited by Barlow, Kaplan, Renninger, and Saito (1987).
MATERIALS REQUIRED
Figure 2 illustrates the set-up that we use at Northwestern University for the simpler version of the exercise; it can be readily modified to fit individ- ual needs. The only piece of apparatus missing from this diagram is the Physioscribe, a chart-recorder described below. Figure 3 is a schematic showing the equipment used in the more advanced version of the exercise. The materials required for the simpler version are listed below. The actual preparation is described in the Appendix.
plexiglas stage leads to Physioscribe hotoelectric 1. channel aluminum 2. Transducer channel 3. ground (to Physioscribe case or outlet)
Figure 2. Basic set-up for Limulus ERG experiment.
106 SQUAREWAVE Prepulse Sync (Trigger) I STlMULATOR Grass SD9 Stimulus Yaveform
Output
LIGHT
SHUTTER DRIVE NEUTRAL DENSITY PHYSIOSCRIBE STORAGE UNIT FILTERS OSCILLOSCOPE (Optional ) Uniblitz 100-28 Hitachi V-134 (Vincent Assoc) , ELECTRONIC ERG SHUTTER A trigger Uniblitz 26L
t Light-proof PREAMPLIFIER shielded PREPARATION boa , I Grass P15 or Channel 1
Figure 3. Schematic for advanced version of the Limulus ERG experiment.
Electrophysiological equipment
1. Physioscribe with Time and Event channel. Biopotential channel, and Transducer channel; three pens with ink bottles connected to the three channels; chart paper for Physioscribe.
2. Wooden, light-proof box with no bottom and a front made by stapling copper screen between two pieces of black cloth. The front is stapled to the top edge of the box so that it opens upward. The interior of the box is painted black and lined with copper screen to which a grounding wire is attached. There should be a hole in the top of the box to hold the shutter and a closeable hole (black cloth attached by Velcro) in one side for the light-adaptation experiments.
3. Aluminum plate (half-inch thick) with several holes drilled to receive banana plugs of grounding wires; fits beneath box.
4. Camera shutter with cable release mounted in hole on top of box.
5. Neutral density filters: three 1.0 log unit filters and one 0.5 log unit filter. These are stacked on top of shutter in various combinations.
6. Two dissecting microscope illuminators; one mounted on top of box with light directed down through shutter into interior; the other directed through the side hole during the light-adaptation experiment. (Light sources other than microscope illuminators could be used as well.)
7. Plexiglas stage with two banana jacks. The eye preparation sits in a
107 petri dish on top of the stage. Two Ag/AgCl electrodes lead from the banana jacks to the preparation; the jacks are connected to the Biopotential channel of the Physioscribe.
8. Photoelectric transducer. This is placed below the Plexiglas stage and connected to the Transducer channel of the Physioscribe.
9. Grounding lead from aluminum plate to case of Physioscribe or power outlet.
Equipment for dissection and mounting of eye
1. 125 ml Erlenmeyer flask containing approximately 75 ml seawater from aquarium; Pasteur pipette with bulb.
2. 1 ml plastic hypodermic syringe with 25G 5/8" needle.
3. Scalpel, forceps, razor blade.
4. Bottom of small (60 x 15 mm) glass petri dish.
5. Lump of plasticene (modeling clay) - approximately 1 cm diameter.
6. Silicone stopcock grease and small spatula.
7. Binocular dissecting microscope with illuminator.
Animal s
1. Limulus polyphemus (2"-4" carapace width), secured to a board with T- pins; one animal per pair of students. The second eye may be used later by another pair.
2. Conditioned marine aquarium; equipped for refrigeration, aeration, and filtration. Alternatively, animals could be kept for at least a few days in unfiltered, unaerated aquaria in a cold room at 10-15ºC.
PREPARATION OF MATERIALS
1. Limulus polyphemus
These animals are so hardy that they are shipped almost dry from some of the suppliers. We routinely order them from Marine Biological Labora- tories at Woods Hole. Massachusetts. We keep them in a marine aquarium at 12-15ºC where they will be fine for months if the aquarium is in good shape. (If animals are obtained from a gulf coast supplier, they should be kept at room temperature.) Animals kept in an inadequately-prepared aquarium may not respond well. Horseshoe crabs are scavengers, eating soft-bodied invertebrates, benthic algae, and detritus. They can be fed any number of marine invertebrate foods, or things such as chopped fish, but they seem to do fine for a short time with no feeding.
We have identified four suppliers for horseshoe crabs, but have actually ordered from only the first two.
108 1. Marine Biological Laboratory (617) 548-3705 ext. 325 Department of Marine Resources Woods Hole, Massachusetts 02543
L95 Limulus (small 2"-4"); one-half dozen (minimum order) $12.50. Animals are kept in aquaria at MBL, are shipped as necessary, and arrive in excellent shape. Keep at 12-15ºC.
2. Carolina Biological Supply Co. (800) 334-5551 2700 York Road Burlington, North Carolina 27215
16-2978 Horseshoe crab $3.35 ea or $16.50/5. Animals are collected and shipped after receipt of order. Exact delivery dates cannot be guaranteed. Sizes vary, but all are usable for the exercise. Keep at room temperature.
3. Ward's Natural Science Establishment (800) 962-2660 5100 West Henrietta P. 0. Box 92912 Rochester, New York 14692-9012
87 W 7530 Horseshoe crab $3.25 ea. Keep at room temperature.
4. Gulf Specimen Company, Inc. (904) 984-5297 P.O. Box 237 Panacea. Florida 32346
Ar-1070 Limulus polyphemus small (2.5-8.0 cm) $4.00 ea ($1.00 extra per animal to select for clear lateral eyes for electrophysiological recording). Keep in room temperature aquarium.
Physioscribe and Physioscribe problems
The Physioscribe is a chart mover and recorder equipped with a number of channels and produced by the Stoelting Corporation, Chicago, Il linois. In addition to the Time and Event channel, the Biopotential channel, and the Transducer channel which are needed for this exercise, a Stimulator module and other input channels are available, together with the necessary leads and cables. Two suppliers are:
E. J. McGowan and Associates 310 Lake Street Suite 110 Elmhurst, Illinois 60126
Carolina Biological Supply Co. address given above
We have found the Physioscribe to be a rugged and reliable piece of laboratory apparatus. Several phenomena, however, are worth mentioning. The extent to which these would apply to other student-model strip-chart recorders is not known.
First, it is important that all three pens are aligned so that they cross the same vertical line on the strip-chart at the same time. In the
109 worst case, the pen on the biopotential channel could be longer than that on the transducer channel. The record would then appear to have a response that preceded the stimulus.
Second, we have sometimes observed "cross-talk" between channels, especially when the sensitivity of the biopotential channel is high. The result is that timing pulses recorded on the time and event channel or light signals recorded on the transducer channel may cause spurious deflections on the biopotential channel which could be misinterpreted as ERGs. The latter circumstance can be reduced or eliminated by keeping the sensitivity of the transducer channel at the lowest level at which the dimmest light flash (i.e., that attenuated by 3.5 log units) will just barely register.
Third, one records response amplitudes that span a large range in some of these experiments. If the sensitivity is high enough to see the smallest response, the pen may be driven to saturation for large responses. Once the pen deflection reaches about 30 mm, it is probably a good idea to decrease the sensitivity. Unfortunately the Physioscribes are not calibrated, so it is difficult to know exactly what the sensitivity is. If the same flash is delivered before and after adjusting the sensitivity, the ratio of the two recorded response amplitudes can be taken to be the ratio of the sensitivities.
Finally, because the recording modules do not appear to be adequately grounded to the case of the Physioscribe, we find it necessary to run an additional wire from a convenient location on one of the modules (we use the nut on the input jack of the stimulator module) to the ground plug of the wall outlet. Failure to do this results in 60 Hz interference on the biopotential channel.
If one were to use a different recorder, it would be necessary to make sure that the low frequency cutoff of the channel used for the ERG is rather low, on the order of 0.1 Hz, so that it reproduces the ERG relatively faithfully.
3. Photoelectric transducers
We obtain these from:
Carolina Biological Supply Co. address given above
69-7567; $127.50 each. 4. Shutters -and -cable releases American Science Center, Inc. 5700 Northwest Highway Chicago, Illinois 60646
AGC Prontor II shutter; $2.00 each. These are cheap, second-quality shutters that we found at ASC. They work well enough if lightly lubricated. Cable releases can be obtained at any camera shop.
110 5. Filters
Cadillac Plastic and Chemical Co. (312) 342-9200 1924 North Paulina Chicago, Illinois 60622
Quarter-inch thick smoky gray plastic which attenuates light by 0.5 and 1.0 log unit is commercially available. We purchased scraps and cut them to 6 cm squares. 6. -- Other items The light-proof box, the aluminum plate, and the Plexiglas stage were of our own design, and constructed in the shops at Northwestern University.
7. Ag/AgCl electrodes
For electrodes, we use pure, uncoated 0.01" or 0.015" diameter silver wire. Three sources are listed below.
Medwire Corporation 121 South Columbus Mt. Vernon, New York 10553
A-M Systems, Inc. 1220 75th Street S.W. Everett, Washington 98203
World Precision Instruments, Inc. 375 Quinnipiac Avenue New Haven, Connecticut 06513
AGW 1010 30' of 0.01" @ $50.00 January 1987
In order to have stable recordings. Ag/AgCl electrodes are desirable. The principle of this electrode is described by Ferris (1974), among others. Essentially, one wants a reversible, non-polarizable electrode that will not develop a potential of its own with respect to the fluid it is in. To prepare Ag/AgCl electrodes, silver wires are made the anode and current is passed. As a power supply. we use a Hewlett-Packard 6202B DC power supply which has a maximum output of 50V and 1A. This is set to provide 4 volts, but this is not critical. Batteries could also be used for power. An ammeter in the circuit is convenient. The microgator clips (smooth alligator clips) that are used to hold the wires in solution rust after some time, and we monitor the current in order to know whether the wires are making contact. The chloriding bath is simply 0.9% NaCl. A silver cathode is not necessary since the reaction at the cathode is not important, but silver prevents contamination of the bath.
Before chloriding, wires are cleaned using soap to remove any grease and then rinsed in distilled water. Four wires are twisted together at the end where they contact the microgator clip. The cluster is suspended in the solution so that the free ends do not touch each other or the cathode. The bath containing the wires is placed in a dark box since the Ag/AgCl is
111 10 KR power supply battery
, microgator clips reference electrode wires to chlorided 0.9% NaCl
Figure 4. Schematic of apparatus for preparing Ag/AgCl electrodes.
sensitive to light. A current of about 1 mA is applied for 30 minutes to create a uniform dark grey AgCl layer. The set-up is shown in Figure 4. After chloriding, wires are rinsed with distilled water, inspected for unchlorided sections, and stored in the dark. Only the portion of the electrode that touches fluid needs to be chlorided. A pair of electrodes should last through at least two laboratory periods, but if they become scratched or if noise or drift is a problem, they should be replaced.
-INTACT -ANIMAL PREPARATION We have used the isolated eye preparation as described in the two versions of this exercise for several years with considerable success. It is, however, feasible to record the ERG from an intact animal. This is the preferred technique for doing research on the horseshoe crab, and it may be preferable for student exercises as well, since it would eliminate the dissection with its attendant technical difficulties. An intact animal preparation would also be more satisfactory for those students who find excision of the eye from the living animal to be aesthetically and morally objectionable.
Our preliminary experience with this preparation is good, although we have tried it on just one occasion using the advanced version of the exercise. We have not yet optimized the conditions for the intact-eye preparation, and therefore, we will not present it here. We will, however, offer the following preliminary observations.
Equipment -for-- intact animal preparation 1. All of the electrophysiological equipment listed previously can be utilized with the intact-eye preparation with the following exceptions. The plexiglas stage must be replaced with some sort of a stage that will hold the entire horseshoe crab - securely pinned to a board - at a suitable angle so that the eye is appropriately oriented toward the light. One of the two silver-chlorided electrodes should be replaced with a wick elec- trode. Same sort of a clamp will be necessary to hold the wick electrode
112 and to support the two banana jacks into which the Biopotential channel lead will plug.
2. None of the equipment listed for dissection and mounting will be necessary, save for the razor blade to scrape the surface of the eye. In addition, a sharp pin will be required to pierce the carapace for placement of the reference electrode.
Preparation of materials
1. It is most important that the crab be immobile. In order to accomplish this, the animal is attached to a small wooden board in the same way as for dissection of the lateral eye. A folded paper towel wetted with sea water is placed on the board beneath the animal. A T-pin is inserted through the edge of the carapace on each side, and these are hammered into the board. It is better to put the pins at an angle (i.e., not vertical) so that the animal can not push up with its legs and slide up along the pins. It will also help to push down on the center of the carapace so that the animal is spread out a bit laterally. A third pin is inserted through the back hinged section (abdomen) so that the hinge joint is immobilized. We have considered using heavy-duty rubber bands instead of pins, but have no experience with this yet.
2. Once the eye is scraped (see below), there may be fluid coming out onto the surface, allowing use of a chlorided silver wire electrode as in the isolated eye preparation. It seems preferable, however, to make sure that there is a fluid bridge by using a wick electrode (Barlow, 1984). These can be made by threading a piece of silk or cotton thread through a plastic tip for an automatic pipetter, and attaching a rubber Pasteur pipette bulb to the back end of the tip. A hole is made in the bulb with a pin and a chlorided silver wire is fed through, leaving enough outside the bulb to attach to the binding post on the stage. To fill the electrode, hold the thread against the outside of the pipette tip, squeeze the rubber bulb and draw in artificial sea water. The amount is not critical, provided that the wire is in the fluid and that the wick itself is wet. Bubbles in the pipette tip will not matter. Cut the wick so that about 2 mm sticks out of the pipette tip.
American Scientific Products, 1210 Waukegan Road, McGaw Park, Illinois, 60085-6788, has been our source for both amber latex Pasteur pipette bulbs (P5002-1) and for polypropylene pipette tips (P5059-80R or P5059-80D; 2", 2-200 µ1 capacity). Note that pipette tips with ribbed tops will not work.
Preliminary procedural tips
1. After the animal is secured on the mounting board, the eye should be scraped with a scalpel to reduce the resistance of the cornea. As in the isolated eye, actually slicing off a thin layer appears to work well. Wald et al., suggest that this is a painless procedure.
2. The board is mounted on a tilted stage so that the eye is oriented up toward the light. The stage must be positioned on the aluminum plate so that the eye will be in the light beam.
113 3. A small hole is made with a sharp pin somewhere on top of the animal for placement of the Ag/AgCl reference electrode, which just needs to make contact with the interior; it need not be pushed in deeply.
4. A wick electrode is brought in contact with the eye by adjusting the clamps holding it.
5. The photoelectric transducer is placed on top of the carapace so that it can collect adequate light.
6. The physioscribe cables are connected, the box is drawn forward over the preparation, and the experimentation can commence.
7. At the end of the experiment, the animal can be taken off the board and put back in the tank.
-SAMPLE -DATA Figures 5-8 show representative ERGS and graphs of data for 1) response amplitude and latency as a function of illumination, 2) response amplitude as a function of duration, and 3) the time course of dark adaptation. These are the key measurements one makes during the course of the laboratory period. The data in Figures 5, 7, and 8 were obtained by students using techniques des-
Log Stimulus Attenuation
Figure 5. Response amplitude and latency vs. illumination. Measurements were made in one dark-adapted isolated eye preparation. An interval of 45 sec was allowed between flashes, which were each 40 msec in duration.
114 Dark-adapted Light-adapted
Log Stimulus Attenuation
Figure 6. Response amplitude vs. illumination for the intact eye preparation under dark-adapted (filled circles) and light-adapted (open circles) conditions.
0 10 20 30 40 50 60 70 80 90 100 200 Flash Duration (msec)
Figure 7. Response amplitude vs. flash duration in two dark-adapted isolated eyes. Stimulus attenuation was 3.0 log units.
115 cribed in the Appendix. The data in Figure 6 were obtained by the authors using an intact-eye preparation. Since the Physioscribe sensitivity is uncalibrated, response amplitude is taken to be simply pen deflection in mm. The absolute size of the response cannot easily be compared among lab groups, but there would be reasonably good agreement if one plotted response as a percentage of maximum. The actual maximum amplitude of the ERG is generally between 100 and 500µV, as determined using the apparatus shown in Figure 3.
Some of these data can be fitted to equations if desired. A relatively large range of the intensity-response function is usually well described by the simple relation R = K1logI + K2, where K1 and K2 are constants. R is ERG ampli- tude, and I is illumination or intensity. This shows that a large range of stimulus intensities are encoded into a relatively small range of responses. Sometimes it has been possible to fit intensity-response curves more completely with the sigmoidal function familiar as the Michaelis-Menten equation, R = RmaxI/(I + s), where Rmax is the response at saturation, and s is the "half- saturation" corresponding to the value of I where R=Rmax/2. An interesting difference can be observed between the isolated eye and in- tact eye stimulus response functions in Figures 5 and 6. The ERG from the iso-
Adapting Flash 0 I I I I I I I I I I I -3 -2- 1 0 2 3 4 5 6 7 8 9 10 Time from Adapting Stimulus (min)
Figure 8. Time course of dark adaptation following a 5 second flash of unattenuated light for two isolated eyes. For one eye (open circles), the control responses are plotted at negative times. The test flashes were at 1.5 log units attenuation. For the other eye (filled circles), the student failed to record the control flashes in the lab report. The test flash illumination in this case was 3.0 log units attenuation. 116 lated eye tends to saturate at higher stimulus intensities, showing the entire form of the curve. whereas the response from the intact eye does not. This ap- pears to be a consistent difference, and it is surprising since the microscope illuminator is rather bright. It may be possible to saturate the response of the intact eye by using a photoflash unit (D. Mayette, personal communication), but then one would not be able to study the effect of stimulus duration.
The dark adaptation curves can usually be fitted very well by a single -t/τ exponential of the form R(t) = Rfinal(1 - e ), or if the response just after extinguishing the adapting light is greater than zero, -t/τ R(t) = (Rfinal - Rmin) (1-e ) + Rmin. where Rmin is the response at the beginning of dark adaptation, Rfinal is the dark-adapted response, and τ is the time constant. To fit these functions to the data, it is best to transform the equations to a form where some function of R is linear with time. For the first of these, this would be
-1n[(R(t) - Rfinal)/Rfinal] = t/τ.
REFERENCES
1. Armington, J. C. (1974) The Electroretinogram. Academic Press, New York. 2. Barlow. R. B. (1984) Circadian rhythms in the Limulus visual system. J. Neurosci. 3:856-870. 3. Barlow, R. B., Kaplan, E., Renninger, G. H., and Saito, T. (1987) Circadian rhythms in Limulus photoreceptors. -J. -Gen. -Physiol. 89:353-378. 4. Barlow, R. B., Powers, M.-K., and Kass, L. (1987) Vision and mating behavior in Limulus. In Symposium on Sensory Biology of Aquatic Animals (A. N. Popper, R. Fay, T. Atema, and W. Travolga, Editors), Springer- Verlag, in press. 5. Chapman, R. M. and Lall, A. B. (1967) Electroretinogram characteristics and the spectral mechanisms of the median ocellus and the lateral eye in Limulus polyphemus. -J. -Gen. -Physiol. 50:2267-2287. 6. Ferris, C. D. (1974) Introduction to Bioelectrodes. Plenum Press, New York. 7. Hanna, W. J. B., Horne. J. A., Renninger, G. H., Kaplan, E., and Barlow, R. B. (1985) The tail of Limulus: an extraocular photoreceptor organ for a circadian clock. Invest. Ophthalmol. Visual Sci. Suppl. 26:113 (abstract). 8. Hartline. H. K. (1927) A auantitative and descriptive study of the electric response to illumination of the arthropod eye. Am. J. Physiol. 83:466-483. 9. Hartline. H. K. and McDonald, P. R. (1947) Light and dark adaptation of single photoreceptor elements in the eye of Limulus. J. Cell. Comp. Physiol. 30:225-253. 10. Kaplan, E. and Barlow, R. B. (1976) Energy. quanta, and Limulus vision. -Vision -Res. 16:745-751. 11. Millecchia, R.. Bradbury, J., and Mauro, A. (1966) Simple photoreceptors in Limulus polyphemus. Science 154: 1199-1201.
117 12. Miller, W. H., Ratliff, F., and Hartline, 9. K. (1961) How cells receive stimuli. -Sci. -Amer. 205(3):223-238 (Reprint #99). 13. Oakley, B. and Schafer, R. (1978) Visual reception: frog electroretino- gram. In Experimental Neurobiology, The University of Michigan Press, Ann Arbor, MI pp. 166-173. 14. Packer, L. (1967) Bioelectric Potentials II. Electrical activity of the Limulus eye. In Experiments in Cell Physiology, Academic Press, New York, NY pp. 269-276. 15. Wald, G.. Hopkins, J., Albersheim, P., Dowling. J., and Denhardt, D. (1968) Electrical activity of a sense organ: the Limulus eye. In Twenty- six Afternoons of Biology (Second Edition), Addison-Wesley Publishing Company, Inc., Reading, MA pp. 112-116.
ACKNOWLEDGEMENTS
We thank Mr. Ed McGowan of E. J. McGowan and Associates, Elmhurst, Illinois, for making a Physioscribe of the latest model available for us to demonstrate. We also thank Dr. J. S. Bjerke for assistance with the manu- script.
APPENDIX
SENSORY PHYSIOLOGY: THE ELECTROETINOGRAM OF THE HORSESHOE CRAB
Introduction
One of the fundamental characteristics of all cells, and consequently, of all living organisms as well, is their ability to monitor external conditions and to detect changes in these parameters. This characteristic - the sensory capability of living organisms - may in general be referred to as "sensitivity". Even the simplest single-celled prokaryotes exhibit sensitivity; bacteria can detect food sources as indicated by their preferential movement up a concentration gradient of nutrient molecules. Photosynthetic organisms from single-celled algae to the most complex plants are exquisitely sensitive to light. It is among the animals, however, that the characteristic of sensitivity is most conspicuously developed.
During the long course of animal evolution, certain cells have become specialized in a variety of ways for carrying out different sensory functions in a highly efficient way. These receptor cells may occur singly, scattered throughout the body, or they may be aggregated into large and complex sense organs which function in remarkable ways. For example, sensory organs often 9 operate reliably over an enormous range of stimulus energies - 10 -fold in the case of the human auditory system. Others can discriminate minuscule differ- ences in stimulus intensity - 0.002ºC in the case of the reptilian pit organ which detects infrared radiation.
Receptor cells are typically specialized for detecting a single specific
118 form of energy. On this basis, they can be categorized into chemoreceptors, mechanoreceptors, photoreceptors, electroreceptors, and thermoreceptors. The receptor cells responsible for sensing all of these different parameters in the external world are quite different in their structural and functional proper- ties. These differences are in large part due to the very different types of stimulus energy to which they are sensitive. At the same time, there are also important similarities between different types of receptor cells. This is because they all function as transducers, converting energy of various sorts into membrane potentials. These receptor potentials then trigger the genera- tion of trains of action potentials, either directly in the axon of the recep- tor cell or, by causing the release of a transmitter substance which ultimately leads to activity, in the axon of an afferent nerve. Consequently, the neural outputs of all different sensory systems are essentially identical. Thus, by studying a specific sense organ, one can learn principles that apply broadly to sense organs in general, as well as features specific to the organ under study. In addition, since there has been evolutionary conservation of the principles employed, one may study sensation in a specific preparation - even from an invertebrate - and elucidate principles that are applicable to sensory systems in all animals.
Some of the principles that can be illustrated are 1) gradation of response as a function both of stimulus intensity and of stimulus duration, 2) gradation of response latency (i.e., the time lag between receipt of the stimulus and on- set of the response) as a function of stimulus intensity, and 3) adaptation under conditions of continuous stimulation. These properties could be investi- gated with intracellular recording from individual receptor cells, or with ex- tracellular recording of action potential discharge at some other cell in the system. Often, however, they can more easily be studied by recording massed potentials or field potentials. These represent the sum of extracellular cur- rent flow from a group of neurons or other cells.
In this exercise, you will record the electroretinogram (ERG) from the lateral eye of the horseshoe crab, Limulus polyphemus. The ERG is a field potential from the eye in response to light; it is analogous to the electro- cardiogram (ECG), which represents the sum of current from depolarization of cardiac muscle fibers. ERGs can be recorded from the eyes of vertebrates and invertebrates by placing a recording electrode in contact with the cornea or external surface of the eye and a reference electrode at some remote point on the animal. The vertebrate ERG is a complex series of potential changes having components that arise from several of the cell types in the retina. Inverte- brate ERGs are much simpler, reflecting the simpler structure of the eye. There are only two components to the Limulus ERG; a transient, corneal-negative potential and a smaller, steady component that is maintained for the duration of the stimulus. Because of the limitations of your recording instrument (the Physioscribe), you will see only the transient component. You will study variations in the amplitude of the response as a function of changes in stim- ulus conditions. Before continuing with the details of your experiment, how- ever, a digression about the biology of Limulus is in order.
Limulus polyphemus is a marine arthropod found in tidal regions along the Atlantic coast. While they are called crabs, they are not crustaceans as are the edible crabs and other "true" crabs. Rather, they are classified in the class, Merostomata, in which they are the only Living species. They are dis- tantly related to the ticks, mites, and spiders - members of the class Arach-
119 nida. The fossil record documents that as a species, Limulus polyphemus has existed for at least 300 million years. All of its close relatives, phylo- genetically speaking, are extinct. Like other arthropods, horseshoe crabs must molt in order to grow, and the lenses of the eyes are shed with the rest of the exoskeleton. Over at least a few years, the age of a horseshoe crab is approx- imately equal to the diameter of its carapace in inches. The maximum size is about thirty inches.
Limulus adults have two large compound lateral eyes and two small simple median eyes. Both have been useful in understanding certain aspects of visual physiology, and the importance of this animal for research in vision cannot be overestimated. This is largely because the recordings of the activity of sin- gle optic nerve fibers and of membrane potentials in photoreceptors were feasi- ble in Limulus many years before such recordings could be obtained from verte- brates. The experiments which you will do were pioneered in the 1930s and 1940s by Professor H. Keffer Hartline who worked at the University of Pennsyl- vania, the Marine Biological Laboratory in Woods Hole, Massachusetts, and at Rockefeller University. In 1964, Professor Hartline was awarded the Nobel Prize in Physiology and Medicine for his work on the visual system of Limulus.
The compound lateral eyes of the horseshoe crab each consist of approxi- mately 300 individual, identical units known as ommatidia. For an illustration of ommatidial structure, a description of the optical properties of compound eyes, and a discussion of some aspects of the visual physiology of Limulus, you are referred to pages 262-265 and 277-279 of Eckert and Randall ANIMAL PHYSI- OLOGY (Second edition).
You will begin the experimental portion of this exercise by excision of one of the lateral eyes from a horseshoe crab, leaving it attached to a section of the carapace. Remarkably, this eye will continue to generate responses for hours with no special precautions concerning oxygenation, buffering, provision of substrates, etc. You will mount your preparation over sea water in a small dish and place electrodes on the surface of the eye and beneath it in the sea water which contacts the back of the eye. The preparation will be housed in a box that serves two functions: First, it provides electrical shielding, and second, it ensures that room light does not reach the preparation. The only light the eye will receive will be provided by a simple optical system. Responses from the eye will be recorded on the biopotential channel of the Physioscribe. A photoelectric transducer located inside the box will be connected to a transducer channel of the Physioscribe in order to record the timing of the light flashes.
After your preparation has been assembled, you will first need to allow the eye to adjust to darkness in the box so that it will be maximally sensitive to the flashes you deliver. You will be able to tell when the eye has attained its maximal dark-adapted sensitivity by looking at its response to weak test flashes. You will then be able to study stimulus-response relationships in the steady-state, dark-adapted eye. In doing this, and throughout the exercise, it is essential that you realize that a single bright flash or too many dim ones spaced too closely together will light-adapt the eye and change its sensitivity.
Once you have determined some aspects of the physiology of the dark- adapted eye, you will proceed to study the time-course of the dark-adaptation
120 process, i.e., how the eye's responses change after exposure to a steady light. Finally, you will determine how the eye's responses change during continuous exposure to a low level of light, i.e., during the process of light-adaptation. You will then have the opportunity to devise your own additional experiments to investigate further aspects of the physiology of vision in the Limulus eye. Questions which can be answered using your experimental preparation will be posed for you to consider.
Objectives
When you have completed this laboratory exercise, you should be able to:
1. describe in general how to use a simple optical system to experimentally stimulate a visual system preparation 2. describe the waveform of the Limulus ERG, and compare it to the vertebrate ERG as illustrated on page 267 of Eckert and Randall ANIMAL PHYSIOLOGY (Second edition) 3. describe the essential structural features of ommatidia and of compound eyes; explain what is meant by dark adaptation, light adaptation, and response latency 4. tell how response amplitude depends on stimulus intensity and stimulus duration in the dark-adapted Limulus eye; interpret data which illustrate these relationships 5. tell how response latency depends on stimulus intensity; interpret data which illustrate this relationship; explain why an experiment to show the effect of stimulus duration on response latency was not suggested 6. describe the time-course of dark adaptation; interpret data which illustrate this phenomenon 7. describe the effect of maintained illumination on the stimulus-response relationship
Laboratory Procedures
1. Instrumentation.
a. Optical system. To stimulate the eye, you will use a simple but effective optical system consisting of three parts - light source, neutral density filters, and a shutter. The shutter is built into the box top; the filters and the light source are positioned above so that they will be accessible. The source provides a large continuous amount of light, the filters attenuate the light to different degrees, and the shutter controls the timing of flashes that reach the eye. This system is preferable to simply switching the light on and off because 1) the duration of the flashes can be more accurately controlled, 2) the intensity changes instantly when the shutter is opened. and 3) the light switch could introduce electrical artifacts. One could change the light intensity by simply changing the voltage to the illuminator, but filters are preferable because intensities can be 1) adjusted more reproducibly, and 2) varied over a wider range.
121 1. Light source. A Nicholas microscope illuminator will be used as the light source. 2. Shutter. The shutter is operated by a push-button cable release. Because it is a mechanical shutter, one must load the internal spring with the small cocking lever before each use. You will also need to set the duration for which the shutter is open. This is done by rotating the toothed wheel at the outer edge of the shutter. It can be set for durations as short as 1/200 of a second (shown as 200 on the shutter) and as long as 1 second (shown as 1). In addition, there is a setting labelled T. This is the timed position, and one trip of the release will open the shutter and leave it open. The shutter will not close until the cable release is pressed a second time. You will use this setting when lining up your preparation with the light beam, and later, when studying light adaptation. 3. Filters. The filters used in this experiment attenuate all wavelengths approximately equally, and are therefore called neutral density filters. Optical density (OD) is specified in log units of 1 attenuation. A 1 log unit filter (OD = 1.0) transmits only 1/10 or 1/10 or 10% of the incident light. A 2 log unit filter (OD = 2.0) transmits 1/102 or 1/100 or 1% of the incident light, and a 0.3 0.3 log unit filter (OD = 0.3) transmits 1/10 or 1/2 or 50% of the incident light. The advantage of using optical density is that the units are additive; a 2 log unit filter and a 1 log unit filter together attenuate the light by 3 log units. In your system,' the filters will simply be stacked on top of the shutter. You will have three 1.0 log unit filters and one 0.5 unit filter to work with in this exercise. Used in various combinations. these will enable you to stimulate at seven different attenuated light intensities (plus, of course, the intensity of unattenuated light). b. Physioscribe. You will be using the Time and Event channel, the Biopotential channel, and one Transducer channel for this exercise. 1. Time and Event Channel: use to set chart speeds and to gauge short time intervals. 2. Biopotential channel: displays the ERG. A signal cable is plugged into this channel; the two banana plugs at the other end of the cable are seated in the binding posts mounted in the lucite stage. 3. Transducer channel: displays the timing of the light flashes. The cable from the photoelectric transducer is plugged into this channel. The transducer itself will be positioned beneath your preparation on the lucite stage inside the Light-tight box. 4. Grounding: the Faraday cage (mounted inside the light-tight box), the heavy aluminum plate, and the Physioscribe will all be grounded.
2. Preliminary testing.
a. Position the bottom of a small glass petri dish close to the binding posts on the lucite stage. 1. Arch the wire electrodes up over the edge and down into the dish to touch its bottom.
122 2. These are silver-silver chloride electrodes, so take care not to scrape the silver chloride off the silver wire core. b. Fill the dish approximately half full of sea water, making sure that the electrodes are immersed. c. Position the lucite stage in the center of the heavy aluminum plate, with the binding posts toward the left end. d. Secure the photoelectric transducer to the lucite stage on the lower level beneath the petri dish if it is not already in place. e. Slide the box forward over the aluminum plate. 1. With the curtain open and the shutter open (use the T setting), position the illuminator so that its light beam shines down through the center of the shutter and into the center of the petri dish. 2. Lower the curtain and secure its edges; close the shutter. f. Turn on the Physioscribe; fill the pens; switch on the illuminator (use the 3 setting). g. Test the biopotential channel as follows: 1. Slowly and carefully turn up the sensitivity knob of the biopotential channel. You should reach the maximum sensitivity level without detecting 60-cycle noise. 2. If you pick up noise below a setting of 70, check the grounding connections or summon your teaching assistant. h. Adjust the sensitivity of the transducer channel and test it out as follows: 1. Using 1/25 second flashes with the light attenuated by 3 log units, set the sensitivity of the transducer channel so that the deflection of the pen is minimal. Begin with a setting of 10, and increase by increments of 5. 2. At this sensitivity, unattenuated light flashed with a duration of 1/25 second would give a large but not maximal deflection; does it? 3. With an attenuation of 1 log unit, observe the response of the transducer channel when flashes of different duration (1/50, 1/25, 1/10 1/5, 1/2, and 1 sec) are given. Note that there is a deflection at the onset of light and, for longer flashes, at the offset, but that there is no plateau while the light is on. This is a limitation of the Physioscribe, but not a serious one, since you really only need to keep track of when the flash began. (You may also note that the timing mechanism in the shutter is not reliable; it tends to stick at longer flash durations. Fortunately, you will be using short flashes for the most part.) When you have completed checking out your system, return the sensitivity setting of the biopotential channel to its minimal level, but leave the sensitivity of the transducer channel as you have adjusted it. 1. Turn the Physioscribe and the illuminator off. 2. Open the box and slide it back to expose the aluminum plate and your experimental set-up.
123 3. Empty the sea water from the petri dish and dry it with a Kimwipe. 4. Construct two or three small clay pyramids in the middle of the dry glass petri dish. These should be about 0.5 cm across and 0.5 cm high. Your teaching assistant will have a sample illustrating the appropriate dimensions at the front desk.
3. Dissection of the lateral eye; mounting the preparation.
a. Pick up a dissecting kit and a horseshoe crab which has been secured to a dissecting board from the preparatory lab. b. First, you will have to collect a small sample of blood in a hypodermic syringe using the method of heart puncture. The heart runs antero- posteriorly in the dorsal midline of the body. 1. Locate the hinge which joins the horseshoe-shaped carapace with the angular posterior section (abdomen) of the body. 2. Aim the needle anteriorly and push it through the elastic hinge cuticle in the midline of the animal's body (see Figure 1) to a depth of approximately 0.5 cm. 3. Pull the plunger slightly and adjust the depth of the needle until the syringe begins to fill with blood. Collect approximately 0.5 ml. The blood will be almost clear or slightly bluish; this is due to the presence of the blue respiratory pigment, hemocyanin, which becomes bluer as it is oxygenated. 4. Lay the syringe with its contained blood aside for future use. c. Examine the lateral eye under the binocular dissecting microscope. Note the facets, each of which represents the lens over one ommatidium. 1. While holding your animal securely, slice off a section of the waxy epicuticle over one of the lateral eyes with a sharp, single-edge razor blade. You should slice deeply enough so that the surface ends of some of the ommatidia are removed with the waxy layer, but not so deeply that you remove most of the eye. 2. With the corner of the razor blade, make a rectangular incision approximately 1.5 cm square through the carapace around the eye (see Figure 2). You need cut only through the cuticle (approxi- mately the thickness of a fingernail); not deeply into the animal, which is only a few millimeters thick in this area. 3. With forceps, lift up the section of excised carapace, separating it from underlying tissues with the tip of a sharp scalpel. Do not, however, poke your scalpel in so far that you damage the soft, bottom side of the eye. d. Pick up the section and, holding it securely along its edges between your fingers, dry the surface of the cuticle carefully with a Kimwipe. The dryness of the cuticle around the eye is a critical factor. e. Take the dry-surfaced section securely in the forceps again, and with the narrow end of a stainless steel spatula, apply a wall (2 mm high) of silicon grease around the edge on all four sides. The integrity of the wall is a second critical factor.
124 1. Why are a dry surface and an intact wall so important? Shortly, you will place one recording electrode in the liquid within the wall where it will contact the surface of the eye. Your reference electrode will be in the sea water bath which will be in contact with the back of the eye. It is essential that the electrical resistance between these two compartments be very high, and this requires 1) making a good wall with no Leaks and no gaps, and 2) not allowing fluid to spill over from one compartment to the other. 2. Electrical short-circuiting across or beneath a faulty wall is a comnon cause of trouble in this exercise. f. Mount the rectangle between your clay pyramids, securing its edges firmly in the clay so that it won't later float off. g. With a pasteur pipette, add sea water to the petri dish to a depth such that its surface is level with the top of the mounted section of cara- pace. Tilt the dish appropriately so that trapped air bubbles escape from beneath the section. h. Expel a couple drops of blood from the syringe onto the cut surface of the eye. i. Place the petri dish containing your preparation in its place on the lucite stage. j. Position one wire electrode (black) in the sea water, and the other (red) in the drop of blood over the surface of the eye. k. Draw the box forward over the preparation. 1. Stack 2 log units of filter on the open shutter, switch the illuminator on (use the 1 setting), and quickly check to make sure that the light beam illuminates our preparation; then close the shutter promptly. m. Position a second Nicholas illuminator to serve as an adapting light source for Experiment IV. 1. Remove the patch covering the porthole on the right side of the box. 2. Aim the light beam of the second illuminator (use the 1 setting) into the box somewhat to the side of your preparation. Then turn off this illuminator and cover the porthole again. n. Secure the curtain and wait 5 minutes for the eye to dark-adapt. o. Nota bene! Take care not to knock or jar your preparation, pull on the electrical cable, or move the second illuminator during your subsequent experiments. If the alignments are disturbed and you have to open the light-tight box again, valuable time will be lost in waiting for the eye to dark-adapt a second time.
4. Final testing.
a. Set the chart speed of the Physioscribe at a slow speed (approximately 1 mm/sec). b. Slowly and carefully turn up the sensitivity of the biopotential channel to maximal. You should observe no greater 60-cycle noise than you did in your preliminary test.
125 1. If this is not the case, summon your teaching assistant. 2. Set the sensitivity of the biopotential channel at approximately 40 to begin your final testing of the system. c. Turn the top illuminator on; use the 3 setting for all experiments. d. Set up and administer a test flash at 1/25 sec of light attenuated at 1 log unit. e. If there is a response, wait 5 minutes and repeat with another identical flash. 1. Continue with repeated identical flashes after 5 minute intervals until the response amplitude no longer changes; at this point, your preparation has reached the stable, dark-adapted state. 2. Proceed to Experiment I. 3. During the final testing, and throughout all experiments, you will need to keep track of the following: Write them on the chart for each flash! ----- a. Absolute time - clock time. b. Attenuation of the light beam - number of log units. c. Flash duration - shutter speed. d. Sensitivity setting of the biopotential channel. f. If there is no response, summon your teaching assistant. You will need to re-check your electrical connections, the alignment of your light beam, the intensity and attenuation of the light, the placement of your electrodes, the quality of your dissection, and the status of your mount. If these all appear to be in order, close the dark box again, wait 5 minutes, and try part d again. 1. If there is still no response, take the eye out of the dish, dry the carapace and re-grease it, re-mount it as described in the previous section, and proceed with final testing once again. 2. As a last resort, you will have to do a second dissection.
5. Experiment I. Response amplitude and latency of response as functions of stimulus intensity.
a. Set the Physioscribe speed at maximum. 1. Run the chart mover only at the times when you will actually be delivering flashes; you will waste enormous amounts of costly paper if you run it continuously. b. The sensitivity settings of both channels should remain as previously set. c. Use 1/25 second flashes for this experiment. d. Begin with the dimmest flash possible (3.5 log units of attenuation); stimulate the eye and record the response. It is important that you get good traces on both recording channels, although the responses will be very small. Readjust the sensitivities of the channels if necessary, and repeat the flash until you get a good trace.
126 e. Continue to stimulate the eye with progressively brighter flashes of light. By stacking different combinations of filters, six additional intensity levels are possible. f. How long should you wait between stimuli so that the response to a given flash is not influenced by the previous stimulus? Your experimental protocol will need to include controls which show that your stimuli were not given too close together. There are several possible controls that could be done. You will have to think about the problem; to devise a suitable control, and include it during the course of the experiment.
6. Experiment II. Response amplitude as a function of stimulus duration.
a. Continue with the previous Physioscribe settings. b. Choose an intensity for which the shortest possible flash (1/200 sec) gives a minimal response; stimulate the eye, and record the response. c. Continue to stimulate the eye with progressively longer flashes of light at the same intensity; there are seven additional settings up through 1 second on your shutter. d. Use the data obtained in your controls for Experiment I to determine the approximate delay time between successive stimuli.
7. Experiment III. Time course of dark adaptation.
a. Slow the chart speed to approximately 1 mm/sec, and allow the chart to run continuously during this experiment. b. Choose a stimulus duration and intensity that will give a half-maximal response. 1. Stimulate repeatedly at suitable intervals until you obtain at least two equal responses. 2. Note the minimal time that must elapse between these flashes to obtain equal responses. This is your dark-adapted control. Make sure that you do not underestimate this time. c. Light-adapt the eye by giving unattenuated light for 10-15 seconds; the T setting of the shutter is useful here. d. Return immediately to the stimulus conditions used for the dark-adapted control flashes. 1. Deliver flashed regularly at the interval previously determined until the dark-adapted control response is once again obtained.
8. Experiment IV. Sensitivity during light adaptation.
a. This experiment is in part a variation on the previous experiments. Here you will measure responses while a second dim light enters the box; this light will serve to partially light-adapt your preparation. As an adapting Light source, you will use a second Nicholas illuminator which you have already positioned to shine through the side porthole. b. Continue with the previous Physioscribe settings.
127 c. To begin the experiment, you should repeat a mini-Experiment I using only three different intensities of light. This will give you data for a second dark-adapted intensity-response curve (Experiment IV-A). 1. Note that since you are running the Physioscribe at a slow speed, you will not have data for a second intensity-latency curve as you had in Experiment I. d. Next, proceed to repeat the dark-adapted control of Experiment III, giving flashes at an intensity which produces an intermediate response repeated at the standard interval (section 7. b. 1.&2.).This will give you data for a second dark-adapted control (Experiment IV-B). e. After 2 or 3 flashes, open the porthole. This should produce a response, the magnitude of which will tell you the intensity of the adapting light from the second illuminator. f. Continue to administer standard test flashes at the standard interval. 1. The response to your first stimulus during light adaptation should be reduced. If this is not the case, you should reposition the light beam of the second illuminator. 2. Administer test flashes with the porthole open until the response has stabilized. This will give you data for a light-adaptation curve (Experiment IV-C ) . 3. Once the response has stabilized, take data for a light-adapted intensity-response curve (Experiment IV-D). Use at least the same three light intensities that you used in 8.c. above. g. Conclude by closing the porthole and continuing with the standard test flashes until the dark-adapted control response is once again attained. These data will enable you to plot a second dark-adaptation curve (Experiment IV-E).
9. Experiment V (Optional). Latency of response as a function of response amplitude under conditions of light adaptation.
a. Design and carry out an experiment which will provide data to answer the question: Is the latency of equal-sized dark- and light-adapted responses the same?
10. Experiment VI (Optional). Time course of dark adaptation under different stimulus conditions.
a. Repeat Experiment III using stimuli of different durations or of different intensities. Your data should enable you to answer two questions: I. In what respect(s) is(are) all dark-adaptation curves similar? 2. In what respect(s) is(are) all dark-adaptation curves different?
11. Experiment VII (optional). Intensity-response curves as a function of response duration under conditions of dark adaptation.
a. Repeat Experiment I using stimuli of different durations. b. Your data should enable you to answer the question: How is response
128 amplitude related to the total amount of light (i.e., intensity x duration) administered? 1. Does an increase in intensity by a factor of 2 exactly compensate for a decrease in duration by a factor of 2? 2. Over what range can intensity and time be traded off? (In humans, the sensitivity to flashes of equal total energy is constant for durations less than 100 msec).
12. Termination of the exercise.
a. Return the dissecting equipment and the dish containing your preparation to the preparatory lab. b. Wipe the grease off the spatula; return it and the container of silicon grease to the front lab bench. c. Clean the salt from the stage of the binocular dissecting microscope. d. Turn the Physioscribe off; empty the pens; return all sensitivity knobs to their minimal settings. e. Check out your data with your teaching assistant
Data Analysis and Report -- 1. Preliminary testing.
a. Strip chart - in Appendix.
2. Final testing.
a. Strip chart - in Appendix.
3. Experiment I.
a. Summarize the data in tabular form. 1. Stimulus intensity in log units of attenuation. 2. Response amplitude in mm 3. Latency of response in mm, convert to seconds. b. Plot response amplitude as a function of stimulus intensity. This will be your dark-adapted intensity-response curve. c. Plot latency of response as a function of stimulus intensity. d. State conclusion(s) which can be drawn from these data. e. Strip chart - in Appendix.
4. Experiment II.
a. Summarize the data in tabular form. 1. Stimulus duration in seconds (from shutter). 2. Response amplitude in mm.
129 3. Latency of response in mm, convert to seconds. b. Plot response amplitude as a function of stimulus duration. c. State conclusion(s) which can be drawn from these data. d. Strip chart - in Appendix.
5. Experiment III.
a. Summarize the data in tabular form. 1. Time in seconds after previous flash for dark-adapted controls. 2. Time in minutes after the end of light-adaptation for experimentals. 3. Response amplitude in mm. b. Plot response amplitude as a function of time following the end of light-adaptation. This will be your dark-adaptation curve. c. State conclusion(s) which can be drawn from these data. d. Strip chart - in Appendix.
6. Experiment IV.
a. Summarize the data in tabular form. b. Plot the data for Experiment IV-A on the graph which you made for Experiment I as described in 3.b. above. c. What was the intensity of the adapting light from the second Nicholas illuminator. How did you determine this? d. For Experiment IV-C, plot response amplitude as a function of time following the beginning of light adaptation. This will be your light- adaptation curve. e. Plot the data for Experiment IV-D on the graph which you made for Experiment I as described in 3.b. above. Your light-adapted intensity- response curve will thus be directly comparable with your two dark- adapted intensity-response curves. f. Plot the data for Experiment IV-E on the graph which you made for Experiment III as described in 5.b. above. g. State conclusion(s) which can be drawn from these data. h. Strip chart - in Appendix.
7. Optional Experiments.
a. Outline your experimental protocol so that your experiment can be repeated by someone else with the same results. b. Tabulate the data in an appropriate way. c. Plot data on graphs if appropriate. d. State conclusion(s) which can be drawn from these data. e. Strip chart - in Appendix.
130 Chapter 7
Methods to Process and Identify Symbiotic Fungi in the Roots of Vascular Plants
Iris Charvat
Department of Botany University of Minnesota 220 BioSciences Center 1445 Gortner Avenue St. Paul, MN 55108
Iris Charvat, an associate professor in the Botany Department at the University of Minnesota, studies the development and cell biology of fungi including vesicular arbuscular mycorrhizae (VAM). She is also interested in the distribution of mycorrhizae in aquatic systems and their possible role in the evolution of land plants. She received her M.S. in botany from the University of Illinois, and Ph.D. in Biology from the University of California, Santa Barbara in 1973. She is presently the Editor of the Mycological Society of America Newsletter. Last year she received the Morse-Amoco Award for outstanding contribution to Undergraduate Education at the University of Minnesota.
131
Introduction:
Vesicular arbuscular mycorrhizae (VAM) are present in the roots of almost all vascular plants. These common, soil-borne fungi belong to the family, the Endogonaceae (Zygomycotina) and produce fungal structures in the cortex of the roots. VAM play a crucial role in the mineral nutrition of plants by transferring phosphorus and other minerals from the soil to the plant. Techniques for obtaining VAM samples from natural sources and from inoculated pots are outlined. Methods of isolating spores, of processing, staining and clearing root samples, and of identifying the fungal structures are discussed
Isolation of Roots and VAM Spores from Natural Sources and from Inoculated Pots. [Wet sieving and decanting methods for spore isolation, modified from Gerdemann (1955), and Gerdemann and Nicolson (1963). See Schenck (1982) for other details concerning methods for spore and root isolation.] Methods:
1. Processing a plant plus surrounding soil or a soil core for isolation of the roots and the VAM spores present in the soil. Remove the plant from the pot or from its natural environment along with the soil around the roots. Cut off the top. Soak the root system in a large container of tap water and then wash the roots with a running stream of tap water to remove the soil. All of the soil particles are collected in a large container. Break-up the larger soil particles with your hands. 2. Collection of intact roots. Save the roots to examine for VAM and/or use as an inoculum. Place the roots in a plastic bag containing a small amount of water so they will not dry out. If the roots will not be examined or used in a couple of hours, put the bag in the refrigerator, or the roots may be fixed in formal-acetic-alcohol (FAA) to be processed and examined later for VAM structures. 3. Collection of additional roots and large pieces of organic debris (to be discarded) on a 2 mm sieve.
Pour the soil-water mixture discussed in #1 through a 2 mm Nalgene sieve that has been placed above its bottom half, a 5 liter solid container. The soil-water mixture is collected in the bottom half. Remove the large pieces of organic matter from the sieve, and then collect the roots by picking off the larger ones and placing them in a small container of water or fixing them in FAA. 4. Collection of spores (propagules) of mycorrhizal fungi on a 38 um sieve. Stir the soil-water mixture and pour part of this soil suspension onto the surface of the 250 um sieve that has been stacked on top of the 38 um sieve in the sink. The top sieve will concentrate most of the soil particles; so only the fine soil particles along with the VAM spores will collect on the 38 um sieve. Use a jet of tap water
133 to wash the spores and finer soil particles through the top sieve. This water will quickly pass through the top sieve, but it is difficult to get the water to drain through the 38 um sieve because of the build up of fine soil particles. The two sieves are separated and a jet of water on the surface of the 38 um sieve permits the water to drain at that spot. Then the two sieves are stacked together again, and the process is repeated until the water washing through the two connecting sieves leaves the bottom sieve colorless. Generally this means the soil in the top sieve is washed 3 times. Note: The water drains slowly through the lower sieve; hence, the 38 um sieve must be continuously checked by separating the two sieves and visually looking at the height of the water. If the water does overflow the lower sieve, spores are lost. 5. Concentration of VAM spores in soil pellet. The screenings are collected from the 38 um sieve by washing them into a beaker using a small stream from a wash bottle. Then, the soil is placed into a series of 50 ml, round-bottom centrifuge tubes for use in a Sorvall RC-2B centrifuge (1/3 soil and 2/3 tap water up to within 2 1/2 cm of the top of the tube). The mixture is thoroughly stirred, the tubes are balanced and then centrifuged at 4000 rpm for 5 minutes. The supernatant containing the light organic matter is decanted. 6. Isolation of VAM spores in 50% sucrose. The pellet containing the spores is suspended in a 50% sucrose solution and is mixed well. The tubes are balanced and centrifuged again at 4000 rpm for 5 minutes. The supernatant that contains the spores are poured over the clean 38 um sieve and immediately washed with tap water to remove the sucrose. The VAM spores are washed into a beaker of water using a water bottle. This entire procedure (#6) is repeated one or two more times to obtain more spores. 7. Isolation of VAM spores free of nematode cysts. The VAM spores can be cleaned further by preparing a gradient sucrose solution in a 15 ml centrifuge tube (clinical centrifuge tubes). Place 3 ml of each of these solutions in this order: bottom layer, 50% sucrose solution; middle layer, 15% sucrose solution. Slowly add the spore suspension into the tube. The water in the spore suspension makes up the 3 ml top layer. Centrifuge at 2750 rpm (#7 on the speed dial of the clinical centrifuge) for 5 minutes. Collect the spores from the first (top) interface and wash off the sucrose solution. These spores will be free of nematode cysts and other contaminants found in the other interface (Tang, 1986). Make a wet mount and examine a small sample of the spores with the dissecting and compound microscopes. 8. Sterilization and storage of spores. Sterilization: Concentrate the spores on the 38 um sieve and then with a squeeze bottle wash the spores into the sterilization solution. Soak the spores in this solution for 20 minutes. Pour the spores onto the filter which is sticking on the funnel, and rinse with distilled water 5 times. make a hole in the filter paper and wash the spores into a 100 ml graduated cylinder. Add more water until the volume is 100 ml. Samples can be removed to determine the spore number (Tang, 1986).
134 Short term storage: The isolated, non-sterilized spores may be stored for 2 days in the refrigerator in a beaker of water sealed with parafilm. Long term storage: Fill a clean petri dish 3/4 full of clean, dry sand and pour the sterilized and rinsed spore suspension over the sand. Allow the water to evaporate from the sand by leaving the cover off. This may take overnight to several days. When the sand is dry, cover the petri dish and seal it with parafilm. Store the sealed and labelled dish at 4º C. until the spores are used.
9. Retrieval of long term stored spores. Pour the dry sand/spore mixture onto the 250 um sieve (top), which has been stacked on the 38 um sieve (bottom). Using distilled water, rinse the spores through the top sieve and collect them from the bottom one (38 um) with a squeeze bottle containing distilled water. The spores can be washed into a petri dish. 10. Isolated spores may be identified and/or used to inoculated other plants.
Variation: A small amount of Calgon or other surfactant may be added to the 50% sucrose solution (#6) to increase the spore yield (Ianson and Allen, 1986).
Materials and Equipment:
Centrifuges-Sorvall RC-2B and a clinical centrifuge. Centrifuge test tubes: 50 ml, round bottom tubes for Sorvall centrifuge 15 ml glass tubes for clinical centrifuge Water bottles with bent spout, filled with distilled water. Sieves: 2 mm Nalgene 250 um mesh sieves 38 um mesh sieves Beakers: 250 ml and 100 ml Petri dishes Clean sand (small amount to fill 12 petri dishes). Rubber tube (long enough to attach to tap water faucet). A plastic attachment at the end will provide the water jet Compound microscopes Dissecting microscopes Preparation of Solutions: 15% sucrose (w/w) - 15 g. sucrose + 85 ml distilled water 50% sucrose (w/w) - 50 g. sucrose + 50 ml distilled water Sterilization solution (Tang, 1986) - 2 g. chloramine T (2% w/v) 0.02 g. streptomycin sulfate (200 ug/ml) a trace of Tween 80 add water until the total volume is 100 ml
135 Clearing and Staining of Roots to see VAM Structures: Basic Method
Methods: 1. Clearing the roots. Wash roots preserved with FAA in water. Heat at 90º C. for generally 1 hour in 20% KOH in the hood. 2. Pour off KOH solution and rinse with 3 changes of tap water or until the rinse is clear. 3. Acidify with 10% HCL and soak for about 4-5 minutes, then pour off. Do not rinse. 4. Cover roots with a 0.01% acid fuchsin-lactic acid solution, which will stain the VAM structures. Heat at 90º C. for 10 to 60 minutes. 5. Rinse off the solution with distilled water and mount the roots on a slide either in water (temporary mount) or in a permanent medium (See Powell and Bagyaraj, 1984). 6. Examine the slides with the compound microscope for VAM structures including spores, arbuscules and hyphae. 7. Allow slides to set and harden on warming tray for two days. Lead weights may be used to flatten root segments. Variations:
1. Length of time in KOH (#1): Some roots such as cattails are destroyed if heated for more than 20 minutes (Tang, 1986); however, most grass roots need to be processed for 1 hour. 2. Type of stain: Other stains work just as well as acid fuchsin; however, one advantage of acid fuchsin is that no destaining is necessary. Trypan blue is a commonly used stain (0.01%) but destaining for 1 hour or more using the lactic acid solution without stain is required. 3. Heating or no heating: KOH (#1). The roots do not have to be heated in the base instead they can be left at room temperature. The minimum time which my lab has used is 12 hours, but shorter periods may be possible. Heavily suberized roots may need to be left for several days or even a week. 4. Phenol: Do not add to lactic acid solution. It is not necessary and is potentially carinogenic. Materials and Equipment: Water bath heat to 90º C. Test tubes and holders Tweezers Slides Coverslips Compound microscopes
136 Preparation of Solutions: 20% KOH 10% HCL Lactic Acid Solution - 875 ml lactic acid 63 ml glycerin 63 ml distilled water Add 0.1 g. acid fuchsin to 1 liter lactic acid solution Discussion: Details of most of the procedures I have discussed are elaborated on in the excellent reference book, "Methods and Principles of Mycorrhizal Research," edited by Schenck (1982). Chapter three by Daniels and Skipper (1982) and chapter four by Kormanik and McGraw (1982) are particularly useful. Kormanik and McGraw discuss the problems with Phillips and Hayman (1970) procedure for clearing and staining roots, and make recommendations for improving the method. Photographs of VAM structures, including spores, arbuscules, vesicles and hyphae are provided at the back of this reference (Schenck, 1982). If these structures are found within the roots of the plant, then the plant if mycorrhizal. Another useful general reference concerning VAM is the recent book by Powell and Bagyaraj (1984). Keys for spore identification are found in the books edited by Schenck (1982) and Powell and Bagyaraj (1984). References:
Daniels, B. A. and H. D. Skipper. 1982. Methods for the recovery and quantitative estimation of propagules from soil. In Methods and Principles of Mycorrhizal Research. Ed. N. C. Schenck. The American Phytopathological Society. pp. 29-36. Gerdemann, J. W. 1955. Relation of a large soil-borne spore to phytomycetous mycorrhizal infections. Mycologia 47. 619-632. Gerdemann, J. W. and T. H. Nicolson. 1963. Spores of mycorrhizal Endogone extracted from soil by wet sieving and decanting. Trans. Brit. Mycol. Soc. 46.235-244. Ianson, D. C. and M. F. Allen. 1986. The effects of soil texture on extraction of vesicular- arbuscular mycorrhizal fungal spores from arid site. Mycologia 78. 164-168. Kormanik, P. P. and A. C. McGraw. 1982. Quantification of Vesicular-arbuscular Mycorrhizae in Plant Roots. In Methods and Principles of Mycorrhizal Research. Ed. N. C. Schenck. The American Phytopathological Society. pp. 37-36. Phillips, J. M. and D. S. Hayman. 1970. Improved procedures for clearing roots and staining parasitic and vesicular-arbuscular mycorrhizal fungi for rapid assessment of infection. Trans. Brit. Mycol. Soc. 55: 158-161. Powell, C. L. and D. J. Bagyaraj. 1984. VA Mycorrhiza. CRC Press, Inc. Schenck, N. C. 1982. Methods and Principles of Mycorrhizal Research. The American Phytopathological Society. Tang, F. 1986. Colonization of Vesicular-arbuscular Mycorrhizae in Typha angustifolia. University of Minnesota. Masters thesis. Acknowledgement: Funding for the Typha-VAM Project has been supported by grants from the BioEnergy Coordinating Office (BECO) to I. Charvat. The techniques covered in this workshop have been developed as part of this project.
137
Chapter 8
Teaching Botany Through Inquiry
Gordon E. Uno
Department of Botany and Microbiology 770 Van Vleet Oval, Room 135 Norman, Oklahoma 7301 9
Gordon E. Uno received his B.A. in Biology with Education from the University of Colorado at Boulder, and his Ph.D. in Botany from the University of California at Berekely. He is currently Associate Professor of Botany at the University of Oklahoma where he teaches a large Introductory Botany course and Economic Botany. He has worked as a consultant for the biological Sciences Curriculum Study (BSCS), and is co-author of four editions of two different high school biology texts produced by the BSCS. His research interests include plant-animal interactions in the prairie, and the use of inquiry instruction in college biology courses.
139
TEACHING BOTANY THROUGH INQUIRY
INTRODUCTION
Introductory Botany is taught in two different formats at the
University of Oklahoma. The first format includes a traditional lecture for 250 students each semester with inquiry-oriented laboratories of 36. The second format, from which most of the following material is taken, is taught completely using inquiry with
48 students in each class. There are three or four inquiry sections each semester, and each has a faculty member or an advanced graduate student in charge. The inquiry sections meet four times a week for
50 minutes each day. In these, lecture, laboratory and discussion are combined, with emphasis on laboratory and discussion. Students are given all the laboratory equipment and materials needed for each day's lesson. See pages labeled Day 8 - Day 13 from the lab prep manual which outlines what students and instructors need each day.
Pages 27-36 are taken from the student's workbook which coincides with the activities that take place in class. Students are given reading assignments, but these and the workbook activities arc completed AFTER the students have participated in class. This insures that the "discovery" aspect of the inquiry method is maintained. Introductory Botany is not a content course but a concept course where students are trained to ask the right questions and to develop botanical/biological concepts on their own. Students are asked to learn facts, however, not as many are given as in a traditional lecture. Inquiry botany is designed for both majors and non-majors, although a majority of students are non-majors.
141 FIRST-DAY INQUIRY ACTIVITIES FOCUSING ON THE SCIENTIFIC METHOD
First-day activities focus on the "scientific method" which serves as the foundation for all inquiry activities throughout the semester. Although not all instructors or scientists agree on the existence of the scientific method or on its elements, I emphasize the following steps:
1) observation of a phenomenon
2) asking questions about the cause ot the phenomenon
3) hypothesis formation
4) prediction about what will happen if contributing factors to
the phenomenon are changed
5) experimentation to test the hypothesis
6) collection and analysis of the data
7) rejection or support of the hypothesis
Using The Scientific Method.
To the students: Suppose you are an objective observer watching an empty theater before the movie begins. You observe a young woman enter the theater by herself and sit in one of the seats. A few minutes later, a young man enters the theater and, without saying anything to the woman, sits down beside her. These are your OBSERVATIONS. Now, based on these observations, what
QUESTIONS might you ask about this situation? Students give a variety of responses including: "Do they know each other?" "Is that the best chair for viewing the movie?" "Is he trying to pick her up?"
We can choose any one of these questions and study it by first forming a hypothesis and then testing the hypothesis. To FORM A
142 HYPOTHESIS, we can convert a question (for example, the first one asked) into a statement--The two people in the movie theater know each other. It this statement is true, then we can manipulate the situation and PREDICT what will happen. For instance, we could move one of the two people to a different seat (you can't ask the people questions) and see if the other person follows. If these two people do know each other as we have stated in our hypothesis, then the other person should follow. We can now conduct the EXPERIMENT and move one of the people to a different seat. If we move the woman, and the man follows, we need to write this down as our DATA. If we continue to move the woman from seat to seat and the man follows each time, then our HYPOTHESIS IS SUPPORTED. But, that does not prove our hypothesis to be correct--we cannot prove a hypothesis to be true, we can only support or reject it.
Students who are asked why the results of the experiment do not prove our hypothesis to be true provide responses such as: "He could be a masher following her", "He could be interested in her perfume and not her," "He might be trying to find the best chair in the theater," etc. This leads to further hypothesis formation and testing.
Review the scientific method with the students and point out the steps that were used in the discussion about the movie theater.
Followup First-day Activity.
This second activity also revolves around the scientific method. Duplicate and cut up the sheet of figures (Grouping
Geometric Figures) and place one set of figures, except Q and R, into an envelope. Students are divided into teams of 2,3 or 4
143 students. Each team gets one envelope with 17 figures in it (A-P
and S). The students should introduce themselves and then be given
the following task: "Arrange the 17 f igures into groups. You can
have as many groups as your team wants, but you need to write down a statement describing why you have placed the figures in each particular group."
On the board, teams write the letters in each of their groups,
but not the descriptions of the groups. The instructor chooses two
interesting groupings (for example, the groupings of Team 1 and Team
4) and divides the class in half. One half of the class arranges
their figures like those of Team 1 while the other half arranges
their figures like Team 4. (Team 1 arranges the figures like Team 4 and Team 4 arranges like Team 1.) These are OBSERVATIONS. Students now try to decide why Team 1 (or 4, whichever is appropriate) arranged the figures as they did, and they write down the reasons.
This is the HYPOTHESIS. The instructor now hands out figures Q and
R and the students TEST THE HYPOTHESIS by placing Q and R into the groups they think Team 1 (or 4) would.
In a general discussion, students state why they think Team 1
(or 4) arranged the figures into their groups and into which groups they think Team 1 would place Q and R. Team 1 now reveals whether the discussion has been accurate. If so, the HYPOTHESIS IS
SUPPORTED. If not, the hypothesis is not supported and must be rejected. Frequently, Q and R will be placed in the right groups, but for reasons other than those Team 1 (or 4) originally stated.
In this case, the hypothesis is supported, but the hypothesis is
144 incorrect. This is an important point for discussion. Summarize emphasizing the steps used in the scientific method.
INQUIRY LABORATORIES ON LEAF ANATOMY AND PHOTOS YNTHES IS
Each student/workshop participant is given a prepared microscope slide of a privet leaf cross section and asked to make observations about the arrangement of cells. Students make a sketch of the leaf, and examples are drawn on the board. The instructor points out that it appears the lower group of cells within the leaf
(spongy parenchyma) appears to be loosely packed and asks how we might be able to determine if this is true. Several responses are given including: "Observe leaf sections cut at different angles to see it cells are widely spaced in these views as well," "Stain cells so they more easily seen." If no one comes up with a desired response, the in structor asks a "prompting question"--"How do we know that air is in a balloon?" A response--"By squeezing or pushing the air out." A similar demonstration is done by placing a fresh privet leaf in very hot water. When this is done, bubbles form at the bottom of the leaf but not at the top. The instructor asks, "What information does this provide?" Several responses include: "The cells in the bottom halt of the leaf are loosely packed and surrounded by a gas that escapes from the leaf."
Secondly, "There are holes in the bottom of the leaf through which gases can escape (or enter)." Finally, "There are holes in the bottom of the leaf but not the top." The instructor asks the students to look at the privet leaf cross section again and see if these holes (stomata) can be found. A discussion is held about the holes and the gases that might enter or leave.
145 Through a series of experiments and demonstrations, the
students now develop for themselves the formula for photosynthesis.
The instructor first shows the clearing and staining process that
reveals the presence of starch within a leaf (hot water, hot
alcohol, iodine). A discussion is held after each of the following
experiments/demonstrations is conducted.
Experiment 1: Plant 1 kept in the light for 48 hours; Plant 2 kept
in the dark for the same length of time. A leaf taken from Plant 1
stains darkly, the leaf from Plant 2 does not stain. Conclusion:
Light is necessary for starch formation.
Experiment 2: Leaf 1 is placed in a bowl of glucose solution for 48
hours; Leaf 2 is placed in a bowl of water for the same length of
time. Both leaves are kept in the dark. Leaf 1 stains darkly, Leaf
2 does not stain. Conclusion: Light is not necessary for the
formation of starch, but glucose is. Secondly, light is necessary
for the formation of glucose.
Experiment 3: We know glucose has carbon in it, but from what source does the carbon come? Plant 1 is kept under a sealed glass
bell jar with a container of potassium hydroxide which removes all of the carbon dioxide from the jar; Plant 2 is kept under a sealed glass bell jar but with no potassium hydroxide. A leaf from Plant 2 stains darkly, but a leaf from Plant 1 does not stain. Conclusion:
Carbon dioxide is necessary for the formation of starch (and glucose), and the carbon in the glucose comes from carbon dioxide.
Experiment 4: How does the carbon dioxide enter the leaf? Plant 1 has half of the top of each leaf coated with vaseline and Plant 2 has half of the bottom of each leaf coated with vaseline. Leaves
146 from Plant 1 stain darkly throughout the leaf while leaves from
Plant 2 stain darkly only in the half of the leaf where there was no
vaseline. Conclusion: carbon dioxide (or some other substance
necessary for photosynthesis) enters the leaf from the bottom of the
leaf. Instructor should relate this information back to the study
of leaf anatomy and previous experiments.
Demonstration 5: Is chlorophyll necessary for photosynthesis? Both
albino and normal green corn seedlings are tested for starch. Only
the green corn seedlings are shown to produce starch. Conclusion:
chlorophyll is necessary for photosynthesis. Discuss importance and
role of chlorophyll.
Demonstration 6: Separation of spinach leaf pigments using paper
chromatography. Demonstrates that chlorophyll is not the only
pigment in leaves. Discuss role of all pigments in a plant.
Demonstration 7: From what source does the hydrogen in glucose
come? Discussion is held about an experiment involving heavy water
and a green plant in sunlight. This demonstrates that water is
necessary for the production of glucose, and that the hydrogen in glucose comes from water.
Experiment 8: What else, beside glucose, is produced in
photosynthesis? Elodea Plant #1 is placed in water under light;
Elodea Plant #2 is placed in water in the dark. Funnels arc
inverted over both of the containers and a test tube is placed at
the end of each funnel. Test tubes inverted over the containers
holding the elodea are tested with a glowing splint. The splint glows brighter when placed in the test tube over Elodea #1 and does
147 not glow brighter when placed in the tube over Elodea #2.
Conclusion: oxygen gas is produced in photosynthesis.
A discussion is held to re-emphasize major points and to
recreate the formula for photosynthesis. Four to five class periods
are used for the development of the formula for and the discussion
of photosynthesis.
KEYS TO INQUIRY
1. Lead students to discover the biological and botanical concepts
for themselves rather than just telling them. For instance:
Traditional Approach Inquiry Approach
A fruit is a ripened ovary What do the following
containing seeds. structures have in
common?
squash, tomato, apple,
watermelon, peanut, etc.
Make a list of responses
and then discuss.
Note: Inquiry takes longer than the traditional approach.
2. Incorporate elements of the scientific method into your course as often as possible. Most importantly, allow students to make
148 observations, to form hypotheses, and to test hypotheses through experiments or demonstrations.
Note: Active discussions are essential to the success of this method of instruction.
3. Use a variety of approaches for each concept you teach--experiments, demonstrations, living materials, discussion of data already collected, films, and slides. You don't have to do an experiment to do inquiry.
4. It is possible to adapt traditional laboratories or demonstrations so that students can discover concepts on their own.
Rule of thumb--Are you telling students information that they could figure out for themselves? Can you give students just enough information about a concept so they can work out the rest?
5. Start asking questions on the first day and encourage discussion from the very beginning.
6. Think up questions you will ask and responses students nay give
BEFORE you get into the classroom. By doing this, you can control the direction of the discussion. This requires that you think through the day's activities. You will find that discussions often stray from your intended path, but as long the discussion is productive, that's okay. Your prepared questions will allow you to get back on track whenever you need to.
7. Ask only one question at a time. Do not ask "yes or no" type questions or any question that has only one correct answer--such as
"Does this leaf stain positively for starch?" You should ask open-ended questions to which there are several responses.
149 8. Don't answer your own questions. Wait for answers from
students--even if it seems like an hour before you hear a response.
If no students respond, rephrase the question.
9. Accept all responses made by students--don't be negative as long
as the students are making a legitimate attempt at contributing to
the discussion.
Try to get several answers to each question you ask.
10. Summarize every day. With inquiry, students sometimes
complain that they don't know what notes to take in class. These
students often seem to be the ones who are passive, who don't
participate, and who just want answers.
11. Try to get everyone involved!
12. If you meet every day, do not use inquiry ALL OF THE TIME.
Students will become annoyed at being constantly asked
questions--mix your teaching methods.
150 Grouping Geometric Figures
Cut apart the figures on this page. Then sort the figures into groups.
Reprinted with permission from Biological Sciences: Patterns and Processes, third edition, 1986, p. T5, Kendall/Hunt Publishing Company and the Biological Sciences Curriculum Study (BSCS). 151 -DAY -8 TISSUE CONCEPT AND EXTERNAL TISSUE OF LEAVES
Each Student: --1. Microscope -Each -Two Students: 1. Dissecting tray 2. One potted Graptopetalum plant with large number of leaves (for epidermis) 3. Finger bowl with 12 wilted leaves of wandering jew (wilted for two days) picked from bed in moderately bright light 4. Reagent rack with dropper bottle of water Demonstration Table: 1. Same as for students Greenhouse Order: 1. 25 Graptopetalum plants 2. Many (300) wilted wandering jew leaves Grocery Order. 1. Nothing
152 -DAY -9 LEAF STRUCTURE USING PRIVET Each Student: --1. Microscope 2. Slide set series "A" (slide #4-A, X sec. privet leaf and #3-A, epidermis of wandering jew for review purposes if necessary)
Each Two Students: --1. Dissecting tray 2. Dropper of water in wooden rack 3. Fresh microtomed X sec. of privet leaf, in small Syracuse watch glass of water. They should be cut about 60-90 microns thick. The basal half of the leaf will give much better midveins. (Keep in cold room and make sure they are thin enough. Check with microscope.) Put 10-15 young privet leaves between 2 potato or carrot slices, then microtome. Use potato for microtoming. Use sliding microtome in Room 19. 4. Bottle of water containing two 10" leaf bearing stems of privet. (Wide mouthed 1 pint bottle) (for macroscopic observation of the leaf)
Demonstration Table: 1. Same as for students 2. To demonstrate the release of gas from the inter-cellular spaces of the leaf: a. Potted coleus and geranium plant with leaves containing air in the intercellular space. (2 plants each) b. Electric hot plate c. Glass 400 ml beaker of hot water. (Start to heat before the 9:30 a.m. section) d. Pair of long (10" or 12") forceps e. Asbestos gloves 3. Rotary microtome (in Room 156) 4. Leaf skeletons to demonstrate veins 5. Leaf model
Assistants: 1. Check the supply of privet sections each hour and be sure for thickness during the time they are microtomed. Do not cut too thick.
Greenhouse Order: 1. Fifty 10" leaf hearing privet stem tips 2. TWOeach coleus and geranium Grocery Order: 1. 1 potato Prep Notes: 1. Set up coleus leaf blades in sucrose and in water 48 hours ahead of testing. (for DAY 11). 2. Set up vaseline coleus 48 hours ahead of testing (for DAY 12). 3. Prepare large glob of gluten without starch (for DAY 10). Soak large glob of gluten in beaker of water for three or four days. Periodically knead and rinse gluten under running water during this time. (Gluten is prepared by adding water to powered gluten to form a dough-like consistency.) 153 Day 10 CHEMISTRY AND THE CHEMISTRY OF CARBOHYDRATES
Each Student: --1. Microscope 2. Slide set series "A" Each Two Students. 1. Dissecting tray 2. Reagent block with a. Dropper bottle of water b. Dropper bottle of iodine (Sufficient concentration to give a positive test for starch grains) 3. Carbohydrates in small bottles: potato starch, glucose, fructose, and sucrose. (Samples will be taken from the bottles of sugar and starch, therefore, it will be necessary to check and refill many of the bottles before they are stored after their use) (to test for solubility). 4. Microtome sections of a potato tuber in a Syracuse watch glass of water. The sections should be between 100 and 200 microns thick. These sections should be kept in a refrigerator over-night. Be sure to check the sections occasionally as they are being cut to ascertain their fitness for study. Use sliding microtome in Rm. 19 - add a few drops of lemon juice to preserve slices. 5. Corn, olive, coconut, cottonseed oils 6. Gluten in finger bowl of water
Demonstration Table: 1. To demonstrate a test for starch in potato tubers: a. One potato tuber per section b. One sharp knife for slicing potatoes c. One 125 cc. dropper reagent bottle of iodine solution d. Two clean petri plates per section 2. Corn or oat seedlings 6" to 8" tall that have been growing in: a. the dark in white quartz sand b. the light in white quartz sand (to show that, initially, seedling gets energy from seed.) 3. Large glob of gluten without starch (test a small glob for starch-should be starch-negative) 4. Beaker with water
Greenhouse Order: 1. Corn or oat seedlings growing in quartz sand and seeds of each. Grocery Order: 1. Large potatoes - 2 per section Prep Notes: 1. Set up carbon dioxide coleus one day ahead of testing (for DAY 12) 2. Prepare gas free water (for DAY 12) by boiling H20 and let it sit there for days.
154 DAY 11 FOOD MANUFACTURE IN PLANTS --PHOTOSYNTHESIS Each Student: --1. Nothing
Demonstration Table: 1. To demonstrate the effect of light on starch formation: a. Varigated coleus plant that has been growing in light, displayed on the demonstration table under a light with leaves which show a good starch test. b. Varigated coleus plant that has been growing in continuous darkness until the leaves are completely starch-free (5 to 10 days in a dark box) c. Plant with leaf mask 2. To demonstrate the synthesis of starch in the dark by destarched coleus leaves floating on a sugar solution a. Support blade of leaf on rim of petri dish with petiole extending into a 4% sucrose solution. Place under dark box for 48 hours before needed for class. These leaves should be starch positive on the veins only. b. Control for above except that water is used in place of a sugar solution. The leaves should be starch negative 3. Equipment for testing for the presence of starch in leaves. (For procedural demonstration only). The following list of materials will be referred to here after as "Leaf Starch Testing Equipment": a. Hot plate and extra hot plate with asbestos pads b. 2 beakers each: 600,400, 250, 250 ml c. One 1 liter bottle of 95% ethyl alcohol d. One 500 cc bottle of iodine solution e One 500 cc. bottle for waste alcohol f. One pair of long forceps (10" stainless steel) g. One box of starch free filter paper, to fit petri plates h. Six petri dishes i. Enamel tray of water and one enamel tray - empty j. Three or four standard lantern slide cover glasses on which to float leaves. k. Box of soda crackers (Matzos). Large square of cheese-cloth l. 1 asbestos glove 4. Labeled dentonstrations of leaves from coleus plants listed above (tested for starch): a. Untreated b. Boiled in water c. Boiled in water and alcohol d. Boiled in water and alcohol and treated with iodine solution. The above demonstrations should be displayed on moist filted paper in a covered petri dish. A complete set "a-d" should be made for one plant above, but only a "d" demo should be prepared for all other tests. 5. Instructors may run tests on the plants on the light and dark plants. Be sure plants have enough leaves for all sections, or see that replacement plants are available. DAY 11 Continued on Next Page
155 -DAY- 11 Continued Assistants 1. Return all demonstrations to the instructor's desk and be sure to replenish the supply of 95% ethyl alcohol on hot plate.
Greenhouse Order: 1. Seven 9" high varigated coleus plants (in 3" pots) from bright light. These should be strongly starch positive. 2. As above except from the dark box and should be starch negative. 3. One plant with leaf mask (see Dr. Uno). Grocery Order: 1. See grocery order, Day 25
156 -DAY -12 FOOD MANUFACTURE IN PLANTS: Photosynthesis Cont'd Student's Table: 1. Nothing
Demonstration Table: 1. To demonstrate the liberation of oxygen by Elodea in bright light: a. Place six 6" stem tips of Elodea beneath a glass funnel with the cut end directed toward the funnel stem. Place the funnel and Elodea in a 1 gallon battery jar filled with gas free water. Support the rim of the funnel from the bottom of the battery jar with three #9 rubber stoppers. The level of the water should cover the tip of the funnel. Place this apparatus in bright light such as a group of flourescent lights. Oxygen should be liberated from the stem tips and will collect in the test tube. Fill elodea collection tube with O2 from tank. Replenish O2 for each section. b. Control: Same set-up as above except that it will be displayed under a small dark box. No oxygen will collect in this test tube if prepared properly. Check test tube each hour. 2. To demonstrate that carbon dioxide enters the leaf mainly through the stomates: a. Starch-free variegated coleus plants with one half the underside of each leaf coated with vaseline. Place plant in bright light. (A series of fluorescent lights should be used as they will not melt the vaseline.) They should remain under this light for at least 48 hours or until the uncoated side becomes strongly starch positive (black). Remove vaseline with toluene. 3. To demonstrate that carbon dioxide is necessary for photosynthesis: a. Expose three starch-free variegated coleus plants in the following environments (label such that students can't see the labels). 1. Variegated coleus plant from the dark box exposed to bright light until the leaves give a positive test for starch. 2. Variegated coleus plant from the dark box sealed under a bell jar from which all of the carbon dioxide has been removed by a strong solution of calcium hydroxide. This solution may be placed in a beaker containing a fluff of gauze which will increase the reacting surfaces. Caution: Do not allow the calcium hydroxide to overflow onto the table. The above plants should be starch negative. 3. Same as above except that instead of removing the carbon dioxide with calcium hydroxide a 5% carbon dioxide gas should be added to the bell jar. Include a small beaker of water with a gauze wick as #2 above for a control. This plant will become strongly starch positive b. A kip carbon dioxide generator with attached wash bottle (Don't seal the tube or top of the CO2 generator) c. The bell jars used in the above experiments should have openings in the top to permit removal of the leaves with long forceps. 4. Starch testing equipment from previous lesson 5. Labeled demonstrations of coleus plants listed above, tested for the presence of starch. Only one demo per plant is necessary 6. Box of salt-free crackers (Matzos, from preceding day) 7. Wooden splints and matches
DAY 12 Continued on Next Page 157 DAY 12 -Continued-
Greenhouse Order: 1. More coleus plants as needed from the light and dark 2. 12 6" elodea stem tips
Grocery Order: 1. Nothing Prep Notes: 1. Soak 30 wheat seeds overnight for pyrogallic demo. (for DAY 16) 2. To renew CO2 generator: Open the valve until all acid has drained into the bottom and middle sections. Carefully lift off the top section and empty out acid. Replace the acid with a 50% HCl and 50% water solution. (NOTE: add acid to water and work under a hood). Fill to one third of middle section. Don't fill up to stopper on side. Carefully spoon in about half a cup of marble chips (CaC03) into the middle chamber. Replace the top section and make sure it is well sealed. After a few seconds, close the valve to stop the reaction.
158 -DAY -13 FOOD MANUFACTURE: Photosynthesis and Chlorophyll
Each Two Students. --1. Potted, variegated Coleus plant 2. To demonstrate separation-of pigments by paper chromatography a. Dropper bottle of spinach chlorophyll ground in acetone and filtered b. 500 ml bottle containing 1 inch of solvent Pet Ether fitted with cork containing hook which holds a 2 inch wide strip of chromatography paper long enough to touch surface of solvent. c. I capillary tube
Demonstration Table: 1. To demonstrate that photosynthesis occurs in chlorenchyma: a. One variegated coleus plant, under a light, with starch in the leaves. This plant should have a large non-green central area. b. One purple Coleus plant with starch in the leaves, under a light. Leave the lights on in the room overnight. 2. Labeled demonstrations of leaves from Coleus plants listed above tested for a) untreated, b) boiled in water, c) boiled in water and alcohol, d) boiled in water, alcohol, and treated with iodine solution. This demonstration should be displayed on moist filter paper in a petri dish. 3. Leaf starch-testing equipment. (See DAY 11) 4. A labeled demonstration of the chlorophyll separation in a large cylinder of sufficient size to be seen at the back of the room. 5. One small flat of albino corn seedlings from the light (in green house). 6. One or two plants with variegated leaves with some leaf areas non-green.
Assistants: 1 Return all demonstration material to its proper place on the demonstration table. Check the supply of all the reagents and if low be sure to replenish the supply. Greenhouse Order: 1. 28 8"-12" variegated Coleus plants 2. 2 purple Coleus plants 3. 3-4 different plants with variegated leaves, some areas non-green. 4. 1 flat of albino corn Grocery Order: 1 1 pound frozen spinach. 2. 1 pint karo (obtain from stockroom) 3. 2 packages yeast (obtained from the micro-stock room) Prep Notes: 1. Start pyrogallic acid demo (see DAY 16) 2. Set up week old yeast culture (for DAY 17). Add 1 pint karo and 2 packages of yeast to 3000 ml of water. 3. Start 4 cups of wheat seeds germinating about three or four days before needed (for DAY 16). Germination is somewhat faster under a light.
DAY 13 Continued on Next Page
159 DAY 13 -Continued-
4. The chlorophyll separation in a cylinder is kept in the refrigerator. When this needs renewed apply the following instructions: a. Fresh chlorophyll extract. Can be prepared by boiling green leaves (frozen spinach) in 80-85% alcohol. b. Cylinder of 50% chlorophyll extract and 50% petroleum ether. (demonstrated separation of chlorophylls from carotinoids). Add petroleum ether to chlorophyll extract and shake. Remove stopper carefully to release pressure buildup. Add a little water and continue shaking. When shaking stops two layers will form. The upper layer should be green and the lower one yellow.
160 LEAF CELLS AND LEAF TISSUES
1. Strip off the skin, or epidermis, of a sedum leaf. What is the color of the epidermis? How many cell layers thick is the epidermis? Based on its location, what might be the function of the epidermis?
Why do you think the tissue beneath the epidermis is called chlorenchyma?
2. With the microscope, examine pieces of the upper and lower epidermis of a zebrina leaf. How many different kinds of cells do you see in the epidermis?
Label the kinds of cells in the top diagram on the next page.
STOMATA
The stomate (plural, stomata) is a small opening between each pair of guard cells. Are the walls of the guard cells of uniform thickness on all sides of the cell? How might this help the guard cells open or close the stomate?
What are the small green bodies in the guard cells? Are stomata present in both the lower and upper epiderm is of the leaves of the zebrina plant? Approximately how many stomata are present in each square millimeter of epidermis from the lower leaf surface? How many stomata in the upper epidermis? (The visible portion of the leaf under the low power of the microscope is approximately one square millimeter in area.) The number of stomata per square millimeter of leaf surface varies in different parts of a leaf surface and also in leaves from different plants as shown by the following table. Average number of stomata per square millimeter of leaf surface Plant In upper epidermis In lower epidermis
Elodea 0 0 Coleus 0 141 Nasturtium 0 130 Black Walnut 0 46 0 Kidney Bean 40 176 Sunflower 85 156 Oats 25 23 Corn 7 0 88 Tomato 12 130 Water Lily 46 0 0
161 162 3. What do you think is the significance of the different number of stomata in the top and bottom leaf epidermis?
4. In what kind of environment do you think you would find plants with very few stomata on their leaves?
5. Examine the cross section of a living leaf with the unaided eye and with a microscope. Compare your observations with a specially prepared slide of a similar leaf. Do all the cells in the leaf look the same? Describe the differences you see.
How many groups of related cells can you identify? Each group of similar cells is called a tissue. These may be similar in position (as in epidermis) and similar in function (as in phloem). Locate the following tissues in your leaf section and then give its function.
Name of tissue Function
Epidermis
Chlorenchy ma
Palisade parenchyma
Spongy parenchyma
Phloem
Xylem
Identify the epidermis, palisade parenchyma layer, the spongy parenchyma layer, the phloem and xylem of the bottom diagram on the previous page.
163 PLANT FOOD
I. Three types of sugar are used by plants as food. They are glucose (grape sugar); fructose (fruit sugar); and sucrose (cane sugar). Starch is also used by plants, but is made up of many linked sugar molecules. Place a small amount of starch in your hand and taste it. In the same way taste glucose, sucrose, and fructose, in this order. Then taste glucose again. List the three sugars and starch in the order of their sweetness. 1) 2) 3) 4) Place a very small amount of glucose, fructose, sucrose, and starch on different corners of a slide, keeping track of where you placed each. Examine them all under the low power of the microscope. Add a drop of water to each and observe again. What happened to the glucose? What happened to the fructose? What happened to the sucrose? Were they destroyed? Are they still sugar? How could you separate them again from the water? What happened to the starch?
Why do you think so?
Add a drop of iodine solution to each of the sugars and starch on your slide, observe, and record the results for 1) glucose 2) fructose 3) sucrose 4) starch 2. Food classification. Name some of the foods you eat, and based on their main chemical content, place them in one of the following categories. Carbohydrates:
Fats:
Proteins:
Which of the above categories of food can provide energy for you? What are some of the uses you make of the energy and molecules that are chemically bound in food?
3. Food observation within plant cells. Place an extremely thin fresh section of potato in a drop of water and observe using the low-power objective of the microscope. Add a drop of iodine solution from the edge of the cover glass and observe again. Record your observations.
Does the potato contain starch? How do you know?
How can you account for empty cells?
164 FOOD SYNTHESIS IN PLANTS: PHOTOSYNTHESIS a. Why do molds grow luxuriantly on bread in a dark box, but green plants die in good soil in a dark box? b. Why do the non-green parts of green plants die if the green parts are removed as rapidly as they begin to grow? c. Why do albino seedlings in good soil die about three weeks after germination, while green seedlings continue to live? d. Why do underground organs of plants in good soil die if the tops above ground are kept from developing?
These and many more questions will be answered by this section about photosynthesis.
TEST FOR STARCH Using a leaf from a coleus plant kept in the sunlight, the instructor will: 1. place the leaf in boiling water to remove the purple pigment (anthocyanin). 2. place the leaf in hot alcohol to remove the chlorophyll. 3. treat the leaf with iodine to determine if starch is present.
The instructor will repeat the previous three steps on a leaf from a coleus plant which has been in a dark room for several days.
Record your observations about the leaves taken from the two different conditions, and the effect of light on the presence of starch.
LIGHT
DARK
What conclusion can you draw about the effect of light on the presence of starch?
A leaf from a coleus plant kept in the dark has been immersed in a 4% solution of glucose. Similarly a leaf from the same plant has been placed in water without sugar. During the treatment, both leaves were kept in the dark for 24 to 48 hours. Using the test for starch above, check for the presence of starch, record your observations about the amount of starch present in the leaves under the two conditions.
4% SUGAR WATER
WATER
What is your conclusion about sugar and the formation of starch?
165 Is light necessary for the change of sugar to starch in leaves? Is light necessary for the synthesis of sugar in leaves? Which of these two processes do you think is photosynthesis? How can you determine if your data and conclusions about coleus leaves are true for leaves in general? Under what conditions might starch become changed to sugar in a plant cell?
To demonstrate the liberation of gas during photosynthesis, a short-necked funnel has been placed over some Elodea in a jar and a test tube inverted over the funnel. The whole apparatus has been placed in the light. The gas that replaces the water in the test tube will be tested with a glowing splint. Draw a diagram of the experimental apparatus and explain what is happening, i.e., for what are you testing with the glowing splint and how was it produced?
To determine if any of the components of air are necessary for photosynthesis, we will use two plants that first have been placed in a dark room for several days to insure that no starch remains in the leaves. One of these plants was placed under a bell jar along with a bottle of KOH which removes carbon dioxide from the air. The bell jar was then sealed to a glass plate with vaseline. The other plant was left uncovered and both plants were placed in the light. After two days the leaves of both plants are tested for starch using the previously described starch test. Record your observations for this experiment. COVERED
Are there any of the components of air necessary for photosynthesis? Explain your answer.
What gas do you think is required for photosynthesis? Into what plant molecule is this gas incorporated ?
Does gas enter the leaf, mainly through the stomata or directly through the epidermal cells? To answer this question, a coleus plant is used because nearly all its stomata are in the lower epidermis of the leaves. The lower surface of some of the leaves are covered with vaseline and the plant placed in the light for several days. Now we will remove the vaseline from the leaf and test for starch. Record your observations. LEAVES WITH VASELINE
166 LEAVES WITHOUT VASELINE
Does gas enter the leaf through the leaf epidermis or through the stomata?
How can you tell?
Write the chemical equation for photosynthesis, indicating both the materials used and the products produced.
PAPER CHROMATOGRAPHY
Basic research in modern cell biology helps to determine the metabolic processes that occur, their significance, and how each activity may affect other activities. Learning more about metabolism frequently requires the isolation and purification of chemicals extracted from cells. Paper chromatography is one technique useful in separating the components of cell extracts. In paper chromatography, chemicals are carried on filter paper by a solvent. Because different chemicals have different solubilities in the solvent, the chemicals will separate from each other as they travel on the filter paper. To demonstrate chromatography of a leaf extract:
1. Filter paper is trimmed so a strip will fit into a test tube without bending. 2. Application of spinach leaf extract is made about 3 centimeters from the end of the strip and allowed to dry. 3. Step Two is repeated several times, with application of the extract to the same place each time, until a large amount of extract has been applied to a small spot on the paper. 4. Test tubes are filled with a small amount of solvent and supported in an upright position. 5. The strip of paper is inserted into the prepared test tube with the treated end down so it dips into the solvent but the spot remains above the solvent. The test tube is stoppered, making sure not to move the test tube during the experiment. 6. Make observations and a diagram showing the results, showing which pigments are present and which ones travel fastest.
How many different bands of color do you see from the spinach leaf extract? What were their colors? What molecules in the leaf are producing these colors and what is their role in the plant?
167 The relationship of photosynthesis to light intensity is stated in the table below. The first column lists the percentages of light intensity compared to that on a clear July day. The other vertical columns in the table represent the relative rates of photosynthesis in the designated plants when exposed to the differing light intensities. Plot the data on the graph below, drawing a curve for each of the five plants.
Light Intensity: Relative rates of photosynthesis expressed in milligrams of percentages of carbon dioxide used per 0.5 square meters of leaf area per . full sunlight hour. Spinach Potato Nasturtium Pine Fern
2 5.5 0.3 0.0 0.0 0.0 5 7.1 2.5 1.1 0.1 0.8 10 8.0 5.0 2.1 0.6 1.6 20 8.9 6.4 3.7 1.2 2.2 30 9.6 7.0 4.6 1.6 2.3 40 10.0 7.3 5.1 1.8 2.3 50 10.4 7.6 5.4 2.1 2.2 60 10.7 7.7 5.6 2.3 2.0 70 10.8 7.8 5.7 2.5 1.9 80 11.0 7.8 5.8 2.7 1.8 90 11.0 7.8 5.8 2.8 1.7 100 11.1 7.8 5.8 2.9 1.6
Light intensity: Percentage of Full Sunlight
168 -
In general, how do changes in light intensity affect the rate of photosynthesis?
Which plant would grow best in full sunlight? In dim light? Would the pine or the fern do better in full sunlight? In dim light?
The rate of photosynthesis in a potato in relation to temperature and the concentration of carbon dioxide in the air is represented in the table below. Draw a curve showing the rate of photosynthesis for each concentration of carbon dioxide.
Temperature in degrees 5 10 15 20 25 30 35 40 45
Rate of photosynthesis at 0.03 percent C02 2.1 6.5 8.6 13.8 8.8 8.0 5.7 2.9 0.0 Rate of photosynthesis at 1.22 percent CO2 2.9 12.4 20.2 28.1 33.7 56.0 48.3 35.6 8.0
In general, how do changes in temperature affect photosynthesis?
How does an increase in C02 concentration affect photosynthesis?
169 SUMMARY
1. What kind of "food" is used by plants and how is it used?
2. Explain, in your own words, the process of photosynthesis.
3. What factors in the environment affect photosynthesis?
4. How do humans make use of the knowledge of the relationships between food manufacture in the plant, growth and development of a plant, and environmental conditions?
5. Scientists grew plants using water molecules that contained a radioactive isotope of oxygen, 180. Here is the resulting formula: 16 18 16 18 C 02 + 2H2 0 ------(CH2 0) + 02 Can you determine which is the molecule from the left side of the equation that yields the oxygen gas on the right side? Explain your reasoning.
6. What are the main products of the light-dependent reactions (light reactions) of photosynthesis?
7. What are the main products of the light-independent reactions (dark reactions) of photosynthesis?
170 Chapter 9
A Laboratory Introduction to DNA Restriction Analysis*
David A. Micklos 1 and Greg A. Freyer 2
1 DNA Learning Center of Cold Spring Harbor Laboratory 2 Columbia University College of Physicians and Surgeons
David Micklos is director of the DNA Learning Center of Cold Spring Harbor Laboratory. He received a bachelor's degree in biology from Salisbury State College, Maryland and a master's degree in science journalism from the University of Maryland. Prior to joining Cold Spring Harbor Laboratory, he worked as a survey researcher for Hill & Knowlton, Inc. and a Peace Corps Volunteer in Botswana Africa. He has taught at both the high school and college levels. Greg Freyer is an assistant professor in the radiation oncology department of Columbia University College of Physicians and Surgeons. He received a bachelor's degree in biology from the University of Cincinnati and a doctorate in biochemistry from the University of Missouri. He did postdoctoral work at Cold Spring Harbor Laboratory and was associate researcher at Memorial Sloan Kettering Cancer Center. In 1985, they founded the DNA Literacy Program as a means to train instructors to better teach recombinant-DNA technology at the college and advanced high school levels. The most visible aspect of the program has been the Vector DNA Science Workshop that has now been taken be more than 600 high school and college educators from New York to California, and from Wisconsin to Alabama. The week-long course gives teachers practical experience with recombinant-DNA techniques and addresses the practical aspect of implementing DNA science labs. The program is now based in the DNA Learning Center, the nation's first "museum" devoted entirely to biotechnology education. For more information about the DNA Literacy Program and the Vector DNA Science Workshops, contact the DNA Learning Center at (516) 367-7240.
*This Workshop was sponsored by Fotodyne, Inc., New Berlin, WI 53151 and presented by William R. Gette, Senior Scientist, Fotodyne, Inc.
171
A LABORATORY INTRODUCTION TO DNA RESTRICTION ANALYSIS*
David A. Micklos DNA Learning Center of Cold Spring Harbor Laboratory
Greg A. Freyer Columbia University College of Physicians and Surgeons
PURPOSE This laboratory demonstrates that the DNA molecule can be precisely manipulated and that it behaves as predicted by the structure determined by James Watson and Francis Crick in 1953. The genotypic analysis performed in this experiment is the basis for DNA fingerprinting and the starting point in the construction of recombinant-DNA molecules.
In this protocol, purified DNA from the bacteriophage lambda is digested with the restriction endonucleases EcoRI, BamHI, and HindIII. Each enzyme has five or more recognition sites in lambda DNA and, therefore, produces six or more restriction fragments. The digested DNA is loaded into a 1% agarose gel. An electric field applied across the gel separates the DNA fragments according to size. The bands of DNA within the gel are made visible by staining with either ethidium bromide or methylene blue.
BACKGROUND INFORMATION Restriction Endonucleases
Restriction endonucleases, or restriction enzymes, are used as molecular scalpels to cut DNA in a precise and predictable manner. They are members of the class of nucleases that display the general property of breaking the phosphodiester bonds that link adjacent nucleotides in DNA and RNA molecules. Endonucleases cleave nucleic acids at internal positions, while exonucleases progressively digest from the ends of nucleic acid molecules.
There are three major classes of restriction endonucleases. Type-I and Type-III enzymes have both restriction (cutting) and modification (methylating) activity. Both types cut at sites some distance from their recognition sequences; ATP is required to provide energy for movement of the enzyme along the DNA molecule from recognition to cleavage site.
*Abridged from DNA Science: A First Laboratory Course in Recombinant-DNA Technology, now in preparation. Copyright 1988 Cold Spring Harbor Laboratory and Carolina Biological Supply Company. Presented as a workshop by William Gette.
173 Type-II restriction enzymes are for several reasons most useful to molecular biologists: 1) Each has only restriction activity; modification activity is carried by a separate enzyme. 2) Each cuts in a predictable and consistent manner at a site within or adjacent to the recognition sequence. 3) They do not require ATP for cutting activity. Today, more than 900 Type-II enzymes have been isolated from a variety of prokaryotic organisms. Enzymes have been identified that recognize 130 different nucleotide sequences; over 70 types are commercially available. To avoid confusion, restriction endonucleases are named according to the following nomenclature: 1) The first letter is the initial letter of the genus name of the organism from which the enzyme is isolated. 2) The second and third letters are the initial letters of the organism's species name. (Since they are derived from scientific names, the first three letters of the endonuclease name are italicized.) 3) A fourth letter, if any, indicates a particular strain of organism. 4) According to most recent nomenclature, a Roman numeral indicates the order in which enzymes, isolated fromthe same organism and strain, are eluted from a chromatography column. However, an earlier nomenclature, in which Roman numerals indicate the order of discovery, is still used. Each restriction endonuclease scans along a DNA molecule, stopping only when it recognizes a specific sequence of nucleotides. Most restriction enzymes recognize a four- or six-nucleotide sequence. Assuming that the four component nucleotides (A,C,T,G)are distributed randomly within a DNA molecule, then any four-nucleotide recognition site will occur, on average, every 256 nucleotides (4 x 4 x 4 x 4). A six-nucleotide recognition site is likely to occur every 4,096 nucleotides (4 x 4 x 4 x 4 x 4 x 4). Many restriction enzymes have recognition sites that are composed of symmetrical, or palindromic, nucleotide sequences. This means that the recognition sequence read forward on one DNA strand is identical to the sequence read backward on its complementary strand Put another way, the 5'-to-3' sequence is identical on each DNA strand. In a general sense, the terms 5' and 3' refer to either end of a single DNA strand. Specifically, they designate carbon atoms on opposite sides of the deoxyribose ring that are joined to form a single strand DNA polymer. The 5' carbon is linked, through ester bonds with an intervening phosphate, to the 3' carbon of the adjacent nucleotide. By convention, the nucleotide sequence is "read from the 5' end to the 3' end of the DNA strand. The situation is confused in duplex DNA, where the strands are "antiparallel;" that is, 5'-to-3' reads in opposite directions on the two complementary strands. To carry out a restriction reaction, solubilized DNA is incubated at 37ºC with one or more endonucleases. The reaction takes place in a buffered solution that provides salt conditions necessary for optimum enzyme activity. Within or very near the recognition site, the enzyme catalyzes a hydrolysis reaction that uses water to break a specific phosphodiester linkage on each strand of the DNA helix. Two DNA fragments are produced, each with a phosphate group at the 5' end and a hydroxyl group at the 3' end. Some endonucleases, such as HindII, cut cleanly through the DNA helix by cleaving both complementary strands at the same nucleotide position, typically in the center of the recognition site. These enzymes leave flush- or blunt-ended fragments.
174 Other endonucleases cleave each strand off-center in the recognition site, at positions two to four nucleotides apart. This creates fragments with exposed ends of short, single- stranded sequences. Various enzymes leave single-stranded "overhangs" on either the 5' or 3' ends of the DNA fragments. EcoRI, BamHI, and HindIII, for example, each leave 5' overhangs of four nucleotides.
Single-stranded overhangs, also called cohesive or "sticky" ends, are extremely useful in making recombinant-DNA molecules. These exposed nucleotides serve as templates for realignment, allowing complementary nucleotides to hydrogen bond to one another. A given restriction enzyme cuts all DNA in exactly the same fashion, regardless of whether the source is a bacterium, a plant, or a human being. Thus, any sticky-ended fragment can be recombined with any other fragment generated by the same restriction enzyme. For each restriction Type-II endonuclease there is a corresponding modifying enzyme that blocks restriction activity by methylating DNA within the recognition sequence. The protruding methyl group presumably prevents binding by interfering with the close molecular interaction between the restriction enzyme and its recognition site. EcoRI methylase, for example, adds a methyl group to the second adenine residue within the EcoRI recognition site.
Nomenclature, Recognition Sequences, and Cutting Action of EcoRI, BamHI, and HindIII
EcoRI E = genus Escherichia Recognizes G A A T T C co = species coli R = strain RY13 I = first endonuclease isolated
BamHI B = genus Bacillus Recognizes A T c c am = species amyloliquifaciens H = strain H I = first endonuclease isolated
HindIII H = genus Haemophilus Recognizes G C T T in = species influenzae TTC d = strain Rd III = third endonuclease isolated
Agarose Gel Electrophoresis
Electrophoresis means literally "to carry with electricity." The method takes advantage of the fact that, as an organic acid, DNA is negatively charged. DNA owes its acidity to phosphate groups that alternate with deoxyribose to form the rails of the double helix. In solution, hydrogen ions are liberated from hydroxyl groups, leaving negatively- charged oxygen ions that radiate from phosphates on the outside of the DNA molecule. When placed in an electrical field, DNA molecules are attracted toward the positive pole and repelled from the negative pole. Sorting of differently-sized molecules is achieved by electrophoresing DNA fragments through a gel-like matrix that acts as a molecular sieve through which smaller molecules can move more quickly than larger ones. The distance moved by a DNA fragment is
175 inversely proportional to the logarithm of its molecular weight. Thus, in a given period of time, smaller restriction fragments migrate relatively far from the origin compared to larger fragments. A research team at Cold Spring Harbor Laboratory, led by Joseph Sambrook, introduced two important refinements to electrophoresis that made possible rapid analysis of DNA restriction fragments.
First, they used a matrix composed of agarose, a highly-purified form of agar. Agarose efficiently separates larger DNA fragments ranging in size from 100 to more than 50,000 nucleotides. DNA fragments in different size ranges are separated by adjusting the agarose concentration. A low concentration, down to 0.3% produces a looser gel that separates larger fragments. A high concentration, up to 2%, produces a stiffer gel that resolves small fragments.
Second, they used a fluorescent dye, ethidium bromide, to stain DNA bands in agarose gels. Following a brief staining step, the fragment pattern is viewed directly under ultraviolet light. As little as 10 nanograms (0.01 micrograms) of DNA can be detected in a band. (An earlier method was laborious and required the use of radio-labeling.)
Currently used electrophoresis methods are essentially identical to those published by the Cold Spring Harbor team in 1973. Molten agarose is poured into a casting tray in which a "comb" is suspended. As it cools, the agarose hardens to form a jello-like substance consisting of a dense network of cross-linked molecules. The solidified gel slab is immersed in a chamber filled with buffer solution, which contains ions needed to conduct electricity. The comb is removed, leaving wells into which DNA samples are loaded. Just prior to loading, the digested DNA is mixed with a "loading dye" consisting of sucrose and one or more visible dyes. The dense sucrose solution weights the DNA sample, helping it to sink into the well when loaded. The negatively-charged dye molecules do not interact with the DNA, but migrate independently toward the positive pole. For example, the commonly used marker bromophenol blue migrates at a rate equivalent to a DNA fragment of approximately 300 nucleotides (in a 1% gel). Thus, the visible movement of the dye allows one to monitor the relative migration of the unseen DNA bands.
Current supplied through electrodes at either end of the chamber creates an electrical field across the gel. The negatively-charged DNA fragments move from the wells into the gel, migrating through the pores in the matrix toward the positive pole of the electrical field.
Following electrophoresis, the gel is soaked in a dilute solution of ethidium bromide. The stain diffuses throughout the gel, becoming concentrated in regions where it binds to DNA fragments. (Alternately, ethidium bromide is incorporated into the gel and electrophoresis buffer prior to electrophoresis.) A planar group of the ethidium bromide molecule intercalates between the stacked nucleotides of the DNA helix, staining DNA bands in the gel.
The stained gel is then exposed to medium wavelength ultraviolet (UV) light. The DNA/ethidium bromide complex strongly absorbs UV light at 300 nm, retains some of the energy, and reemits visible light at 590 nm. Thus, the stained restriction fragments appear as fluorescent orange bands in the gel, when exposed to ultraviolet light.
It is important to understand that a band of DNA seen in a gel is not a single DNA molecule. Rather, the band is a collection of millions of DNA molecules, all of the same nucleotide length.
176 REAGENTS, SUPPLIES , AND EQUIPMENT
0.1 mg/ml lambda DNA 0-10 µl micropipetor + tips restriction enzymes electrophoresis box EcoRI power supply BamHI 1.5 ml tubes HindIII permanent marker 2X restriction buffer test tube rack loading dye 37ºC incubator distilled water microfuge (optional) 1% agarose solution transilluminator/camera 1X TBE buffer (optional) 1µg/ml ethidium bromide solution parafilm (optional) (or 0.025% methylene blue solution)
PRELAB PREPARATION
1) Aliquot for each lab group: 20 µl 0.1 µg/µl lambda DNA (store on ice) 25 µl 2x restriction buffer (store on ice) 2 µl each of BamHI, EcoRI, and HindIII (store on ice) distilled water loading dye (NOTE: The volumes and concentrations of DNA have been optimized for ethidium bromide staining, which is the most rapid and sensitive method. If you prefer to use methylene blue staining, increase DNA concentration to 0.3-0.4 µg/µl Volumes used remain as stated.)
2) Prepare 1% agarose solution (40-50 ml per lab group). Keep agarose liquid in a hot water bath (about 55ºC) throughout lab period.
3) Prepare 1X Tris-Borate-EDTA (TBE) buffer for electrophoresis (400-500 ml per lab group). (NOTE: The volumes of agarose solution and TBE buffer needed vary according to electrophoresis apparatus used. The volumes used here are based on typical "mini-gel" systems.)
4) Set water bath to 37ºC. (NOTE: An aquarium heater makes a very good constant-temperature bath.)
177 LABORATORY PROTOCOL I. Prepare Restriction Digest (30 minutes, including incubation)
1) Label four 1.5 ml tubes, in which you will perform restriction reactions: B BamHI E EcoRI H HindIII -- no enzyme 2) Set up matrix to use as checklist as reagents are added to each reaction:
DNA 2X buffer BamHI EcoRI HindIII H20
3) Collect reagents, and set in test tube rack on lab bench.
4) Set micropipetor to 4 µl, and add DNA to each reaction tube. Touch pipet tip to side of reaction tube, as near bottom as possible, to create capillary action to pull solution out of tip. (It is not necessary to change tips when adding same reagent.)
5) Buffer should be added to reaction tubes before enzymes. Use fresh tip to add 5 µl restriction buffer to clean spot on each reaction tube. (Same tip may be used for all tubes, provided tip is not touched to solution already in tubes.) 6) Use fresh tips to add 1 µl EcoRI, BamHI, and HindIII to appropriate tubes. 7) Use fresh tip to add 1 µl deionized water to -- Tube. 8) Pool and mix reagents:
-- by sharply tapping tube bottom on lab bench.
-- with a short, several-second pulse in microfuge. (Make sure tubes are placed in balanced configuration in rotor.) 9) Place reaction tubes in 37ºC water bath. 10) Let reactions incubate for minimum 20 minutes. (Reactions can be incubated for longer periods of time. After several hours, enzymes lose activity and reaction stops.) ** OPTIONAL STOP POINT: Following incubation, reactions can be frozen at -10 to -20°C until ready to continue. Thaw reaction before continuing to Step III-1.
178 II. Cast 1% Agarose Gel (20 minutes) Seal ends of gel-casting tray and insert comb. Place gelcasting tray out of the way on lab bench, so that agarose poured in next step can set undisturbed. (Gel is cast directly in box in some electrophoresis apparatuses.)
Carefully pour enough agarose solution, at approximately 55ºC, into casting tray to fill to depth of about 5 mm. (Liquid agarose container should be just cool enough to hold comfortably in the hand.) Gel should cover only about 2-4 mm height of comb teeth. Large bubbles or solid debris can be moved to sides or ends of tray, while gel is still liquid, using toothpick or pipet tip. Do not move or jar casting tray whileagarose is solidifiing. As it polymerizes -- in about 10-15 minutes -- agarose will change from clear to cloudy. Touch corner of agarose away from comb to test if gel has solidified.
When agarose is set, unseal ends of casting tray. Place tray on platform of gel box, so that comb is at negative end. Fill box with 1X Tris-Borate-EDTA (TBE) buffer to level that just covers entire surface of gel. Too much buffer will channel current over top rather than through gel, increasing time required to separate DNA. (TBE buffer can be used several times; do not discard until told to do so.) Gently remove comb, taking care not to rip gel. (Buffer solution helps lubricate comb teeth.) Make certain that holes (sample wells) left by comb are completely submerged. If "dimples" are noticed around wells, slowly add buffer until they disappear.
OPTIONAL STOP POINT: Cover electrophoresis tank and save gel until ready to continue. Gel will remain in good condition for at least several days, so long as it is completely submerged in buffer. (Gels can be cast and sealed in a small volume of buffer inside zip-lock plastic or "seal-a-meal" bags. Before sealing, squeeze out as much air as possible. Include enough buffer to completely surround gel, and refrigerate.)
III. Load Gel and Electrophorese (40-60 minutes)
Add 1-2 µl loading dye to each sample: -- Set up four individual droplets of loading dye (1-2 µl each) on small square of parafilm or wax paper. Withdraw contents from reaction tube, and mix with a loading dye droplet by pipeting in and out. Immediately load dye mixture according to Step 3. Repeat successively, with clean tip, for each reaction.
-- Add 1-2 µl loading dye to each reaction tube. Mix by tapping on lab bench, pipeting in and out, or spinning tubes for short several-second pulse in microfuge. (Make sure tubes are placed in a balanced configuration in rotor.) This mixes loading dye with reactants.
179 3) Use micropipetor to load entire contents of each reaction tube into separate well in gel, as shown in diagram below. (A piece of dark construction paper beneath the gel box or a piece of black electrical tape affixed to gel box bottom makes wells easily visible.) - Steady pipet over well using two hands. - Be careful to expel any air in micropipet tip end before loading gel. (If air bubble forms "cap" over well, DNA/loading dye will flow into buffer around edges of well.) - Be careful not to punch tip of pipet through bottom of gel. - Gently depress pipet plunger to slowly expel sample into appropriate well. If tip is centered over well, reaction solution will sink to bottom of well. 4) Close top of tank, and connect electrical leads anode to anode (red-red) and cathode to cathode (black-black). Make sure both electrodes are connected to same channel of power supply.
5) Set power supply at 100-150 volts, and turn unit on. (Alternately, set power source on lower voltage and let gel run for several hours. Check to monitor progress of bromophenol blue band.)
6) Ammeter should register approximately 50-100 milliamps. (Current through two gels is double that for single gel at same voltage.) This confirms that current is flowing through gel. If no current is detected, check connections, and try again.
7) Shortly after current is applied, loading dye can be seen moving through gel toward positive side of electrophoresis apparatus. It will appear as a blue band -- eventually resolving into two bands of color. The faster-moving, purplish band is the dye bromophenol blue. The slower-moving, aqua band is xylene cyanol. Bromophenol blue migrates through gel at same rate as a DNA fragment approximately 300 basepairs long.
8) Electrophorese for 20-60 minutes. Good separation will have occurred when the bromophenol blue band has moved 4-8 cm from wells. CAUTION: If gel is allowed to run for too long, dye and DNA samples will electrophorese out end of gel into buffer.
9) Turn off power, and disconnect leads. 10) Carefully remove gel from electrophoresis chamber, and place in disposable weigh boat or other shallow tray. ** OPTIONAL STOP POINT: Gel can be stored in a zip-lock plastic bag for viewing and photographing the next day. However, over time DNA will diffuse through gel, and bands become indistinct.
11) Stain and view gel using one of following methods.
180 IVA. Stain with Ethidium Bromide and View (10-15 minutes) INSTRUCTOR: Before using this method, familiarize yourself with Safety Note: Responsible Handling of Ethidium Bromide at the end of this laboratory. Your instructor may complete staining. RUBBER GLOVES MUST BE WORN DURING STAINING, PHOTOGRAPHY, AND CLEANUP.
Flood gel with ethidium bromide solution (1 µg/ml).
Allow to stain for 5-10 minutes. (Staining time depends on thickness of gel. Gentle rocking of staining tray will aid diffusion of ethidium bromide into gel.)
Use funnel to drain off ethidium bromide solution into storage container for reuse. Rinse gel and tray under running tap water to remove excess ethidium bromide solution. (Chlorine in water will inactivate traces of ethidium bromide.)
If desired, gels can be destained in tap or distilled water for 5 or more minutes to remove background ethidium bromide.
OPTIONAL STOP POINT: Staining intensifies dramatically if rinsed gels set overnight. Cover or wrap gel in plastic wrap to prevent dessication. (Weigh boats containing stained and rinsed gels simply can be stacked together overnight.) View under ultraviolet transilluminator or other W source. CAUTION: Ultraviolet light can damage your eyes. Never look at unshielded W light source with naked eyes. Only view through filter or safety glasses that absorb harmful wavelengths.
IVB. Stain with Methylene Blue and View (30+ minutes)
Flood gel with 0.025% methylene blue. Allow to stain for approximately 15 minutes. Retrieve stain and rinse gel in tap water. Let gel soak for several minutes in several changes of fresh water. DNA bands will become increasingly distinct as gel destains. (Heat and agitation facilitate rapid destaining.)
OPTIONAL STOP POINT: For best results, continue to destain overnight in small volume of water. Cover container to retard evaporation. (Gel may destain too much if left overnight in large volume of water.) View gel over light box.
181 V. Photograph(10 -15 minutes) 1) FOR W PHOTOGRAPHY.. - Use Polaroid high-speed film Type 667 (ASA 3,000). - Set camera aperture to f/8 and shutter speed to B (bulb). Make a 2-4 second exposure by depressing shutter release for desired length of time. (Shutter remains open as long as release is depressed.) FOR WHITE-LIGHT PHOTOGRAPHY: - For Polaroid film Type 667, set aperture to f/8 and shutter speed to 1/125 second. Exposure times vary according to mass of DNA in lanes, level of staining, degree of background staining, thickness of gel, and density of filter. Experiment to determine best exposure. When possible, stop lens down (higher f/ number) to increase depth of field and sharpness of bands.
2) Place left hand firmly on top of camera to steady. Firmly grasp small white tab, and pull straight out from camera. This causes a large yellow tab to appear. 3) Grip yellow tab in center, and in one steady motion, pull straight out from camera. This starts development.
4) Let develop for recommended time -- 45 seconds at room temperature. Do not disturb print while developing. 5) After full development time has elapsed, separate print from negative. Begin peeling back at end nearest yellow tab. CAUTION: Avoid getting caustic developing jelly on skin or clothes. If jelly gets on skin, flush immediately with plenty of water.
DISCUSSION
1) Troubleshooting Electrophoresis What electrophoresis effect would occur: a) If gel box is filled with water instead of TBE buffer? b) If water is used to prepare gel instead of TBE buffer? What effect would be observed in the stained bands of DNA in an agarose gel:
C) If casting tray is moved or jarred while agarose is solidifying in step II.2? d) If gel is run at very high voltage?
e) If a large air bubble or clump is allowed to set in agarose?
f) If too much DNA is loaded in a lane?
182 2) Two small restriction fragments of nearly the same size appear as a single band, even when sample is run to very end of the gel. What could be done to resolve the fragments? Why would it work?
3) Shown below is an ideal fragment pattern for a digest of lambda DNA with BamHI, EcoRI, and HindIII. Using the ideal gel as a reference, how many fragments can be identified on the photograph of gel obtained in this laboratory?
Account for differences in separation and band intensity between the experimental gel and the ideal gel.
B E H -- BamHI EcoRI HindIII No enzvme
183 4) Linear DNA fragments migrate at rates inversely proportional to the log10 of their molecular weights. For simplicity's sake, basepair length is substituted for molecular weight. a) Construct a matrix like the one shown below, which gives the size in basepairs (bp) of lambda DNA fragments generated by BamHI and HindIII digests: BamHI EcoRI HindIII bp dis bp dis bp dis 16,841 *23,130
*Pair appears as single band. **Does not appear on this gel. b) Use ideal gel shown above. For each of the three digests, carefully measure distance (in mm) each fragment migrated from the origin. Measure from front edge of well to front edge of each band. Enter distances into matrix. c) Match sizes of known BamHI and HindIII fragments with bands that appear on the ideal digest. Label each band with kilobasepair (kbp) size. For example 16,841 bp equals 16.8 kbp. d) Set up semi-log graph paper with distance migrated as the X (arithmetic) axis and log basepair length as the Y (logarithmic) axis. Then plot distance migrated vs. basepair length for each BamHI and HindIII fragment. (Alternately, calculate log bp and plot directly on the Y axis of standard graph paper. To finish problem, compute anti-logs in step g.) e) Connect data points with a best-fit line. f) Locate on X axis the distance migrated by one EcoRI fragment. Using a ruler, draw a vertical line from this point to its intersection with the best-fit data line.
g) Now extend a horizontal from this point to the Y axis. This gives the basepair size of this EcoRI fragment. h) Repeat steps f and g for each EcoRI fragment.
For Further Experimentation 5) Design and carry out a series of experiments to study the kinetics of a restriction reaction. Determine approximate percentage of digested DNA at various time points. Repeat experiments with several enzyme dilutions and several DNA dilutions. In each case, at what time point does reaction appear complete?
6) Design and test an assay to determine the relative stability of BamHI, EcoRI, and HindIII at room temperature.
184 185 SAFETY NOTE: RESPONSIBLE HANDLING OF ETHIDIUM BROMIDE
Like many natural and man-made substances, ethidium bromide is a mutagen and cancer suspect agent. Therefore, this protocol has been written to limit its use to a single step that can be performed by the instructor in a controlled area. With responsible handling, the dilute ethidium bromide solution used in this experiment poses minimal risk.
The greatest risk is the posssibility of inhaling ethidium. bromide powder when mixing a 5 mg/ml stock solution. Therefore, we suggest purchasing ready-mixed stock solution from a supplier.
The stock solution is then diluted by the instructor to make a staining solution with a final concentration of 1 µg/ml. This is near the threshold of detection of ethidium bromide in mutagenicity assays -- approximately 0.5 µg/ml. Experimental data suggest that treatment with 5,000 ppm available chlorine (10% bleach solution) inactivates 95 percent or more of ethidium bromide in solution. Thus, treatment of fresh staining solution should bring ethidium bromide concentration to a maximum of 0.05 µg/ml- well below the threshold of detection. Following bleach treatment, ethidium bromide concentration in stained gels and in reused staining solution is considerably less. a) Always wear gloves when working with ethidium bromide solutions and stained gels. b) Limit ethidium bromide use to a restricted area located next to a sink with running water.
C) Following gel staining, use funnel to retrieve as much of the ethidium bromide solution for reuse or disposal as outlined below. d) Following lab, wipe down camera, transilluminator, and staining area with 10% bleach solution.
e) Flood stained gels with 10% bleach solution and let stand for 15 or more minutes. Drain, and collect stained gels in plastic bag. Discard in regular trash.
f) To discard staining solution, mix with equal part 10% bleach solution. Let stand for one or more hours, and discard down drain.
186 Chapter 10
Resource Partitioning in Potentially Competing Insect Taxa
John A. Haarstad
Cedar Creek Natural History Area University of Minnesota 2660 Fawn Lake Dr. N.E. Bethel, MN 55005
John A. Haarstad received his B.A. degree (biology) from Carleton College. His M.S. (ecology) and Ph.D. (entomology) degrees are from the University of Minnesota. He is currently a Post-doctoral Research Associate at Cedar Creek Natural History Area, Bethel, Minnesota where he is conducting ecological research on various insect communities, and working on the Station's reference insect collection.
187
RESOURCE PARTITIONING IN POTENTIALLY COMPETING INSECT TAXA
The idea that many biotic communities are competitively structured has both supporters and critics (Salt, 1983). Competitive structuring implies that interspecific competition is or has been an important force in determining which species are present in an area as well as their relative abundances and resource use patterns. If such communities exist it seems reasonable that there should be a limit to the similarity of coexisting species (Hutchinson, 1959). Studies on patterns of resource partitioning in coexisting, potentially competing, taxa provide real world information on how similar coexisting species are. However, it is important to bear in mind that interspecific competition may be relatively unimportant in many communities. There are a variety of factors that may play a role in determining which species are present in a community, how abundant each is, and the niche each occupies. Such factors include geographic barriers and the opportunity for colonization, abiotic factors such as soil and climate and physical disturbance, and other biotic factors including resource supply, predators, pathogens, and parasites. Thus it becomes important to select communities that are likely candidates for competitive structuring. Members of parasitoid and folivorous insect communities may find themselves on a coevolutionary mobius strip of defense and counter-attack where interspecific competition is relatively unimportant (Price, 1980; Lawton and Strong, 1981). In other communities competition may have been important historically but is no longer demonstrable because of past divergence (Connell, 1980). Ideally, one should select a set of similar species sharing a resource that is apparently limited in supply. Their populations should show a relatively rapid response to any change in resource supply or manipulation of suspected competitor population sizes. OLD FIELD ANT COMMUNITIES Ants are a diverse group with roughly 660 North American species and are excellent candidates for studies of resource partitioning (Schoener, 1983). Some tropical species are leaf-cutters and many western species forage for seeds (Hansen, 1978), but most temperate species are generalist scavengers foraging for other arthropods or feeding at plant nectaries and tending honeydew secreting insects (Carroll and Janzen, 1973). Any habitat is likely to have upwards of six coexisting species (Post and Jeanne, 1982) and it becomes a challenge for the investigator to unravel their niche relationships (Lynch, et.al., 1980). A long term investigation would include species' distributions across habitats (i.e., which species are habitat generalists or specialists) and a comparison of their seasonal abundance patterns. In this exercise we will concentrate on analyzing the niche relationships of an old field ant community at a single point in time. Niche dimensions to be examined include space (distribution within a field), time (diurnal activity periods), and food (resource preference). PROCEDURES AND ANALYSIS The instructor should locate a potential study site and set out baited petri plates prior to the field exercise. Plates with ants should be collected and frozen so that students have time to acquaint themselves with the species to be encountered and acquire some practice at doing counts. When large numbers are attracted to the baits, it is sometimes necessary to count by tens or twenties. Students must learn not to dote over a plate and to be satisfied with a good approximation of the numbers present.
189 Establish a grid of petri plates (e.g., one hectare grid at 10 meter spacing = 100 petri plates). Early in the morning, bait each plate with a ca.. 10 gram mixture of honey and tuna (pancake syrup and catfood if under a tight budget). Groups of students then visit the grid periodically throughout the day and record weather conditions, surface temperature, and the species and numbers of each visiting each plate. In 20 of the spaces betwen grid points, place divided petri plates containing different types of baits in each quadrant (e.g., pure tuna, Grape Nuts cereal soaked in cooking oil, grapes, centimeter cubes of sponge soaked in honey). Students observing these plates will count the number of individuals of each species attracted to each bait type and also make note of any aggressive encounters or displacements that occur. Additional information that could be recorded are discovery time, recruitment rates, and differences in food handling. For example, are large items dismembered by many individuals of a small species or are they ignored and subsequently carried off by a larger species. 1) Prepare a map of the study area locating grid points and conspicuous patches of vegetation. Superimpose upon this map the ant species collected at each grid point. Are certain species associated with particular patches of vegetation? Do aggressively dominant species appear to be segregated spatially? 2) Calculate the extent of spatial overlap between each species using the formula: 2c/a+b where a = no. baits containing species a, b = no. baits containing sp. b, and c = number baits containing both spp. a and b. 3) Are the same species present at the baits throughout the day or is there a diurnal succession of species? If species change throughout the day what do you think might be responsible (e.g., innate activity patterns, physiological responses to ambient temperature, aggressive displacements)? How might you test these ideas?
4) Calculate the extent of temporal overlap between each species using the formula TO = 1 - .5 S |pit - pjt| where pit and pjt are the proportions of the ith and jth species active during a particular time interval (t). 5) Do the ant species found in your field differ in resource preference? Which prefer sugar baits, protein baits, or show no preference? Do they differ in the size of bait they can handle?
6) Calculate the extent of food overlap between each species using the formula FO = 1 - .5 S |pif - pjf| where pif and pjf and the proportions of the ith and jth species attracted to a particular type of food (f). 7) Calculate total overlap along the dimensions of space, time, and food by multiplying the 3 overlaps calculated above. Which species are most sim^_ilar (dissimilar) in their niches? Which species do you suspect are the most serious competitors? Does the extent of niche overlap necessarily reflect the intensity of interspecific competition? See Colwell and Futuyma, 1971. 8) Which are the dominant and which the passive species in your ant community? Does displacement occur via aggressive fighting (e.g. leg or antennal pulling) or does some chemical appear important? For example, do any species gaster-flag or show evidence of anal secretions? See Adams and Traniello, 1981. ADDITIONAL QUESTIONS AND SUGGESTIONS The work and analysis done previously has helped to define the realized niches of the ants in a particular old field community. Use of space, daily activity pattern, and food preference in the
190 absence of all other species would be a species' fundamental niche. Many of the following questions are designed to look for 'niche shifts' when some of the community elements have been altered. 1) Place baited petri plates along transects passing through various habitats and collect the ants visiting them. Are some of the ants observed in old fields also found in wooded areas? What new species are encountered? Which species have the broadest (narrowest) niches? How might you quantify 'niche breadth'? 2) Establish a grid of baited petri plates in another abandoned field. Are the same species present in similar abundances or is this ant community different? What might account for any differences in species composition? For example, is the field more xeric? If the two fields differ somewhat in the complement of ant species, is there any evidence of a temporal niche shift in a species found in both fields? For example, does it forage earlier (or later) in the day when another species is present? 3) Test whether any diurnal changes in species abundances are due to innate activity patterns or physiological responses to ambient temperature by shading a colony and bait with a plywood board. 4) Locate a bait site that is dominated by a particular species having a nearby colony, and isolate that colony from the bait by encircling it with aluminum flashing rimmed with Tanglefoot Do other species of ants now use the bait? 5) Compare food preference of an ant species at a site where it is the only one to utilize the baits and at sites where other species are present. Is there any evidence of it being displaced from more desirable resources by another species? 6) Crush several ants suspected of chemically repulsing other species in petri plates containing honey/tuna baits. After removing the dead ants place these and uncontaminated baits near the colony of another species of ant Are both types of baits utilized with equal rapidity? REFERENCES Adams, E.S. and J.F.A. Traniello (1981). Chemical interference competition by Monomorium minimum. Oecologia 51: 265-270. Carroll, C.R. and D.H. Janzen (1973). Ecology of foraging by ants. Ann. Rev. Ecology and Systematics 4: 231-257. Colwell, R.K. and D.J. Futuyma (1971). On the measurement of niche breadth and overlap. Ecology 52: 567-576. Connell, J.H. (1980). Diversity and the coevolution of competitors, or the ghost of competition past. Oikos 35: 131-138. Creighton, W.S. (1950). The ants of North America. Bull. Mus. Comp. Zool. Harvard 104: 1-585. Hansen, S.R. (1978). Resource utilization and coexistence of three species of Pogonomyrmex ants in an Upper Sonoran grassland community. Oecologia 35: 109-117.
191 Hutchinson, G.E. (1959). Homage to Santa Rosalia, or why are there so many kinds of animals? Amer. Naturalist 93: 145-159. Lawton, J.H. and D.R. Strong (1981). Community patterns and competition in folivorous insects. Amer. Naturalist 118: 317-338. Lynch, J.F., E.C. Balinsky, and S.G. Vail (1980). Foraging patterns in three sympamc species, Prenolepis imparis, Paratrechina melanderi and Aphaenogaster rudis. Ecol. Entomology 5: 353-371. Post, D.C. and R.L. Jeanne (1982) Rate of exploitation of arboreal baits by ants in an old-field habitat in Wisconsin. Amer. Midland Nat. 108:88-95. Price, P.W. (1980). Evolutionary biology of parasites. Princeton Univ. Press, Princeton, N.J. 237 pp. Salt, G.W. (editor) (1983). A round table on research in ecology and evolutionary biology. Amer. Naruralist 122: 583-705. Schoener, T.W. 0983). Field experiments on interspecific competition. Amer. Naturalist 122: 240-285.
192 Chapter 11
Supercooling and Freezing in Winter Dormant Animals
William D. Schmid
Department of Ecology and Behavorial Biology University of Minneosta 109 Zoology Building Minneapolis, MN 55455
William Schmid received his B.S. in Wildlife Management from the University of Minnesota (1959) and his Ph.D. in Zoology from the University of Minnesota (1962). He currently is a Professor in the Department of Ecology and Behavioral Biology at the University of Minnesota. His current research interests include studying the seasonal patterns of supercooling and production of cyroprotectant or antifreeze compounds in the red spider mite (Trombidium), and the role of white- footed mice (Peromyscus leucopus) as host for the tick vector (Ixodes dammini) which harbors the spirochete (Borellia burgdorferi) of Lyme disease. He has a 1987-88 National Research Council Research Fellowship, and will be a USA- USSR Interacademy exhange fellow, September 1 through December 15, 1987. He also received a Fulbright Lectureship, Novosibirsk State University, Academgorodok, USSR, March 1 through June 1, 1988.
193
Supercooling and Freezing in Winter Dormant Animals
William D. Schmid Department of Ecology and Behavioral Biology University of Minnesota Minneapolis, MN 55455
Winter ecology is an extensive field of investigation that includes meteorology, microclimatology, phenology, natural history, physiology and biochemistry. Many organisms have special morphological, physiological and behavioral adaptations which are necessary for survival in seasonally cold regions. Organisms that live in such regions have four general categories of adaptation to winter stresses: (1) die, (2) migrate, (3) remain active and (4) become dormant. Death, migration and winter activity are all adequate adaptations to the winter cold season, but it is winter inactivity or dormancy of many animals that has been the focus of research in my laboratory during the past decade. Animals in a state of winter torpor are in one of two groups: (1) they can survive freezing (frost tolerant), or (2) will die if ice forms within their bodies so that they develop antifreeze chemicals to depress their freezing points and deeply supercool (frost resistant).
A small land snail, Vallonia perspectiva, is an example of the latter case. The cooling and freezing curve for a single specimen is shown in Fig. 1 which illustrates the application of thermocouple thermometry. The phenology of supercooling by this species is shown in Fig. 2 where the winter specimens are seen to supercool to much lower temperatures than those of summer. On the other hand, if we measure supercooling points of frost tolerant animals it is usually found that internal ice formation occurs at higher temperatures during the winter. A summary of supercooling point measurements is given for a variety of Minnesota species in Table 1. In all but one case, Mordellistina unicolor, frost tolerance is associated with relatively warm supercooling temperatures; i.e., those species which can survive freezing usually promote internal ice formation during the winter season. The examples provided here illustrate applications of the thermometry techniques to be described now.
The goldenrod gallfly, Eurosta solidagensis, will be used to demonstrate the techniques of thermometry as applied to studies of supercooling and freezing in winter dormant organisms. The gallfly is distributed widely across the eastern United States and winters as a third instar larva in diapause within the ball galls of goldenrod (Solidago sp.) plants. Third instar larvae collected in early fall will typically supercool to -15 ºC., and do not survive freezing. However when these larvae are collected in midwinter they will supercool to -9 ºC., and survive freezing (frost tolerant). Ball galls can be collected in midwinter and stored in a freezer until needed for laboratory measurement of supercooling points of the resident larvae. Ladybird beetles, Hippodamia convergens, in winter diapause, obtained from commercial suppliers, can be used to the demonstrate deep supercooling (frost resistance) adaptation to winter cold exposure. It is important to keep in mind that new, original discoveries can be made with these methods by utilization of local organisms which have never been studied for their adaptations to winter survival.
The term "freezing" simply indicates the formation of ice: phase change from liquid water to solid ice as temperature drops. "Supercooling" on the other hand refers to a
195 -20 -20 ...... ! 0 5 10 15 20 25 30 MINUTES MONTHS
Fig. 1. Cooling and freezing Fig. 2. The annual cycle of curve for a terrestrial snail, supercooling points for Vallonia Vallonia perspecriva, collected perspecriva. The open circles from its wintering site in (o) represent active specimens January. 1981. Point (a) is the and the closed circles (e) supercooling point (-17.6 º C), represent inactive animals with the sudden rise in specimen epiphragms over the aperature temperature from a to b is due to of their shells. At no time exothermic release of heat by during the year could this snail internal ice formation. and survive if frozen. Stippled bars freezing continued from b to c. at the top of the graph represent The snail did not survive the coldest microclimate temper- freezing. After Schmid (1986). ature of each month through the winter season After Schmid (1986).
Table 1. Supercooling and Frost Tolerance of Some Minnesota Animals in the Minter
SPECIES S.C.P. (ºC) N Survival Minter Habitat Pnphi biam Rain ..pt&rionazu Mtnk Frog Aquatic Rm.2 pi* Leopard Frog Aquatic &mu .yzvotim Mood Frog Terrestrial By& vasieobr Grey TmFrog Terrestrial tryla chl-y.o.~eZi. Gmn Trcc Frog Terrestrial &La CrVCifm Sprlng Peeper Terrestrial Bufo ollmGaw knerlcan Toad Terrestri a1 Ambystma MsmL. Jefferson Salamander Terrestrial Snai 1s VaLtoniU paspectiva Land Snail Terrestrial Ccurtrocoptu Land Snall Terrestrial Insects Terrestrial UOldELZiUtoM WliCObr Gall Beetle Buwsta soZidag&. Goldenrod Gallfly Terrestri a1 ~chclvtesnivimluo Snow Flea Terrestrial PZatypsyZZa matoris Beaver Beetle Terrestrial Ticks tor vadabiZie Wood Tick -14.2 =1.6 15 0 Terrestri a1 Delmaemtor azbipictu. Uinter Tick -17.4 t2.5 9 0 Terrestri a1 ~desdamnini Deer TIck -16.3 '2.4 10 0 Terrestrial nltes 7hmbidim sp. Red Velvet Mlte -7.4 t.6 11 11 Terrestrial Spiders Dictynu sp. Spider -32.1 t.7 8 0 Terrestrial ustaph-Zippus sp. Jmping Spiders -23.8 t2.3 38 0 Terrestrlal
After Schmid ( 1986) 196 freezing temperature below what is expected from the solute (colligative effect) concentrations in the aqueous solution of organisms. It is the temperature of spontaneous ice formation seen in intact animals as the temperature is gradually lowered. For this reason surface contact thermometry is necessary. Invasive placement of temperature sensors might provide artificial sites for ice nucleation that are not present in the intact animal. Although both thermistors and thermocouples are adequate choices of temperature sensors, the latter will be described and used in this exercise. In addition to the specimen cooling system, measurement of supercooling and freezing requires a sensor, a detector (high input impedance microvoltmeter for amplification) and a recorder.
The COOLING SYSTEM consists of a liquid bath into which a test tube containing the specimen and sensor can be placed for removal of heat. In the simplest case, this might be a thermos filled with a solution of automotive antifreeze which has been left in a deep freeze overnight. A dewar flask filled with antifreeze solution or ethyl alcohol can be cooled by addition of chips of dry ice. For shallow cooling temperatures, a styrofoam chest filled with crushed ice and rock salt might be sufficient. Controlled cooling baths, programmed for specific cooling rates and temperatures are commercially available and usually more convenient than these simple systems; e.g., Neslab or Forma. The choice of cooling system will depend upon your specific needs and budget available for equipment expenditures.
The SENSOR for continuous measurement of specimen temperature is constructed of fine (.002 inch diameter = 3 mil) thermocouple wires. Type T or copper-constantan wires are readily available and easily fabricated into sensors for contact thermometry. If the reference junction of the thermocouple pair is held at zero degrees C. in a bath of chipped ice and distilled water, the thermoelectric voltage can be easily converted to temperature by values given in Table 2. For example, if a cooled specimen had an exotherm at -.668 mV, its supercooling point would convert to -17.6 degrees C.
Table 2. Thermoelectric voltage for copper-constantan (Type T) thermocouples.
DEG C 0 1 2 3 4 5 6 7 8 9 10 OEG C 0 0.000 0.039 0.078 0.117 0.156 0.195 0.234 0.273 0.312 0.351 0-391 0 10 0.391 0.630 0.470 0.510 0.549 0.589 0.629 0.669 0.709 0.749 0.789 10 20 0.789 0.830 0.870 0.911 0.951 0.992 1.032 1.073 1.114 1.155 1.196 20 30 1.196 1.237 1.279 1.320 1.361 1.403 1.614 1.486 1.528 1.569 1.611 3 0 40 1.611 1.653 1.695 1.738 1780 1.822 1.865 1.907 1.950 1.992 2.035 40
The fine wires of the sensor or measurement junction of the thermocouple are soldered to heavier (24 AWG) copper-constantan lead wires for hook-up to the microvoltmeter or amplifier. The heavier lead wires are also used to form the reference junction where temperature is constant and problems of thermal inertia are not important. The fine wires of the sensor junction are necessary because they
197 have very low heat capacity or thermal inertia necessary to detect the heat released (latent heat of fusion of water) when ice forms inside of a specimen at its supercooling point. Structural durability for the fine. 2 mil sensor wires can be improved by using wire with a teflon sheath for insulation: the teflon covering provides strength as well as insulation. The insulation can be burned from the end of the wires which can then be scraped clean and twisted together to form electrical contact for the sensor junction. For greater integrity of the sensor thermocouple junction, the twisted wires can be wiped with solder. This junction must be held in close contact with the specimen as it is cooled.
A 15 ml conical centrifuge tube with the fine wire sensor junction inside at the bottom of the cone is a very simple system into which a specimen can be placed for cooling and measurement of its supercooling point. A more reliable system, required for small specimens can be constructed from a piece of plastic rod. The rod is cut into halves along its longitudinal axis (through its diameter) and the thermocouple sensor wires are glued onto the flat cut surface. A specimen is positioned against the sensor junction and held in place with a foam rubber pad that is itself taped to the plastic rod. The rod is then placed in a test tube and lowered into the cooling bath. Variations on the construction of the sensor junction and specimen holder must be made to accommodate the organism under study and the cooling rates desired. For example, added insulation might be necessary to control the cooling rate in the absence of a programmed cooling bath.
The DETECTOR-RECORDER operate as a unit. The sensitivity and amplification of the detector and the sensitivity of the recorder are adjusted to give readout precision to accurately detect the thermoelectric voltage change at supercooling. Two detectors of thermoelectric voltage that I have used successfully are the Keithly 155 microvoltmeter and the Omega 2809 digital thermometer. Both instruments have the necessary high input impedance and output connections to a recorder. Both the Gould 105 and the Fisher Recordall 5000 have been satisfactory recording devices for continuous records of temperature change. It is certainly feasible to use other combinations of devices for detection and recording of thermoelectric voltages from a Type T thermocouple sensing system; e.g., an A-D card in a personal computer might be used to capture the record of thermoelectric voltage change in digital form for later printout to hard copy. The sensitivities of the detector and recorder can be adjusted to provide the most useful scale on final output of strip chart paper. If the minimum temperature records do not go below -25 degrees C., adjustment of sensitivities to give full scale recorder pen deflection of 0 to -1 mV will be easily translated to a scale of temperature. Likewise, a full scale pen deflection of 0 to -2 mV will accommodate cooling to about -55 degrees C. Such systems of sensor-detector- recorder with subzero cooling can be used to examine many different examples of organism-cold season adaptation seen in patterns of supercooling and freezing. The experimental cooling rates used are best defined by knowledge of normal variations in temperature which are experienced by organisms in nature (Figs. 3-5). A nominal cooling rate often used in experiments of supercooling of insects is -1 degree C. per minute. The effects of cooling rate upon measured supercooling point and upon freezing survival should always be checked as a source of experimental bias. Biochemical changes accompany the seasonal patterns of supercooling and some of these can be easily detected by relatively simple methods.
Paper chromatography will detect polyhydric alcohols which are often associated with seasonal development of freeze tolerance or deep supercooling. A single gallfly larva, after measurement of its supercooling point, is homogenized in .5 ml of cold 70% ethanol. This crude extract is centrifuged to settle tissue debris and the
198 -20 -45 5 12 34 5 67 8 9101112 JASONDJFMAMJ MARCH, 1987 MONTHS Fig. 4 Daily high and low Fig. 3 Long term weather station temperatures recorded at the records for the Twin Cities. The open University of Minnesota bars represent average monthly low Forestry and Biology Station. and high temperatures, while the lines are record highs and lows.
I November December January February March Fig. 5 Microclimate temperature variation recorded through the winter (1980-81) at the soil surface, beneath leaf litter, in Elm Creek Park Reserve, Hennepin County, Minnesota.
199 supernatant is saved. Aliquots of 5 uL are spotted onto Whatman No. 1 paper along with mixed standards (5 uL of .1 M) glycerol, sorbitol and trehalose. Ascending chromatograms are developed in a solvent system of n-butanol:pyridine:water = 6:4:3 for about seven hours. The chromatograms are dried and sprayed with periodate and iodine-starch as described by Somme (1964). The developed chromatogram is dried and sprayed with 0.01 M aqueous solution (.23 g/100 ml) of potassium periodate and dried again. Then it is sprayed with the iodine-starch solution (35% saturated sodium tetraborate, .3% potassium iodide, .9% boric acid and 2% soluble starch) and dried. Glycerol and sorbitol will appear as white spots on a dark blue background, while trehalose appears as a light gray smudge on the same background. Chromatographic separation (migration) is approximately proportional to molecular weight so that glycerol is closest to the solvent front and trehalose is farthest away. The amount of each compound can be estimated by the size (area) of each spot. It is necessary to run standards of different concentrations in order to estimate concentration by spot area. Thin layer chromatography and high performance liquid chromatography are obvious alternatives to this relatively crude method of analysis. Nuclear magnetic resonance is also being used to estimate the seasonal production of cryoprotectant and/or antifreeze biochemicals by dormant animals of seasonally cold regions.
The materials and methods described in this brief presentation can be used to illustrate adaptations for survival during cold seasons of the year in laboratory courses of animal and plant physiology as well as physiological ecology. However, students with interest, curiosity and scientific aptitude will be able to use this exercise as a guide for original research wherein their findings will provide us with new and useful information about the adaptations of organisms to their environment. I close this discussion by thanking three people who have helped me develop my interest and skill to examine problems of cold temperature adaptation: Wally Saatala, Bob Maxwell and Rick Lee.
200 Appendix 1: Supplies
Suppliers of general laboratory equipment: Fisher Scientific 50 Fadem Road Springfield, NJ 0708 1
Cole-Parmer Instrument Company 7425 North Oak Park Ave. Chicago, IL 60648
Bio Rad Laboratories 32nd and Griffin Richmond. CA
Supplier of low temperature controlled baths: Neslab Instruments, Inc. 871 Islington St. Portsmouth, NH 03801
Supplier of biochemical standards for paper chromatography: Sigma Chemical Co. P. O. Box 14508 St. Louis, MO 63178
Suppliers of thermometry equipment and supplies: Omega Engineering, Inc. One Omega Drive Box 4047 Stamford, CT 06907
Keithly Instruments Cleveland, OH 44139
California Fine Wire Co. Grover City, CA
Thermometries 808 U.S. Highway I Edison, NJ 08817
Supplier of specimens for supercooling study: Fountain's Sierra Bug Co. P.O. Box 114 Rough and Ready, CA 95975 Source of ladybird beetles, Hippodamia convergens, in diapause. About $10 per quart
There are certainly alternative sources to those above, but this listing gives you someplace to start. There is no intention of special endorsement for any of these suppliers.
201 Appendix 2: Thermocouples
Thermocouples generate a thermoelectric voltage whenever there is a temperature difference between the junctions of the two dissimilar metals of their construction. Thermoelectric describes the phenomenon whereby heat energy is transformed (transduced) to measureable electrical energy, and vice versa. Although the irreversible conversion of electrical current 2 into heat (i.e., heat = I R) comes under the category of thermoelectric, it is the reversible phenomenon that we are more concerned with in regard to measurement of environmental or biological temperatures. The Peltier effect: Junctions of dissimilar metals are heated or cooled depending upon the direction in which a current passes through them. This effect is proportional to the first power of current and not to I2 as in the case of irreversible heating of conductors. Battery I Heated _-______------(I)
The Thompson effect: An emf (voltage) will arise within a single conductor wherever a temperature gradient is present. The total Thompson effect along a conductor depends only upon the temperature difference between the two ends of the conductor. The Seebeck effect is the algebraic sum of the Peltier and Thompson effects, and it is this total effect that is useful to us in application of thermocouples to thermometry. An electric current and its emf are present in a series of two different metals if the two junctions are at different temperatures; e.g., copper and constantan, where constantan (Cn) is an alloy of 60% copper and 40% nickel. High input impedance mvoltmeter
--
Thermoelectric sensitivity, dV/dT (mV/ºC) of thermocouples made of the following materials with platinum (reference junction = OºC):
Antimony 47 Aluminum 3.5 Nichrome 25 Platinum 0.0 Iron 18.5 Nickel -15 Copper 6.5 Bismuth -72 Silver 6.5 Constantan -35
So the expected thermoelectric emf from copper-constantan (Cu-Cn) or the Type T thermocouple is: (+6.5) - (-35) = 41.5 mV/ºC). Table 2 provides a listing of thermoelectric voltages for the Type T thermocouples, and it can be seen that the emf per degree Celsius change is very close to the theoretical and quite linear over a wide temperature range. 202 BIBLIOGRAPHY
Asahina, E. 1969. Frost resistance in insects. Adv. lnsect Physiol., 6:1-49. Baust, J. G., et al. 1979. Overwintering strategies of gall insects. Physiol. Zool., 52:572-580. Belozerov, V. N. 1982. Diapause and biological rhythms in ticks. In : Obenchain, F. D. and R. Galun, Eds. PHYSIOLOGY OF TICKS. Pergamon Press, Oxford. 509 pages. Boss, K. J. 1974. Oblornovism in the mollusca. Trans. Arner. Micros. Soc., 93:460-481. Burke, M. J., et al. 1976. Freezing and injury in plants. Ann. Rev. Plant Physiol., 27:507-528. Committee E-20, ASMT. 1974. MANUAL ON THE USE OF THERMOCOUPLES IN TEMPEFWTURE MEASUREMENT. American Society for Testing and Materials, Philadelphia, PA. 252 pages. Danks, H. V. 1978. Modes of seasonal adaptation in insects. I. Winter survival. Can. Ent., 110: 11 67-1 205. Danks, H. V. 1987. INSECT DORMANCY: AN ECOLOGICAL PERSPECTIVE. Biol. Survey Can., Ottawa, Canada, 439 pages. Douzou, P. 1977. CRYOBIOCHEMISTRY. Academic Press, New York. 286 pages. Duman, J. G. 1979. Thermal-hysteresis factors in overwintering insects. J. lnsect Physiol., 25:805-810. Duman, J. G., et a1.1982. Anti-freezes of terrestrial arthropods. Comp. Biochern. Physiol., 73 A:545-555. Flint. H. L. 1972. Cold hardiness of twigs of Quercus rubra L. as a function of geographic orgin. Ecology, 53:1163-1170. George, M. F., et al. 1974. Low temperature exotherms and woody plant distribution. Hort. Sci., 9:519-522. Hirsch, A. G., et al. 1985. A novel method of natural cryoprotection. Plant Physiol., 79:41-56. Hochacha, P. W. and G. N. Sornero. 1984. BIOCHEMICAL ADAPTATION. Princeton University Press, Princeton, NJ. 537 pages. Krog, J. O., et al. 1979. Thermal buffering in Afro-alpine plants due to nucleating agent-induced water freezing. Nature, 282:300-301. Lee. R. E. 1980. Physiological adaptation of Coccinellidae to supranivean and subnivean hibernacula. J. lnsect Physiol., 26:135-138. Lee, R. E., et al. 1986. Low temperature tolerance in insects and other terrestrial arthropods: bibliography II. Cryo-leters, 7: 1 13-1 26. Leopold, A. C.. Ed. 1986. MEMBRANES, METABOLISM AND DRY ORGANISMS. Cornell University Press, Ithaca, NY. 374 pages. Lozina-Lozinskii, L. K. 1974. STUDIES IN CRYOBIOLOGY. John Wiley, NY. 259 pages. Morrissey, R. E. and J. G. Baust. 1976. The ontogeny of cold tolerance in the gall fly, Eurosta solidagensis. J. lnsect Physiol., 22:431-437. Murphy, D. J. and S. K. Pierce. 1975. The physiological basis for changes in the freezing tolerance of intertidal molluscs. J. Exp. Zool., 193:313-322. Perry, T. 0. 1971. Dormancy of trees in winter. Science, 171 :29-36. Salt, R. W. 1961. Principles of insect cold-hardiness. Ann. Rev. Ent., 6:55-73. Salt, R. W. 1966. Effect of cooling rate on the freezing temperatures of supercooling insects. Can. J. Zool., 44:655-659. Schmid, W. D. 1982. Survival of frogs in low temperatures. Science, 215:697-698. Schmid, W. D. 1986. Winter ecology (in Russian). Ekologia, 17(6):29-35. Somme, L. 1964. Effects of glycerol on cold-hardiness in insects. Can. J. Zool., 42:87-101. Storey, K. B. and J. M. Storey. 1986. Freeze tolerant frogs. Can. J. Zool., 64: 49-56. Weiser, C. J. 1970. Cold resistance and injury in woody plants. Science. 169:1269-1278. Zachariassen, K. E. and H. T. Hammel. 1976. Nucleating agents in the haemolymph of insects tolerant to freezing. Nature. 262:285-287. Zachariassen, K. E. 1980. The role of polyols and nucleating agents in cold-hardy beetles. J. Comp. Physiol.. 140:227-234. Zachariassen, K. E., et al. 1982. A method for quantitative determination of ice-nucleating agents in insect hemolymph. Cryobiology, 19:180-184.
203
Chapter 12
The Use of Yeast for Teaching Microbiological Techniques and Principles
Robert J. Doyle
Department of Biology University of Windsor Windsor, Ontario, Canada N9B 3P4
This workshop was not included in this proceedings volume because it was not received in time for publication.
205
Chapter 13
Ideas to Stimulate the Non-Major Biology Student
A. Understanding Human Energy Requirements- Roberta B. Williams
B. Ideas to Stimulate the Non-Major Biology Student-Haven Sweet
C. Biology from the Human Perspective-Barbara Newman
Note: This workshop consisted of three separate subtitles and presentations.
207
Understanding Human Energy Requirements - A Laboratory Exercise
Roberta B. Williams
Department of Biological Sciences University of Nevada, Las Vegas 4505 Maryland Parkway Las Vegas, NV 891 54
Roberta Williams is an instructor and the undergraduate laboratory coordinator for the Department of Biological Sciences at the University of Nevada, Las Vegas. She recieved a B.S. from chestnut Hill College in Philadelphia (chemistry) and a M.S. from the University of Nevada, Las Vegas (botany). Williams teaches Human Biology, a science course for non-majors, and is involved with teaching post-baccalaureate science classes to primary and secondary teachers. Her research interests are involved with developing new laboratory experinces for all grade levels. Roberta hosted the 1985 ABLE meeting and is currently Labstracts editor.
209
INTRODUCTION
The American public is becoming increasingly more conscious of
their individual responsibility for wellness and the role of nutrition and exercise in maintaining good health. The news me-
dia bombards us daily with tidbits on the latest research con-
cerning nutrition and exercise and their correlation with cancer
and heart disease. We are aware that obesity is American's num-
ber one malnutrition problem and that solving this problem is
one of the least understood areas in the science of nutrition.
We hear that anorexia nervosa and bulimia are increasing at an
alarming rate as some of our brightest young females become more
and more obsessed with weight. But how many of us know what our
own "ideal weight" should be?
Teaching a course in Human Biology to college non-science majors
has made me realize how little students really know about their
own energy metabolism. They seem to be aware of nutritional re-
quirements, but what happens after they eat is a mystery. To
solve this mystery, I designed a laboratory exercise that deals
with daily caloric intake, energy metabolism and body composition.
This exercise enables the individual student to evaluate his cal-
orice intake and caloric expenditure for an average day and ex-
amine why he or she may be gaining or losing weight. The exercise
then enables the student to determine his or her percent body
fat and calculate what their "ideal weight" should be. Each semes-
211 ter the majority of my 200 students rate this particular labo- ratory experience as their favorite and the one from which they learn the most.
One of the objectives of the lab is to understand energy meta- bolism, the process by which the body stores or releases energy from nutrients consumed. Since the body's total energy needs fall into three categories: energy to support basal metabolism, energy for muscular activity and energy to digest and metabolize food. The students are taught how to calculate each of these using their own height and weight.
Basal metabolism is the minimum amount of energy the body needs at rest in the fasting state. Certain processes necessary for the maintenance of life proceed without conscious awareness. The beating of the heart, the inhaling of oxygen and the exhaling of carbon dioxide, the metabolic activities of each cell, the main- tenance of body temperature, and the sending of nerve impulses from the brain to direct these automatic activities are some of the basal metabolic processes that maintain life. Their minimum energy needs must be met before any calories can be used for physical activity or for the digestion of food.
The basal metabolic rate (BMR) is the rate at which calories are spent for these maintenance activities. The BMR varies from per- son to person and may change for one individual with a change in circumstance, physical condition or age. The BMR is lowest when
212 a person is lying down in a room with a comfortable temperature and is not digesting any food. At this time, the least amount of oxygen is needed and the least amount of heat is being generated by the activities of the cells. During sleep, the person is more relaxed, but there is more muscular activity, so BMR tends to be slightly higher.
The BMR is influenced by a number of factors. In general, the younger the person is, the higher the BMR. This is due to the increased activity of cells undergoing division. After growth stops, the BMR decreases about two percent per decade throughout life (Food and Nutrition Board, 1974). Body surface area, but not weight, influences BMR. Two people with different shapes who weigh
the same will have different BMR. A short, stout person will gen- erally have a slower BMR than a tall, thin person even if they weigh the same. The tall thin person has a greater skin surface
from which heat is lost by radiation and so must have a faster metabolism to generate the lost heat. Another factor that influ- ences BMR is gender. Males generally have a faster metabolism rate
than females due to the greater percentage of muscle tissue in the male body. Muscle tissue is always active while fat tissue is com- paratively inactive. Conditions such as fever, malnutrition and hormonal secretions can temporarily alter BMR. However, the latest
research shows that physical training and conditioning does not
213 seem to influence BMR after exercise has ceased (personal com- munication, Golding).
The second component of energy metabolism is physical activity voluntarily undertaken and achieved by the use of skeletal muscles. This amounts to an average of about 30% of the total caloric expenditure, while BMR accounts for 60%. Unlike basal metabolism which cannot be changed at will, physical activity can be increased or decreased by an individual. Contraction of muscles uses a large number of calories, and in a moving body the heart must beat faster, this also accounts for additional caloric useage. A heavier person uses more calories performing
the same task in the same time as a lighter person, because it takes extra effort to move the additional body weight. The
longer an activity lasts, the more calories are used; therefore, measurement of physical activity is expressed as calories per weight per unit time.
The final component of energy expenditure has to do with pro- cessing food. When food is taken into the body, many cells be- come active. Muscles move the food through the intestinal tract by speeding up their rhythmic contractions, while the cells that manufacture and secrete digestive juice begin to do their jobs.
Al l these cells need extra energy to participate in digestion,
absorption and metabolism of food. In addition, the presence of
food stimulates general metabolism. Al l of this is referred to
214 as Specific Dynamic Action of food (SDA) and represents about 10% of the total calories expended by a person per day.
The week before the lab is scheduled, students are told to keep a written diary of thier physical activities for three consecutive days. Every minute of the day should be accounted for (1440 minutes in all). Figure 1 is an example of the activity diary forms I pro- vide my students. In addition to their activity diary, the student are told to keep a written food diary for the same three days. The food diary should list all the food and beverages consumed through- out the three day period. Students are reminded not to forget to include snacks and alcoholic beverages. I use three days activities and food consumption and take an average to represent a typical day.
I have found it you ask a student to keep the diaries for only one day, they will pick a very atypical day. Both of these diaries are brought to lab and used to calculate the individual 's energy expendi- ture and caloric intake. The second objective of the laboratory ex- perience is to determine if the individual's energy input balances with their energy output. Weight gain or loss depends on the dif- ference between these two factors. A difference of 3500 calories can mean a pound more or less of body weight.
The third objective of the laboratory exercise is to have the stu- dents determine their own percent body fat and to understand the relationship between body fat and body weight. Body fat plays im- portant roles in maintaining health. It serves as an insulator from
215 heat, cold and mechanical shock and as an energy supply to be
used when glycogen reserves are exhausted. Without a protective
layer of fat, the body is fragile and unable to withstand envi-
ronmental stresses. Some body fat is essential, but excess body
fat serves no useful function in a society where food is abun-
dant and easily obtained and the hazards of being obese are
numerous.
Fat normally makes up about 18% of an adult males's body and
about 22% of an adult female's body. The rest of the body com-
position is muscle, bone, other connective tissues and water.
The relative amounts of muscle and bone vary widely from person
to person. An athlete or person doing heavy physical work, whose
skeleton has become dense through constant stress on the bones,
may have a slender figure with no excess fat tissue and still be
heavier than another person of the same height, sex, age and body
shape. An ideal weight for a person cannot be stated on the basis
of height alone. The "ideal weight" tables published by insurance
companies are merely averages for the U. S. population and have
little scientific validity. At best, they can serve as arbitrary
measures for too little or too much body weight. A person more
than 10% over the weight on the tables is overweight; a person 20%
over is obese. Similarly, a person 10% below the weight on the
tables is underweight.
A direct measure of the amount of body fat can be obtained by means
216 of the skinfold test. In this lab one student measures three or four skinfolds on another student using inexpensive calipers
(Fat-0-Meters). The fat attached to the skin is roughly pro- portional to total body fat, and the measurements can be easily converted to percent body fat.
PROCEDURES
Basal Metabolic Rate. There are numerous ways to measure BMR some of which entail twelve hours of fasting and elaborate instru- mentation. From these methods, Boothy (1936) has developed cal-
culations and charts to give a mathematical estimate for BMR. I
have adapted these for use in this laboratory exercise.
Scales and height-measuring devices are set up in the laboratory,
and each student determines his or her own height and weight. The
calculations required assume the individual is fully clothed and
wearing shoes with a one-inch heel. Figure 2 is used to determine
body surface area. A straight line drawn from height to weight on
Figure 2 intersects the middle column to indicate the corresponding
surface area in square meters. Table 1 is used next to determine
a BMR constant for the subject's sex and age. The surface area de-
termined in Figure 2 is multiplied by this factor to determine
calories per hour. This number in turn is multiplied by 24 hours to
obtain the student's BMR in calories per day.
For example, take a 17 year old male who weighs 170 lbs. (77.3 kg)
and is 6 feet tall. His body surface area from Figure 2 would be 1 .99
217 square meters. From Table 2, he would have a basal metabolism rate constant of 41.5 calories per square meter per hour. This, multi- plied by 1.99 square meters, equals 82.6 calories per hour and, by
24 hours, equals a BMR of 1982 calories per day. Numbers are rounded off because this method is not accurate enough to make decimals meaningful.
Physical Activity. The term physical activity is used in this exer- cise to mean that energy expended during non-sleeping periods by skeletal muscles. The amount of energy expended will depend on the size of the body, the type of activity and the length of the acti- vity. Table 2 lists the total energy expended (cal/kg) for various activities. Each activity in the diary prepared by the student the previous week should be grouped into one of the categories listed.
Multiply the minutes spent in that activity by the appropriate factor and by the body weight in kilograms. Obtain a grand total for the three days and divide that by three to get an average caloric expenditure per day.
If the 17 year old's activities for the three days included:
1440 minutes of sleeping (no activity)
1440 minutes of sitting (very light)
360 minutes of walking (light)
360 minutes of standing (very light)
90 minutes of weight lifting (heavy)
218 450 minutes of driving (very light)
180 minutes of jogging (moderate) he would have expended an average of 1044 calories/day calculated in the following manner:
1440 min. no activity X 0.0 cal/kg X 77.3 kg = 0 cals
2250 min. very light X 0.01 cal/kg X 77.3 kg = 1739 cals
360 min. light X 0.02 cal/kg X 77.3 kg = 557 cals
180 min. moderate X 0.025 cal/kg X 77.3 kg = 348 cals
90 min. heavy X 0.07 cal/kg X 77.3 kg = 487 cals
Total expenditure for three days = 31 31 cals
Average daily expenditure = 1044 cals/day
Specific Dynamic Action. To estimate the students SDA, add the calories calculated for BMR and those calculated for average daily physical activity and multiply by 10%. For the 17 year old example, this would be: [1982 cal (BMR) + 1044 cal/day (acti-
vity)] X 0.1 = 303 cal/day (SDA).
Total Energy Requirement For Average 24 Hour Period. Add all
three figures, BMR, physical activities and SDA to obtain the num- ber of calories needed in one day. The average 70 kilogram adult male requires about 2700 cal/day, while the average 55 kilogram
adult female needs only 2000 cal/day. Our 17 year old example is younger and somewhat more active than the average person, and
therefore, has a higher energy requirement (3329 cal /day).
Energy Intake. Using the data compiled during the week on the
219 subject's food intake, have each student calculate the number of calories consumed from commercial calorie guides. I have found nutri- tion text books contain excellent appendices that include very com- plete calorie guides. For my class I have made numerous copies of
the calorie guide from Hamil ton and Whi tney (1985). Measuring the amount of calories consumed is more accurate than measuring the number of calories expended. If the figures are within 10% of each other, the subject is most likely maintaining his or her weight. In order to maintain proper energy balance within the body, the cal- ories expended must be replaced by an equal number of calories from
food. If this does not occur, the subject will lose weight. If the number of calories consumed exceeds the number of calories expended,
the subject will gain weight. An overage of 500 calories per day for one week can result in gaining a pound (3500 calories equal one pound
of body fat). By the same token, expending 500 more calories than are
consumed each day for a week can result in the loss of one pound.
Estimation of Percent Body Fat.
When individuals gain fat, much of the adipose tissue is added to
subcutaneous accumulations that are found in various parts of the body. This subcutaneous fat can be pinched up by the thumb and fore-
finger. As an individual gets fatter, these skinfolds get larger.
Calipers have been designed to measure skinfolds in several parts
of the body. Any single measurement does not give an accurate picutre. At least three measurements, tricep, illium and abdomen
220 must be taken on women and four measurements, chest, abdomen, il- lium and axilla are necessary for men. All these measurements can be done in the laboratory, provided the females are wearing two piece outfits. The more measurements that are done, the more ac- curate the results will be, however, these minimal measurements give a fairly reliable estimate.
All measurements are taken on the right side of the person being measured. The fold of skin should be firmly grasped between the left thumb and the other four fingers and then lifted. Pinch and lift the fold several times to make certain that no musculature is grasped. Hold the skinfold firmly and place the contact side of the calipers below the thumb and fingers. Do not let go of the fold.
Take the reading to the nearest half millimeter. Release the grip on the caliper and release the fold. To make sure that the reading is accurate, repeat the measurement two or three times. Unless each measurement is consistent (within 1-2 mm) reliability will be poor.
The tricep skinfold is measured on the back of the upper right arm, half-way between the elbow and the tip of the shoulder, while the arm is hanging loosely at the subject's side. Grasp the skinfold parallel to the long axis of the arm, and lift it away from the arm to make sure no muscle tissue is caught in the fold. The il- lium skinfold (hip) is measured with a diagonal fold, just above the crest of the hip bone, on an imaginary line that would divide
221 the body into front and back halves.
The abdominal skinfold is measured vertically one inch to the right of the navel.
The chest measurement is taken diagonal l y, mid -way between the nipple and the armpit.
The axillary (side) measurement is taken vertically at the
level of the nipple on an imaginary line that would divide the body into front and back halves.
When students do skinfold measurements on one another, lack of
experience can produce errors. The most common errors occur when
the mid-point is incorrectly marked or measured, when the cali-
per is too deep (muscle involved) or too shallow (only skin
grasped), and when the arm being measured is not hanging loosely
at the subject's side.
The percent body fat can be determined from these measurements
using charts that accompany the calipers or by the use of two
equations. The equations were developed by Jackson and Pollock
(1978) and are widely used in physical fitness evaluation programs.
For males the four measurements are summed and percent fat is cal-
culated (Golding, 1982) as follows:
2 % fat = .27784 (X1) - .00053 (X1) + .12437 (X2) - 3.28791
where X1 = sum of four folds
X2 = age of subject. It is suggested that from a health and aesthetic standpoint,
222 adult males should have 16% or less body fat, and adult females should have 23% or less body fat. At no time should the percent body fat of an adult male drop below 5% or that of an adult fe- male below 10%. Severe medical problems can occur with too lit- tle body fat as well as with too much body fat. Marginal obesity occurs when the body fat of an adult male increases to over 20% or that of an adult female increases to over 30%. The percent body fat of an individual can change with exercise or diet. You can remain at the same weight while changing your percent body fat. Well conditioned athletes, such as marathon runners and swimmers, usually have about 10-12% body fat, while football players and weight lifters may have as high as 19-20% body fat.
Life style can play an important role in an individual's physical makeup (Golding, 1982).
A "target" or "ideal" weight can be calculated using the percent fat figures. Target weight is defined as the lean body weight
(LBW) plus a desirable percentage of fat. If a 20 year old male student is 210 pounds and has 23% fat, he may wish to know what he should weigh with 16% fat. To calculate this he would multiply
23% times 210 to determine that he is carrying 48.3 pounds of fat. Subtracting his fat weight from his total weight will give his LBW of 161 .7 pounds. At 16% fat, this LBW equals 84% of the student's total weight. To determine what his weight should be with 16% fat, divide the LBW by 84% (161.7/.84 = 192.5). A tar-
223 get weight of 192.5 pounds is thereby obtained.
SUMMARY
What makes this laboratory experience successful? I think it is the fact that although we are all aware of calories, none of us really sit down and evaluate what those calories really mean. Pre- pared foods give the caloric value on box tops and popular maga- zines frequently publish calorie expenditure guides. What this means on a day-by-day basis is rarely given more than a passing thought. In this exercise, the students get a chance to see what their actual energy balance is. Even if they only make this cal- culation once in their life time, it is something they will rem- ember.
We are what we eat, and many researchers are now finding that patterns for both obesity and underweight may be set very early in our life time. Overweight or obesity is seen in more than 10% of school-age children in the United States, in about 15% of peo- ple under 30, and in 25 to 30% of adults. Among older people, a third of the men and half of the women are obese. Certain subgroups of the population have a markedly higher incidence of obesity than others: the lower socioeconomic classes, blacks, Mexican Americans,
Native Americans and Eskimos (Hamilton, 1985). Statistics show that some people become fat in childhood and others later on. Research has shown that early onset obesity is especially resistant to treatment. According to the fat-cell theory, early overfeeding is
224 is thought to stimulate fat cells to increase abnormally in num- ber. The number of fat cells then become fixed in adulthood.
Thereafter, a gain in weight can take place only through an in- crease in the size of fat cells. The larger the number of fat cells in his body, the more hungry the person will be. Thus, peo- ple with abnormally large numbers of fat cells will tend to be abnormally hungry and to overeat.
The causes of underweight are as diverse as those of overeating.
Psychological factors may contribute in some cases and metabolic ones in others. Clearly, heredity is involved. Early underfeeding may limit the fat-cell number, just as overfeeding my increase it.
Habits learned early in childhood, especially food aversions, may perpetuate the problem. The demand for calories to support high levels of physical activity and growth often contributes; an extremely active boy during his adolescent growth spurt may need more than 4000 calories a day to maintain his weight. Such a boy may be too busy to take the time to eat that much. The concepts studied with this laboratory exercise help to make students aware of their body composition. I think this exercise is not only valuable for college students; rather, secondary science and health students can also gain much knowledge that can influence their fu- ture lives. A local elementary health and physical education teacher uses parts of this exercise with his students and claims he has had great success.
225 REFERENCES
1. Boothby, W. M., Berkson, J. and Dunn, H. L, (1936), Studies
of the Energy of Normal Individuals: A Standard for Basal
Metabolism, with a Nomogram for Clinical Application,
Ameri can Journal of Physiology, 11 6. pg. 468-484.
2. Boothby, W. M. in Handbook of Biological Data, ed. W. S.
Spector. (1956). Saunder's Publishing Co., Philadelphia,
Pa. pg. 259.
3. Food and Nutrition Board, Commi ttee on Recommended Allo-
wances, (1974). Recommended dietary allowances, 8th ed.
National Academy of Science, Washington, D. C.
4. Golding, Lawrence, Myers, C. R. and Sinning, W. E. (1982).
The Y's Way to Physical Fitness, National Board of YMCA,
Chicago, IL.
5. Hamilton, E. M. N and Whitney, E. N. (1979). Nutrition:
Concepts and Controversies, West Publishing Co., St. Paul,
MN .
6. Hamilton, E. M. N, Whitney, E. N. and Sizer, F. S. (1985).
Nutrition: Concepts and Controversies, Third Edition,
West Publishing Co., St. Paul, MN.
7. Jackson, A. S. and Pollock, M. L. (1978). Generalized equa-
tions for predicting body density of man. British Journal
of Nutrition, 40, pg. 497.
226 227 228 Figure 2.
Nomogram to estimate body surface area from height and weight.
A straight line is drawn from the subject's heights (Scale 1) to the subject's weight (Scale 3). The point at which the line intersects Scale 2 will give the subject's body surface area in meters squared. Adapted from Boothby, W. M., J. Berkson and H. L. Dunn, Studies of the Energy of Normal Individuals:
A Standard for Basal Metabolism with a Nomogram for Clinical
Application, American Journal of Physiology, 116, (1936);
468-484 with permission of the publ isher.
229 230 Table 2. Examples of Daily Energy Expenditures
TOTAL ENERGY EXPENDED TYPE OF ACTIVITY (cal/kg of body weight/min)
NO ACTIVITY: Sleep 0.0
VERY LIGHT: Sitting, standing, 0.01 driving, typing, playing musical instruments, sewing, ironing, walking slowly
LIGHT: Walking at moderate speed, 0.02 light housework, garage work, restaurant trades, gol f , sai l ing, table tennis, volleyball, carrying light loads
MODERATE: Walking fast or jogging, 0.025 weeding and hoeing, scrubbing floors, carrying heavy loads, cycling at moderate speed, skiing, dancing
HEAVY: Walking quickly up hill, 0.07 climbing stairs, basketball, weight lifting, swimming, climbing, football
SEVERE: Tennis, running 0.11
VERY SEVERE: Wrestling, boxing, racing 0.14
Modified from Food and Nutrition Board, Recommended Dietary Al lowances , 1980.
231
Ideas to Stimulate the Non-Major Biology Student
Haven C. Sweet
Department of Biological Sciences University of Central Florida Orlando, FL 3281 6-0990
Haven C. Sweet received his B.S. degree from Tufts University (Botany) and his Ph.D. from Syracuse University (Plant Physiology). After completing a Post- Doctoral fellowship at Brookhaven National Laboratory, he worked at the Johnson Space Center with the Apollo program. In 1971 he moved to the University of Central Florida where he developed and teaches the non-majors biology course, along with a variety of upper level biology and computer courses. His creative interests involve developing computer programs for both research and educational purposes.
233
Ideas to Stimulate the Non-Major Biology Student
ABLE workshop- June 18,1987 Haven Sweet University of Central Florida Orlando, FL 32816
Introduction
The computer programs described in this workshop were created as part of a National Science Foundation grant to develop new laboratory exercises involving the computer. As the programs were written, I incorporated them into my non-majors biology course and revised them according to student reactions. After several years of improving these exercises, I evolved an approach which is successful from an educational point of view, is compatible with very limited budgets, and is transportable to other institutions. This discussion presents the philosophy, as well as specific implementation plans, for incorporating computer programs into a non-majors biology teaching laboratory.
All to often, educators assume that using a computer within a course requires enough machines for each student. Although it is possible to have numerous computers in the laboratory and perform very meaningful activities, most schools do not have the necessary resources for this approach. A single computer is often adequate if the computer program compliments the lab exercise. Just as one can develop a meaningful lab experience around a single spectrophotometer or analytical balance, likewise it can be done around a single computer.
In fact, the educational objectives are often better achieved by de-emphasizing equipment, especially when the teacher avoids "showcasing" the computer. If students perceive the computer as the main focus of the laboratory, they tend to do nothing except wait for their turn.
Assuming that only one or two computers are available, the computer programs selected should meet the following criteria; they should accomplish something that could not otherwise be done (or requires additional equipment); they should eliminate repetitive tasks that do not promote learning (remember that repetition, per se, is not bad); they must be very easy to use so the student does not loose sight of the exercise's objective; and the activity must be only a small component of the entire lab experience. The programs described below were designed to meet these criteria.
All programs are written for the Apple ][ computer and will run with 48K of memory and a single disk drive. Although some programs will display 80 columns if the computer is capable, all work with 40 columns. A machine such as the Apple // C is particularly well suited for use in labs since it is highly portable. While the programs could be adapted to machines other than the Apple, conversion would be a major undertaking with most of the programs.
235 In the following discussion, I will start with the major programs and then describe the smaller exercises. The exercise instructions which are provided to the students have been abbreviated in this manuscript; the complete versions are available in "Non-Majors Biology Laboratory Notebook" (Burgess Publishing Co., 1984).
A. Analysis of nutrients; 1. General Using this program, students enter the foods they consumed during 24 hours and the computer lists all the nutrients in each food, gives a daily total, and compares it to the recommended daily allowance (RDA). In addition to allowing students to assess their personal diets, this exercise shows how nutritional needs are usually met, even by the typical diet of a college student.
This exercise requires 1 computer for 6 to 8 students in a 2 hour lab period. Using the computer to look up nutritional data and to perform the menial computations for each element eliminates the tedium from this task. If a fast printer is available, the time spent on the computer is reduced since students would not have to copy information from the screen. Since determining nutrients is easy, and since they are not graded on how they eat, students are more honest about recording the foods they eat.
To insure that students do not sit and wait for a free computer, the computer calculations provide only a portion of the exercise; they are combined with manual calculations of calories burned over a twenty-four hour period. Although I had written a computer program to calculate metabolic rate, I found students derived no understanding from running it. By making the computations, they recognize some factors considered in determining metabolic rates. Alternatively, it would be reasonable to include exercises such as measuring body fat or performing an assay for vitamin C, especially if the lab is three hours long.
2. The program There are a variety of nutrient analysis programs on the market that run on computers other than the Apple ][. Although some trial and error may be necessary to select a program which is easily run by students, it should be possible to purchase a suitable program. The version I developed utilizes nutrition data which were originally derived from the USDA nutrition handbook and were subsequently modified by Dr. Javed Aslam and Apple Computer. My program disk contains 8 files, all of which must be present for the program to function. Nothing else can be stored on the disk due to a lack of space. The disk must not be write protected since data are temporally stored during the program's operation. The RDA values given are those set by the US Academy of Science and are often much higher than found in other countries.
Beginning several weeks in advance, I repeatedly announce in class that the students will need to record their daily activities and diet to insure they come to lab with real data. Encourage the students to be honest when recording their diet since
236 they are usually surprised to find that their nutrient intake is adequate. If they think you will be grading the way they eat, they will not be as honest.
3. The exercise a. Preface As a preface to the exercise, I provide excerpts from a series of articles dealing with diets, dieting, and the genetic nature of being overweight. These have been omitted from the following material which is distributed to the student.
b. Introduction Although it is not possible to taste or otherwise detect the nutrients in foods, chemical tests can measure which are present in what concentrations. Performing these tests are well beyond the scope of this course; however, we can use the data accumulated by others (e.g., the "Composition of Foods," Agricultural Handbook #8) to evaluate our diets. In addition, while the energy you expended over a 24-hour period is also very difficult to measure, we can use indirect methods to estimate approximately how much energy you burned. Thus, the objective of this laboratory is to determine the quantity of nutrients and energy that you received during a typical day, and then to compare these values to the quantity that you require. This exercise should be very illuminating since you may find that the way you eat is actually adequate to meet your needs.
To accomplish these objectives, you will have to accurately record both your activities and foods consumed for a 24-hour period. When you are recording your data, BE HONEST! You will not be graded on the way you eat or on your activities, but rather on the way you complete the lab exercise. For this experience to be beneficial to you, you should try to select a day that is representative of the way you eat and live. Thus, while it would be much easier to record your activities on a day spent sick in bed, or inventory your foods when you didn't have money to eat, the results would be of no value to you.
For maximum accuracy, carry a note pad with you and record throughout the day, rather than trying to recall the information at the end of the day. Record accurate estimates of the quantity of each food eaten, and the intensity of each activity, on Form 1. Please be neat so both you and your instructor can easily check the figures.
When in the laboratory, you will determine the nutritional content of foods using the Apple computer to record and calculate your data. Calculations needed for estimating your energy expenditure will be done by hand (or a calculator if you have one), so while you are waiting for an available computer, you should work on the second section of the exercise.
c. Food Intake Use Form 1 to record all foods you eat during the day. For nutrient analysis, it is important to estimate as accurately as possible the quantity of each food consumed. Since the data on the computer have been converted to ounces, cups, tablespoons or teaspoons, use the English units of measurement in your estimates. To help you
237 estimate, it may be helpful to measure out a cup, tablespoon, and teaspoon of a food, and then observe it on a plate or in a bowl. A piece of meat the size of your hand is about 3 to 4 ounces, while a single slice of American cheese is about 1 ounce.
When recording your foods, be sure to list any dressings, sauces, spreads, gravies, etc.; also indicate how the food was prepared since fried food has added fat, while other methods remove fat. For mixed dishes, estimate and record the amounts of the major ingredients. For example, one serving of macaroni and cheese might be listed as .05 cup cooked macaroni plus 1 ounce American cheese. For foods that are often served differently, indicate how yours was served.
Thus, include any cream and sugar you use in your coffee, butter and/or spreads used on your toast, the type of salad dressing you used, etc. If you took any vitamin pills, record the ingredients from the label and manually add them to the computerized totals when you are finished.
Once you are in the lab, enter each food type into the computer. Type in the general class of the food rather than the specific name: for example, if you had angel food cake, type "cake," while for brown rice, enter "rice." The computer will list all the types of cake or rice that are on file. Select the appropriate food by entering the number beside the food, then pressing "RETURN." If none are appropriate, press "RETURN" without entering a number. Since you are only permitted to enter a single word, to select peanut butter, type the word "peanut;" then from the displayed list of peanut products select "peanuts, miscellaneous" to see another list which includes peanut butter. If you misspell the word or enter a word that is not in the program, the computer will progressively drop the last letter of the word and scan for a match. If you are not sure of the spelling of the word, you only need to enter the portion of which you are certain.
After selecting the food, you will be asked the quantity you consumed. If the units given seem inappropriate (e.g., you consumed 3 ounces of juice, yet you are asked how many cups you had), press the "ESC" key and select the more appropriate units. Then proceed to enter the quantity that you consumed.
The computer has room for only 40 foods, so you should consolidate your entries. Thus, if you had 3 cups of coffee each with a teaspoon of sugar, one spoonful on your cereal, and one with iced tea, then you only need to enter sugar one time. When asked how many teaspoons of sugar you consumed, enter 5.
After entering all foods for the entire day, press the "ESC" key and verify that the foods you entered are correct. If you decide to add additional foods, press "A." To delete a food, enter "D." If you are satisfied with the list, press "C" to enter the computational portion of the program.
The computer will read the nutritional data from the disk and correct the values for the quantity you consumed. You will then be asked for the information needed to compute your RDA. Summary data which includes both the totals for the foods you
238 consumed, plus the recommended daily allowance that you should strive for are then presented. To observe all three screens of data, press "RETURN" to advance to the next screen of information. Record this summary data using Form 3 in your notebook. If the "ESC" key is pressed, you will see a detailed listing of all the nutrients contained in each food; since there is not enough room to see all 24 nutrients at one time, press the right arrow key to see data to the right. Each time an arrow key is pressed, two new nutrients move onto the screen and two exit from the other side. If the TOTAL of the nutrients is not visible at the screen's bottom, press the "V" key to view the bottom of the listings.
Pressing the "RETURN" key will display the complete name of each food and quantity you consumed. A second press of the "RETURN" key returns you to the nutrient table.
After viewing the detailed data, press "ESC" and you can choose to review the data again, print the data (assuming there is a printer attached), re-enter the food selection program to make minor modification to your diet, or end the program. If you wish to try computing the nutrients from a second day, please wait until everyone else has had a change to accumulate their day's data.
d. Energy Expenditure Record your activities for the same 24-hour period of Form 1. To record the time spent, you can either enter the actual clock time the task was initiated and completed (best with long activities such as sleeping) or directly enter in the minutes spent doing the activity. Each minute of the day should be accounted for, so the total must equal 1440 minutes. Treat flights of stairs as separate functions. On your Form, record the time spent on stairs as "walking," then also list the number of flights you ascended and descended. Count 14-1 5 steps as being a flight of stairs.
When you are in lab, use the description in Form 4 to determine what energy level would be most appropriate for each activity you performed. Then transfer the information to Form 2, enter number of minutes spent performing each activity into the box below the appropriate energy class. Total the number of minutes spent in each activity level at the bottom of the sheet. Transfer this data to Form 4 and multiply the minutes spent at each level with the energy consumed per Kg body weight per minute. Be sure to also include the number of flights of stairs traversed. When the total is multiplied by your body weight, you have computed the amount of energy you used for muscular activity for the day.
To estimate your total energy expended, use Form 5. The basal energy used to maintain your body must first be estimated, and then corrected for your age. In addition, since our metabolic rates apparently increase when we eat more calories, and decrease when we consume less, we also need to estimate how much metabolic compensation could occur in your case. When the basal rate is added to your muscular energy consumption, the result is the approximate amount of energy that your body expended during the day. The internal compensation for extra calories serves as an error term.
239 Form 1. Work sheet
Foods consumed during 24 hours.
Food Quantity Consumed Food Quantity Consumed
Activities during 24 hours.
Flights of stairs: Up: Down:
240 241 242 243 Form 5. Estimation of Total Energy Expenditure
A. Calculate the energy spent on basal metabolism; the basal metabolism rate (BMR) for males is 1.0 Kcal/kg body wt./hour while the rate for women is 0.9 Kcal/kg body wt./hour.
Kcal/kg/hour X kg X 24 hours = Kcal (BMR) (your body (energy for basal weight) metabolism per day)
(should be 1000-3500)
B. Correct BMR for age; approximately 8 fewer Kcal are needed each year after 18.
Kcal- ( X 8 Kcal) = Kcal (Energy for basal (# years you are (age corrected energy metabolism) older than 18) for basal metabolism)
C. Compute the range that your metabolism rate could vary in response to an increase in diet; if you are of normal weight, use a metabolic compensation factor of 0.2. If you are overweight, use a factor of 0.1.
Kcal X = Kcal (age corrected (metabolic (compensation energy for basal compensation energy) metabolism) factor)
D. Compute your total energy expenditure for the day.
Kcal + Kcal = ± (energy for mus- (age corrected (energy (compensa- cular activity) energy for expenditure) tion energy) basal metabolism)
244 e. Discussion How many Kcal did you consume during 24 hours (see Form 3)? . How many Kcal did your body burn during the same period (see Form 5)? . Assuming that these values are typical for you, and that your metabolism runs at a constant rate, what is the difference between your intake of calories and the calories required for your activities plus BMR? (Use negative numbers if your metabolism consumes more than you took in) . If this discrepancy were repeated daily for five years, how many surplus (deficit) Kcalories would you have eaten? Considering that one pound of fat contains 4,090 calories, how many pounds of weight gain (loss) could this discrepancy translate into? . If you continued to consume foods at the same rate and you kept your activity level unchanged over the next fifty years, what would your weight be?
Now return to Form 5 and recompute what your energy expenditure would be in five years assuming your body weight had increased by this amount, and your age was five years older. How many calories would your body be burning per day five years from now? . Why is this value different from your present metabolic rate?
What internal compensation rate did you compute for your caloric intake? (see Form 5) . Is this value large enough to permit you to maintain a stable weight without changing your activity level and/or food consumption? . Do these calculations agree with your own personal experience? I.e., do you find you tend to gain (lose) weight unless you work at maintaining a stable weight? . Please explain.
Our diets should provide less than 30% of the total calories needed in the form of fat. How many grams of fat did you consume (Form 3)? . Since fat contains 9 calories/gram, how many calories of fat did you consume? . How many total calories did you consume (Form 3)? . What percentage of your calories were derived from fat? . Is this value within the recommended limits of fat? . What foods were the major source of fat in your diet?
VITAMINS Do you feel that your nutrient intake as recorded during this study is typical of your normal eating habits? . If not, please explain.
Reviewing Form 3, did you meet the RDA for all nutrients? Which nutrients were consumed at levels less than approximately 80% of your RDA?
Which nutrients were consumed at levels which exceeded the RDA by more than approximately 50%?
Do you see a need to take nutrient supplements?
245 B. Health Hazard Appraisal 1. General With this program, students answer a set of questions depending on their age, sex and race (the 3 major risk factors which affect mortality). Then, depending on the student's family history and personal habits, the computer provides a personalized assessment of their risk of death. The three objectives of this exercise are to convince an 19 year old he/she is mortal; to show how one's activities can influence survival; and, to show how changing life style can improve life span.
There should be 1 computer for at least every 7 students if this is to be done in a two hour lab period. If a printer is available, it should be used since there is much text for the student to read. To insure that students learn additional material while waiting for the computer, I combine this exercise with a variety of investigations that relate to common causes of mortality. Thus, the students use the microscope to compare lung tissue from smokers' and non-smokers' lungs, view lung tissue damaged by emphysema, observe a variety of cancerous tissues, compare coronary arteries with various states of atherosclerosis, survey attitudes on risk factors, and measure cardiovascular fitness. Other exercises that might be appropriate are measuring body fat, observing the effects of nicotine on ciliated organisms, or studying heart anatomy.
This program performs an enormous number of calculations and looks up a vast quantity of information. It would be unreasonable to expect students to perform this amount of work manually, so the use of a computer is appropriate. The resulting information is personalized, and as such, is more meaningful to the student.
2 The program Students of different races, ages and sexes will be asked different questions. Some of the questions for females are very personal since the program is looking for links to cervical cancer and breast cancer; it is probably best to encourage people not to look over each other's shoulders while they are at the computer. Sometimes I designate separate machines for males and females.
One factor that hurts students is their excessive driving which can skew the data; a 15 year old who drives 60,000 miles per year could have the same risk of death as a 41 year old male. Another confusing aspect is that there are no listed risk factors for certain causes of death. This is an omission of the original study, presumably due to a lack of data. Thus, while logic would assume that not knowing how to swim would be a risk factor in swimming deaths, non-swimmers are less likely to go near the water, so it may not be a risk factor.
There are 32 files on the disk, all essential for the program to work. There is no room on the disk for anything else. The disk must not be write protected since data are stored during the program's operation. These data files are from "Prospective Medicine" by J. Hall and J. Zwemer, Methodist Hospital of Indiana, 1979, and represent the best correlations between observed mortality and associated risk factors.
246 3. The exercise a. Preface: -Readings omitted -
b. Introduction The objective of this week's laboratory is to observe some of the factors which present a risk to our survival. Due to our myopic perspective, we tend to accept or ignore risk factors which are of enormous significance, while being unduly concerned about factors having a minimal effect on survival: In addition, there is also a tendency to assume that we are condemned by our genetic heritage. Thus, some people assume that since cancer or heart disease runs in their family, there is nothing they can do to avoid a similar fate.
Fortunately this is not the case. For example, although a woman's risk of breast cancer is approximately two to three times greater if a mother or sister also had the disease, 90% of the women contracting the disease have no relatives with the breast cancer; in addition, of those who do develop tumors in one breast, very few will also form a mass in the other breast. This evidence indicates that, although the disease has a genetic component, most people contract breast cancers due to environmental reasons. Thus, instead of resigning yourself to the fate of your ancestors, you should attempt to modify your environment and reduce your chances of such a fate.
Hopefully, at the completion of this exercise, you will have a better understanding of the causes of death that are most prevalent for persons like you. In addition, you should understand which factors influence survival and which are most important for you to modify.
c. Causes of Death 1. Major risk factors The three risk factors which have the most influence on our mortality are beyond our capability to change; these are our age, sex, and race. To demonstrate their influences, I have selected a few conditions as examples. In all of the following discussions, the numbers presented for death rates are based on actual mortality data. They represent the number of people who died out of a population of 100,000 persons of the same sex, age and race.
The effects of age can be seen in this data derived from a white male;
CHANCE OF DEATH WITHIN 10 YEARS DUE TO: CURRENT HEART LUNG AUTO AGE DISEASE CANCER ACCIDENTS SUICIDE CIRRHOSIS 20 17 3 58 1 2 50 11
247 Note that while the death rate due to most conditions increases with age, some, such as auto accidents, decline with maturity. In addition, suicide appears to be independent of age.
To demonstrate the effects of sex on death rates, consider the following data derived from 20-year old whites.
CHANCE OF A 20-YEAR OLD DYING BY: AGE 50 AGE 60 CAUSE MALE FEMALE MALE FEMALE Heart Note that females have a Disease 1,829 361 6,398 1,565 much lower incidence of many diseases. However there are Lung Cancer 382 172 1,467 541 examples of males having similar rates (lesions of Breast Cancer 4 44 1 10 1,094 the central nervous system-- i.e., strokes) and reduced Homicide 442 125 524 149 rates (breast cancer) when compared to females. Suicide 739 322 1,014 447 Vascular Lesions of C.N.S. 244 251 738 654
The effects of race are also pronounced, although for many of the factors, the difference appears to be cultural in origin.
CHANCE OF A 20-YEAR OLD DYING BY: AGE 50 AGE 60 CAUSE BLACK WHITE BLACK WHITE Many mortality factors are Homicide 4,038 442 4,683 524 very much weighted against blacks. Some (auto acci- Cirrhosis 1,241 436 1,947 966 dents) are about equal for both groups, and a few Vascular Lesions (suicide) affect whites more of C.N.S. 848 244 2,117 738 than blacks. The prevalence of strokes in blacks is Auto Accidents 1,290 1,164 1,691 1,390 probably due to genetic causes, since more blacks have Suicide 505 739 602 1,014 high blood pressure.
The tables that follow show the death rates that a 20-year old could expect up through age 60. Similar data are also available for other age groups.
-Tables omitted -
248 2. Other risk factors Numerous other factors have been correlated with different diseases; i.e., persons with a particular trait, condition, habit, occupation, etc., are noted to have a higher incidence of a given disease. However, our interest is in those factors that are responsible for causing the disease, and not those traits which result from the illness. For example, severe weight loss is correlated with many diseases, although the weight probably reflects the progression of the disease rather than the cause of it.
The field of prospective medicine deals with determining which environmental, background and health factors are significant in increasing or decreasing mortality. The primary interest is to identify those traits that could be altered and thereby result in an increased life span and improved quality of life. Remember that a shortened life span is frequently preceded by illness and debilitation. Since the quality of life is very difficult to measure, we will limit our concern to life span.
The fundamental assumptions of prospective medicine are: a. Everyone is subject to the risk of death. b. The risk of death can be described by an average for a person of a given race, sex and age. This risk is based on actual mortality figures for people in that group. c. This average risk can be adjusted for an individual according to key factors such as the person's family history, health condition and personal habits. Some factors will increase the person's chance of death, while others will reduce it. d. If intervention changes some negative factors, the person's chance of survival will increase.
Subjective Rating of Risk Factors - Section omitted -
RISK OF DEATH ASSESSMENT After entering your age, sex and race in the Apple computer, you will be asked a series of personal questions. The particular questions asked will differ depending on the basic parameters of sex, age and race. Your responses to the questions permit the program to assess your risk of dying from the ten most common causes of mortality for persons of your age, sex and race. Your answers are strictly confidential since all data is erased at the completion of the program.
Because most deaths of persons under 25 are due to conditions for which risk factors have not been derived, you may find it more informative to enter your age as 25 if you are younger.
You will then be shown in descending order of frequency the ten most common causes of death for persons with your basic parameters. Your personalized risk of death over the next ten years is given as a percentage, followed by how the value compares to that of others with your characteristics. If your personal percentage of
249 survival is less than one, your chances of surviving the next ten years is greater than that of the sample population; if it is greater than one, your risk of death is greater.
The remaining information presents the data on which the percentage of survival was based. Listed under "FACTORS INFLUENCING RISK" is each of the risk factors and a risk value. A risk value of less than one means that your response to this risk factor has improved your chance of survival. The composite risk value is the sum of all risk values greater than one added to the product of all those less than or equal to one. This composite risk is then multiplied by the number of persons (out of a sample of 100,000 of your age, sex and race) who would be expected to die. This new value is then converted to a percentage to assess your probability of death.
The second phase of the program will permit you to determine the effects of intervention. If you were to immediately correct those risk factors that are in your power to alter, your survival rate should improve. Finally, the program will give you an attainable age. This value is your mortality age assuming you correct the stated risk factors.
RESULTS SUMMARY AGE SEX RACE
Deaths/100,000 People Your Risk Assessment; Cause of Death in Your Age Group Present After Intervention 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. Given your current habits, the potential number of years you have left is equivalent to those of a year old. How could intervention influence your potential life span?
If you were to institute a public health campaign to encourage people in your age group to alter two or more personal habits, what would you focus on?
250 C. Reaction meter 1. General This program tests students' reaction time as they are presented increasingly more complex tasks to perform. The program simulates a reaction meter, although the computerized version does a great deal more than reaction meters. Since this exercise comprises a small portion of a larger exercise on physiological testing, only one computer is needed for 15 to 20 students. This program is also used in our Human Physiology course.
2. The Program The program is self contained, and as such, can be copied to any disk without fear of loosing essential program files. The program and how to modify it have been described in the fall 1986 issue of Labstracts.
3. The exercise a. Introduction The objective of this week's laboratory exercise is to let you try some tests that are often run by doctors. The tests are used to diagnose illness or abnormalities since they indicate if the patient's values are within ranges that are considered normal. Although some of these tests are part of routine physical exams, do NOT consider this lab exercise a substitute for such an exam. We are doing the tests for demonstration purposes only and will not attempt to diagnose or interpret the values.
b. Human Reflexes A reflex mechanism requires an organ for perceiving a stimulus, an organ which can react, and a communication network in between. The communication system can range from very simple reflex loops (arcs) to highly integrated networks involving higher brain functions.
Many reflexes appear programmed; that is, the appropriate response to a given stimulus has been built into the nervous system. The spinal reflexes serve as an excellent example. For instance, if you burn a finger on a hot object, the spinal reflex immediately causes withdrawal of the finger from the hot object. By the time the higher levels of the nervous system are notified of the event by sensations of pain, the finger is well away from danger.
Other reflexes, such as eye reflexes, require that the brain evaluate the stimulus and determine the proper response.
In addition to observing some reflexes, we will measure reaction time of both simple reflex arcs and those involving the higher brain centers.
1. Pupillary Reflex - omitted - 2. The Blink Reflex - omitted - 3. Patellar Reflex - omitted - 4. Plantar Reflex - omitted -
251 5. Reflex and Reaction Times A person's reaction time depends on numerous factors which vary depending on the reflex type; for example, the response speed of the perceiving organ, the speed that nerves conduct the impulse, the number of jumps the impulse must make (synapses) from one cell to another, the speed with which the effector muscle contracts, and the health of the organs involved. Thus, when dealing with a fairly simple reflex arc with few synapses, response time will be rapid. However, if a response requires thought or decision making, more neural pathways must be traversed and the response time is longer.
To demonstrate the influences of these various factors, we will use the Apple computer to simulate a reaction time meter. Simply press "RETURN" to start each test. When you perceive the signal, press the space bar. The computer will tell you how long it took you to react. Be careful not to accidentally press the key while resting your hands on the keyboard.
After one or two practice tries with a section, record the reaction time for three runs and average them. Also note the number of erroneous key presses. Then press the "ESC" key to go to the next test.
PLEASE DO NOT BANG ON THE KEYS!!!
a. Reaction time to sight Test the subject's reaction to the "X" that appears on the screen. Does it make a difference if it is positioned at random on the screen?
b. Reaction time to sound How does the reaction time with sound compare to that with vision? . Look in your book at the portions of the brain which are involved with hearing and vision. With this information in mind, how do you account for the differences in reaction time?
When both vision and hearing are used, which are you responding to? . Which do you perceive as appearing first, the bell or the "X"?
It will probably surprise you to find that the computer prints the letter first, then makes the beep! How do you account for this discrepancy in what your senses are telling you?
c. Reaction times involving decisions How does having to choose before responding affect the reaction rate?
d. Rate of nerve conduction How much longer does it take for a impulse to reach the brain from your feet than your face?
252 e. Involvement of the higher centers How is your reaction time influenced when complex interactions are involved? . By subtracting the time needed to just find the correct number key from the time needed with the math problems, it is possible to determine how much time is needed just for computation.
f. The action of stimuli and depressants Although the tests in this section are strictly optional, please try to complete the questions below. How would you expect alcohol to influence reaction time?
Alcohol will slow down nerve conduction and also the rate of synapses. Since driving a car requires the higher brain centers, evaluate visual impres- sions and perform the correct response, which of the above tests would you assume would be most similar to driving?
If your overall reaction time was slowed by only one second, how much further would you travel if you were driving at 60 miles/hour?
RESULTS
REACTION TIMES NUMBER OF TEST 1 2 3 AVERAGE MISSES Stationary X Random X Click Click & X Arrows Random 0 Words in center Random words Select number Solve Math Touch cheek Touch ankle D Field of vision 1. General This program simulates an expensive piece of specialized equipment for measuring 'a visual field of view. Due to the inability to vary and control the spot brightness, it could not be used in a clinical setting; however, it is very informative to students. The computer flashes spots of light around the screen, and if it is in the students field of view, he/she presses a key. The program intelligently tries to define the limits of the visual field. If fewer spots are sampled, the exercise can be completed in five to ten minutes and only one computer is needed for 15 to 20 students. This program should be combined with other measurements of human physiology.
253 2. The program VISION requires a series of graphics programs which are run by the HELLO program. If it is to be moved to a separate disk, be sure the necessary programs are also present.
VISION permits students to measure their field of view if the room is darkened (overhead lights wash out the top of the screen). The intensity of the screen is important because if the spots are too bright, persons with poor visual fields may still be able to see them. The number of spots tested is set at 120, but this can be altered when the program begins to run. When the program is finished, it will display the spots seen with a + , those missed with a box; a / indicates it was both missed and selected at different times. I currently attempt to interpolate the actual field of view based on the spots seen and missed. However, this routine is very slow and can be eliminated by inserting a line 4001 GOT0 4246 It is also possible to save the pictures on disk, but this is not a good idea for most labs since the disk will quickly fill up. To include this option, delete the line 4275 GOT0 4300
3. The exercise Vision testing is a routine part of any examination. However one aspect which is frequently overlooked is measurement of the visual field. Many diseases cause a gradual, progressive loss of peripheral vision (the ability to see to the sides) which is unnoticed by the patient. Checking the visual field is very tedious, so it is an ideal task for the computer,
Position yourself directly in front of the Apple computer's screen and cover one eye. A flickering spot will appear at the center of the screen and remain there through the test. Adjust your distance from the screen until the "twitching" spot is at the center of your vision, and the off center point of light disappears in your blind spot (about 4 inches from the screen). Don't worry if the dots are out of focus. Eliminate reflections of room lights from the screen, and be sure that the intensity of the screen is turned down.
Once in proper position, press "RETURN" and wait; be sure to keep your gaze on the twitching spot at all times. You will be shown approximately 150 spots located apparently at random around the screen. When you see a spot appear, press "RETURN" before it disappears. If you press the key when there is no spot on the screen, you will hear a beep. The points will appear at a more rapid pace, and remain on the screen for a shorter period of time, as you become used to the test. Depending on your peripheral vision and on the intensity setting of the monitor screen, you may be unable to see some of the peripheral spots. To insure that you have not shifted your position, your blind spot will be tested 9 times.
When the computer has tested you with enough points, it will give a double beep, then plot all the points that were tested. If you saw a point, it will be represented with a plus (+). If you missed the point, it appears as a box. Points which were both seen and missed at least once are designated with a slash (/). Your
254 maximum field of vision is then plotted by connecting the appropriate points. If the line appears at the perimeter of the screen, it indicates that your peripheral vision is excellent. Using the diagrams below, indicate approximately your field of view for the eye you tested.
Do these results appear normal?
E Measure body fat 1. General This program provides an alternative to using calipers for measuring fat and eliminates tedous and confusing calculations. Depending on student's age and sex, he/she measures the circumference at 3 locations on their body. The computer calculates the percent body fat. An interesting exercise can be constructed where this method is compared to results from use of calipers. The exercise can be combined with a variety of different physiological or health-related exercises. Only one computer is needed for 15 to 20 students.
2. The program BODY FAT also requires a series of graphics programs which are run by the HELLO program. If it is to be moved to a separate disk, be sure the necessary programs are also present.
3. The exercise Standard weight tables are misleading since they tend to underestimate weight. In addition, persons with well developed musculature will usually be classed as overweight because muscles weigh more than fat. However, our concern is not with weight, but rather the proportion of the body that is fat.
There are two indirect means of measuring fat; both are based on the principle that excess body fat is deposited just under the skin. The first method involves making a series of circumference measurements on the limbs and/or torso. The second method uses calipers to measure the thickness of the sub-dermal layer of fat. Both techniques have been calibrated according to the only accurate measurement technique; the method is based on Archimedes principle and involves weighing a
255 person in air and while submerged in water. Since fat is less dense than muscle, the difference between these two weights indicates the proportion of fat. Although the indirect methods are not as accurate, they are easier to perform.
To use the circumference method, follow the instructions given on the Apple computer. According to these measurements, what proportion of your body is fat? Does this agree with your assessment of your excess or lack of body fat?
Skinfold calipers are valuable to use since, if properly used, they differentiate between the underlying muscle and fat. Using calculations, the overall percentage of total body fat can be determined.
Four sites will be used: (1) triceps (halfway between elbow and shoulder on the back of the arm); (2) biceps (halfway between elbow and shoulder on the front of the arm); (3) subscapular (diagonal fold below shoulder blade); and (4) iliac crest (diagonal fold on the side, above the hip bone).
To do skinfold measurements, take hold of the skin and subcutaneous fat (no muscle) in the left thumb and forefinger; be sure you are measuring directly on the skin since clothing thickness can skew the results. With the right hand, slowly close caliper tongs around the fold and measure to the nearest .5 mm. Total the four readings from the calipers and use the appropriate table (below) to estimate your percentage of fat.
How do these readings compare to those obtained with the tape measure? Which method do you think would be more accurate? Why?
DETERMINING % FAT IN MEN DETERMINING% FAT IN WOMEN TOTAL % TOTAL % TOTAL % TOTAL % MM FAT MM FAT MM FAT MM FAT 20 9 90 27 15 15 60 31 25 11 100 28 18 16 63 33 30 13 110 29 21 17 64 34 35 15 120 30 24 18 69 35 40 17 130 31 27 19 72 36 45 18 140 32 30 20 75 37 50 20 145 32 33 22 78 38 55 21 155 33 36 23 81 39 60 22 165 34 39 24 84 40 65 23 180 35 42 25 87 41 70 24 185 36 45 26 90 42 75 25 200 37 48 27 93 43 80 26 210 38 51 28 96 44 54 29 99 45 57 30
256 F lung capacity 1. General This program simplifies calculations required to derive lung capacity using a Collins respirometer.
2. The program LUNG also requires a series of graphics programs which are run by the HELLO program. If it is to be moved to a separate disk, be sure the necessary programs are also present.
3. The exercise The exchange of oxygen and carbon dioxide between the body and the atmo- sphere occurs in the lungs. While evaluation of this transfer would be the most meaningful indication of the lung's health, it is difficult to measure directly. However, many conditions which affect gas exchange also influence both the amount of air that enters and leaves the lungs, and the volume of air that the lungs can hold. Thus, by measuring the capacities of the lung and the rate at which air can be moved into and out of the lung, an estimation of the lung's health can be assessed.
We will be using a spirometer to measure how much air is transferred between your body and the atmosphere. The device is an inverted bell which floats in a can of water; as you exhale air into the mouth piece, it enters the bell, raising it upward. The bell is attached to a pen so that as the bell rises, a pen moves down on graph paper. There are several volumes that we can measure.
lnspiratory Vital Reserve
Tidal Volume
Expiratory Reserve
1. Tidal volume is the amount of air that is inhaled and exhaled during normal breathing. 2. lnspiratory reserve volume is the maximum amount of air that can be taken in during exertion. 3. Expiratory reserve volume is the amount of air that can be exhaled during maximum exertion. 4. Vital capacity is the sum of these three volumes and represents the most amount
257 of air your body can consume. 5. Residual volume cannot be measured by spirometers. This is the amount of air that always remains in the lungs. 6. Total lung capacity is the sum of the residual volume and the vital capacity. Capacities for these measurements vary with a person's height, sex and age.
Typical values are: MALES FEMALES TIDAL VOLUME ...... 500 ml 400 ml INSPIRATORY RESERVE 3000 ml 2250 ml EXPIRATORY RESERVE .. 1000 ml 750 ml VITAL CAPACITY ...... 4500 ml 3400 ml RESIDUAL VOLUME ...... 1500 ml 1200 ml TOTAL CAPACITY ...... 6000 ml 4600 ml
With the valve near the mouthpiece open, gently press the respirometer chamber down to force out the trapped air, then raise it to bring in fresh air. Set the respirometer chamber so that the pen is about 1000 ml above the bottom of the graph. Remove a mouthpiece from the alcohol and wash it well before placing it on the spirometer. After taking a few breaths through the mouthpiece, close the valve so the chamber moves with each breath. Sitting erectly, inhale as deeply as possible and then exhale as completely as possible; record these two values. Using the Apple computer, enter your height, sex and age; then enter the value at maximum inspiration and that for maximum expiration. The program will compute your vital capacity, and then compare it to your predicted volume.
What is your tidal volume? What is your vital capacity? What vital capacity was predicted for you by the computer?
What percentage of the predicted value was your actual vital capacity? Considering values of less than 75% to indicate potential problems, are your values too low? If yes, then do you know of any personal health problems which could account for this?
G. Line fit 1. General LINE FIT is a simple statistics program that we use in the beginning classes to introduce the concept of variability. Students collect data, then enter the X and Y values in the computer. The program calculates the best fitting line to the data. The program can be combined with any exercise where data are collected.
2. The program LINE FIT is a free-standing program and can be transferred to any disk as needed.
258 3. The exercise No student hand-outs are provided.
H. cluster analysis 1. General CLUSTER ANALYSIS is a program that demonstrates simple taxonomic principles using calculations too complex for students to perform. The program's concept was described in great detail in The American Biology Teacher (47(1), 41 -47; 1985). Students first determine which are the significant characteristics that differentiate a series of objects or species (eg. number of leaveslnode, color etc). Then the characteristics of each object is quantified (plant A has 10 leaves/node and is red {red=1}, plant B has 1 leaf/node and is white {white=2} etc.). The data are entered in the computer and the program clusters the data by similarity. A dendrogram visually presents the relationships. Depending on the complexity of the data used, one computer could provide for the needs of ten to fifteen students. This exercise could be combined with manual methods of developing taxonomic keys.
2. The program Included with this program is the sample data file, HARDWARE.RAW DATA, which is referred to in the article. This version of the program does not permit students to save data on the disk. If you want to enter a data set and save it for students to process, do the following; press CTRL RESET as soon as the program begins to run. Type the number 7 and press RETURN, they type RUN and press RETURN. This deletes a program line that tells the program not to ask if the data should be saved.
3. The Exercise No student hand-outs are available. References:
Gealler, H. and G. Steele: The 1974 Probability Tables of Dying in the Next Ten Years From Specific Causes. Methodist Hospital of Indiana, Indianapolis, Indiana. 1974
Hall J. and J. Zwemer: Prospective Medicine Methodist Hospital of Indiana, Indianapolis, Indiana. 1979
Sweet, H. Non-Majors Biology Laboratory Notebook. Burgess Publishing Co. Minneapolis, Minnesota. 1984
Sweet, H. "The Use of Clustering Techniques by Students on an Apple Computer". The American Biology Teacher 47(1), 41-47; 1985
Sweet, H. "Use of a Computer as a Reaction Meter". LabStracts; Fall, 1986
259
Biology from the Human Perspective
Barbara Newman
Biomedical Sciences Department Southwest Missouri State 901 South National Avenue Springfield, MO 65804-0094
Barbara Newman received her B.S. Ed. and M.S. Ed. degrees from Southwest Missouri State University, Springfield, Missouri. Her areas of emphases are social studies, biology, chemistry and secondary education. Since 1986, Barbara has taught and/or been lab coordinator for general biology for non-majors, genetics, botany, zoology, anatomy and physiology. Currently, she teaches and coordinates laboratory sections for human anatomy and concepts in biomedical sciences. Her research interests include science education at the elementary and collegiate levels. She is currently developing a series of instructional videotapes for human anatomy.
261
This Material is Part of the Unit: "METHODS OF SCIENCE'
Objectives: 1) to realize need for tools to extend our senses: 2) to realize need for precise language in conveying ideas.
Laboratory Activity: the Black Bag Investigation
A. Problem: Each student will be given a sealed black plastic bag containing a number of objects, some of which may be fragile. Close your eyes and use only your sense of touch through the bag surface to determine what is in the bag without opening it or destroying its contents. Mentally assign numbers to each object and write word descriptions of the ob- jects next to each number. If possible, use these descriptions to identify the object. Identification of coinage and paper money as to denomination and country of origin entitles you to keep it!
Tentative descriptions and identifications: 1.
2.
3.
4.
5.
6.
B. Language is an important tool in conveying specific ideas efficiently. Choose three of the objects tentatively identified. Now assume that you are a caveman living about 10,000 BC, and using language relating to their experience describe each of the three objects. You may not use any term that a primitive man would not be familiar with (i.e., hard as metal, smooth and round like a marble). In what way does the caveman's description differ from yours? Object No. 1
Object No. 2
Object No. 3
263 Open the bag.
1. Was any object damaged by crude manipulation?
2. Was your touch sensitive enough to locate every object?
3. Did past experience bias your identifications?
4. In what way would you further investigate the natural objects?
5. Is there an item that requires more than one sense to identify it?
6. How does this activity compare to the scientist investigating a living cell, the ocean floor, outer space?
264 Steps in the Effective Establishment of Safety Rules for the Laboratory A Model Experiment
LABORATORY SAFETY This group activity, when carried out as described, will not only introduce students to some common laboratory safety rules, but can at the same time encourage students to cooperate in groups and serve as an introduction to the scientific method. (3-4 students per group)
LEARNING OBJECTIVES After this activity, students should be able to: 1) list at least six (6) common laboratory safety rules in their own words; and 2) set up an experiment following the scientific method.
PREPARATION Instructors should be familiar with common laboratory safety rules and procedures. Instructors should know the location of safety equipment such as the fire extinguisher and the first-aid kit. Instructors should be familiar with any specific institutional safety policies.
DEFINE THE PROBLEM The problem is to develop an effective way of teaching and reviewing laboratory safety rules.
RESEARCHING THE PROBLEM Review of the literature related to science teaching indicates little available data on this type of problem. (Discussion of safety devices evident in the lab -- signs, first-aid kit, fire extinguisher, exits, etc.)
IDENTIFY THE HYPOTHESIS Direct student involvement using group cooperative effort to produce a list of 10 common laboratory safety rules can be more effective in teaching and reviewing lab- oratory safety rules than other methods commonly used.
EXPERIMENT Test group -- Sections of BMS*110 labs taught by instructor X using the methods just presented. Control group -- Sections of BMS 110 labs taught by instructor X using any other method. The same laboratory exams with questions covering lab safety will be given to both groups.
OBSERVING AND STATING RESULTS The number of correct responses to lab safety questions will be tallied for both groups and compared.
FORMULATION OF A THEORY Probably not applicable due to the numerous uncontrollable variables. (Discuss several variables.)
* BMS (Biomedical Sciences)
265 STUDENT-GENERATED SAFETY SUGGESTIONS for use in the BMS 110 LABORATORY
1. Know the location of safety devices (fire extinguisher, first- aid, safety glasses, emergency exits and telephone with emergency number(s) EXT 5911. 2. No smoking in lab. 3. No "horseplay" in lab. 4. Be alert--know the correct way to use chemicals and equipment BEFORE using them. 5. Pay attention to and follow instructions for lab activities, especially when helpful suggestions are given during the course of the laboratory period. If you happen to be out of the laboratory, make sure that someone else covers for you if important instructions were given during your absence. 6. Keep the working area as clean and uncluttered as possible. Make sure area is clean and equipment is left in the proper condition at the completion of the lab period. 7. Be alert to personal safety, i.e., wash hands after lab, wear appropriate protective clothing, don't pick up broken glass with your hands, confine long hair and loose clothing. 8. When possible, work with a partner. 9. Ask for help when uncertain. 10. ENJOY LAB!
266 These Activities are Part of the Unit: "METHODS OF SCIENCE"
The material used for the following activity has been adapted from a 1975 NABT work- shop, "Biology Teaching and the Development of Reasoning, " developed by Anton Lawson, University of California, Berkeley, et al. It is patterned after Piaget's theories con- cerning the mental processes used in problem solving. Distinguishing between students using concrete and formal reasoning patterns is important in structuring laboratory experiences. The majority of concepts taught in biology are formal while the majority of students use concrete reasoning patterns to interpret them. (Lawson & Renner, 1975). Characteristics of students using concrete reasoning patterns include: the need for reference to familiar actions, objects, and observable properties; the need for step-by-step instruction in a lengthy procedure; and inability to recognize their own reasoning inconsistencies. Students who use a formal reasoning pattern can reason with concepts, relationships, abstract properties and theories, and use symbols to express ideas. Given certain overall goals and resources, they can plan a lengthy procedure and they are critical of their own reasoning. The series of problems should be reviewed with particular emphasis given to the writ- ten explanations requested at the end of each problem. The first problem illustrates the student's ability to use combinatorial logic--the ability to link a set of associations or correspondences with each other in many possible ways. The second problem explores the reasoning processes used to separate and control variables. Isolating variables is difficult for many beginning biology students and is critical to many introductory exper- iments. The third problem deals with proportional reasoning. Can the student recognize and interpret the relationship between relationships?
Objectives: 1) to stimulate thinking about problem solving; 2) to introduce methods of problem solving; 3) to assist the instructor in discriminating various thought patterns used by the students, i.e., concrete or formal reasoning.
A. The Fruit Problem A small population of individuals isolated on a Pacific Island have a diet limited to the fruit of four different plant species. You as an investigator wish to study the effects of their diet on longevity. In preparation for this study, you'll want to compile a list of all the possible combinations of these fruits in their diet. Write down each possible combination of fruit. Use the letters to save space: Eam = E Nassel = N Arro = A Volam = V
Looking back, how did you approach this problem: Did you think at once of a way to do it? Did you first think of a way that had to be modified?
267 B. The Mealworm Problem An experimenter wanted to test the response of mealworms to light and moisture. To do this, he set up four boxes as shown in the diagram below. He used lamps for light sources and constantly watered pieces of paper in the boxes for moisture. In the center of each box he placed 20 mealworms. One day later he returned to count the number of mealworms that had crawled to the different ends of the boxes.
dry wet dry wet
I I ;::-- - - I I,
I" I - dry dry wet wet
The diagrams show that mealworms respond to (response means move toward or away from): a) light but not moisture c) both light and moisture b) moisture but not light d) neither light nor moisture
Please explain your choice.
268 C. The Frog Problem* Professor Ranidae, a herpetologist, conducted an experiment to determine the number of frogs that live in a pond near the field station. Because he could not catch all of the frogs, he caught as many as he could, put a white band around their rignt front legs, and put them back into the pond. A week later he returned to the pond and again caught as many frogs as he could. Here is the Professor's data: First trip to the pond: 55 frogs caught and banded Second trip to the pond: 72 frogs caught. Of those 72 frogs, 12 were banded. Estimate the total number of frogs in the pond. Total =
The professor assumed that the banded frogs had mixed thoroughly with the unbanded frogs, and from his data he was able to approximate the number of frogs that inhabit the pond. If you can compute this number, please do so. In the space below, explain how you calculated your results.
* This problem can be made more interesting by using a population of 'pill' bugs housed in an aquarium containing sphagnum. Sample the population, mark the sample with nail polish, return them to the tank. Take the second sample a week later and estimate the population. However, the problem will no longer as clearly separate the concrete vs formal reasoners.
269 This Material is Part of the Unit: "METHODS OF SCIENCE"
Objectives: 1) to improve skills in critical reading; 2) to improve skills in evaluating evidence; 3) to create awareness of the difference between fact and inference; 4) to learn specific meaning of common words used in science.
Below is a statement of facts. Following that is a list of inferences supposed to have been drawn solely upon the basis of the facts given. Mark with the letter C the infer- ences that may be correctly drawn solely from the facts given; with the letter N those that are not proved solely by the facts given.
FACTS: A grain of corn and a marble were placed upon moist blotting paper in a glass dish in a warm (20°C) dark room. Another grain of corn and a marble were placed upon moist blotting paper in a glass dish in a warm (20°C) sunlighted room. Both grains germinated (the embryos grew). This experiment was repeated many times with the same results.
INFERENCES: 1. Grains of corn will germinate on moist blotting paper. 2. Corn grains will not germinate unless they are in glass dishes. 3. Moist blotting paper is necessary for germination of grains of corn. 4. Light is not necessary for the process of growth. 5. The presence of a marble is necessary for germination of grains of corn. 6. Moisture is necessary for germination of grains of corn. 7. All seeds will germinate in both light and darkness. 8. Glass dishes do not prevent germination of grains of corn. 9. The presence of a marble is not necessary for germination of grains of corn. 10. Light has no effect whatever upon growth of embryos of grains of corn. 11. Light does not prevent germination of grains of corn. 12. Glass dishes have no effect whatever on germination of grains of corn. 13. Heat and moisture increase growth of embryos of grains of corn. 14. Grains must have heat to germinate. 15. Grains of corn germinate in either light or darkness. 16. Grains of corn must be placed in moisture in a warm room to germinate. 17. Light is not necessary for germination of grains of corn. 18. A grain of corn placed on moist blotting paper in a wooden dish in a warm (20°C) dark room will germinate.
270 Laboratory Activity and Assignment: Certain common words in science have specific meanings that are important in conveying ideas. Similar words tend to be confused with these by the novice. Learn the definitions and distinctions between the following "confusing word pairs". Write a sentence that illustrates the correct usage of the term.
Al. Accuracy - the degree of correctness of a measurement or a statement. Accuracy means closeness to the true or established value or truth. (a result) A2. Precision - the degree of refinement with which a measurement is made or stated. Precision implies "repeatability" or "obtaining the same value over and over with repetition". (an action)
B1. Qualitative (an adjective) - relating to distinctions of type, not amount. Ex. A qualitative assay procedure would indicate what different types of com- pounds are present. B2. Quantitative (an adjective) - relating to a measurable amount or quantity. A quan- titative procedure is concerned with the measurement of phenomena, either amount of mass or number of items. Ex. A quantitative assay procedure would indicate how much of a particular substance is present.
C1. Error - the difference between a measured, observed, or calculated value and the real or true value. Errors are usually random and more difficult to detect or minimize than mistakes. (result, the distance from accuracy) C2. Mistake - a mistabulation or miscalculation of figures. Mistakes can be corrected by checking each item and computation. (action)
D1. Analytical Technique - a technique designed in scope to determine the quantitative and/or qualitative aspects of a small sample, usually for assessment. Analytical techniques use samples of a larger amount to determine what is in that larger amount. ("taste test"!) D2. Preparative Technique - a technique designed in scope to separate qualitative aspects of a mixture, usually for purposes of isolation or purification. Prep- arative techniques use as large a sample as permitted by the particular technique so as to recover as much of the separated material as possible. (wine making! ; ultracentrifuge)
El. Fact - A real event observable in past time that can be described without mental elaborations. E2. Inference - a deduction or conclusion drawn from facts. Inferences tend to be more mentally predictive as to future events and situations and do not describe real events occurring in past time.
271 This Material is Part of the Unit: "SCIENTIFIC MEASUREMENTS"
The occupational metric problems are adapted from ideas in: Metric Measurement in Food Preparation and Service, Lynne Nannen Ross, Registered Dietitian, Iowa State University Press, 1978.
PROBLEM: Brownies are on the school lunch menu. The cook must prepare one brownie (5cm X 5cm) for each of the 200 children. The new British cook is familiar only with metrics. 1. Convert the recipe to metric. 2. Convert the baking temperature to Celsius degrees. 3. Determine the (minimum) capacity of the bowl needed for mixing the ingre- dients. 4. Determine the size of pan or pans needed for baking. (NOTE: The oven dimensions are 2 feet X 3 feet) 5. Brownies must be ready by 11:30 a.m.! TODAY!!!
Recipe for Brownies ounces unsweetened chocolate Temperature: 350°F ( ºC) cups shortening cups sugar Time: Bake 30 minutes medium eggs cups flour Amount: 200 5cmX5cm squares tsp baking powder tsp salt tsp vanilla cups broken nutmeats
ANALYSIS OF THE PROBLEM: 1. The total volume of the brownie batter will be liters. 2. That volume of batter will require a mixing bowl with a minimum capacity of gallons. 3. How many servings are to be prepared 4. What size is each serving? cm X cm. 5. How many square centimeters of brownies will be needed? 6. What size pan(s) will the oven accommodate? cm2 7. What pan or pans will accommodate the brownie batter and fit into the oven?
CONVERSION AIDS: 1 tsp = 5 ml 1 oz = 28g 1 cup = 250 ml(g) ºC = 5/9(ºF-32)
POSSIBLE SOLUTIONS: 1. Prepare brownies in 2 batches using a liter bowl and bake in cm X cm pan. 2. Prepare brownies in a liter mixing bowl and one cm X cm pan or two cm X cm pans.
272 PROBLEM: The mobile clinic serves several communities in southwest Missouri. You must try to spend a minimum of two hours at each of the 14 small town clinics each week. Is it possible to serve each of the 14 clinics two hours each week? If not, how many clinics can you visit in a forty-hour week? Your home and office are in Springfield. ANALYSIS OF THE PROBLEM: 1. What is the speed limit in Missouri? 65 m/hr = km/hr 2. What is the speed limit on winding roads? 45 m/hr = km/hr 3. What is scale on maps? 1 cm = km 4. How is distance converted to time? Determine kilometer distance and divide by kilometers/hour. Multiply the decimal part of an hour by 60 to find minutes.
POSSIBLE ITINERARIES: 1. MON: Springfield Nixa Highlandville Spokane Return to Springfield TUE: Springfield Reeds Spring Cape Fair Cassville Return to Springfield WED: Springfield Purdy Monett Aurora Return to Springfield THR: Springfield Marionville Crane Elsey Return to Springfield FRI: Springfield Billings Clever Return to Springfield
2. MON: Springfield Billings Clever Nixa Highlandville Return to Springfield TUE: Springfield Spokane Reeds Spring Cape Fair Cassville stay WED: Cassville Purdy Monett Aurora Marionville Springfield
THR: Springfield Crane + Elsey Springfield
EVALUATION OF ITINERARIES: Using a 15 centimeter ruler and the following information, determine: 1. the distance between towns 2. the time required to travel those distances 3. the cost of traveling each route map: 1 cm = 6 km car averages 40 miles/gallon 1.6 km = 1 mile gas averages 75 cents/gallon lodging averages 25 dollars/night
RECOMMENDATION: The most efficient route for time and money: Route 1 hours cost Route 2 hours cost Other hours cost
273 274 THE METRIC SYSTEM REVIEW
1. Some things that won't change (much)! Replace the English term with the most appro- priate metric term. Identify your calculations by number and letter on the bottom and back of the pages.
a. milestone -- 10-gallon hat foot-long hot dog pound cake inch worm an ounce of prevention is worth a pound of cure don't budge an inch give him an inch, he'll take a mile Denver is the Mile High City
2. a. What is the temperature for boiling eggs? ºC b. What is normal body temperature? ºC c. Comfortable room temperature is 76ºF or ºC
3. What is the numerical value indicated by the following prefixes? Example: micro = 0.000001; hecto = 100 a. kilo = b. deci= c. milli = d. centi = e. deka =
4. Approximately (within reason!) how many? liters in a gallon of gas liters in a quart of milk milliliters in a cup of coffee grams in a quart of milk kilograms in 5 pounds of flour yards in a football field kilometers in a football field inches longer is a cigarette that has an extra millimeter cubic centimeters in a 10-gallon hat dollars to make "megabucks"
5. One mile equals 1.6 kilometers, so 65 miles per hour now reads kilometers per hour.
275 This Material is Part of the Unit: "MICROSCOPE USE AND CARE"
Objectives: 1) to introduce the topic of microscopy; 2) to demonstrate how lenses work; 3) to increase the working vocabulary; 4) to promote the use of inquiry in investigation.
The microscope is an important optical tool that performs two functions in biological studies: 1) The microscope increases the ability to resolve details that cannot be observed be- cause of the limits of our own biological optical sense organ, the eye. To resolve is the ability to separate and make visible the individual parts of an image i.e., the headlights of an approaching vehicle appear as one until they are within our ability to resolve them as two. The human eye resolves spaces between two real ob- jects if they are more than .2 mm apart. If objects are closer together than this, our eyes see them as touching. We cannot resolve biological cells with our eyes alone as most cells are smaller than .2 mm and average 0.02 mm in diameter. The primary function of the compound microscope is to increase the resolving power (or resolution). The degree of resolving power is determined by the quality of the lens system and the light used. The practical limit of resolution of any optical system using white light is about 0.0001 mm or 0.1 µm (1 micrometer = 0.001 millimeter).
2) The second function of the microscope is to increase the apparent size or to magnify the object being resolved. Magnification produces an image that is large enough for us to resolve when cast upon the retina of our eye. Because our eyes cannot resolve objects below .2 mm, the microscope lens system must magnify objects so that their apparent size is at least as large as this. Magnification is achieved by refraction of light on curved lens surfaces. The pheno- menon of refraction causes a bending of light as it moves from one curved transparent substance (glass lens surface) into another substance (air). The amount of magnifica- tion is determined by the arrangement and curvatures on the surfaces of the lens system of the microscope. The measure of magnification is determined by how many diameters the objects image appears to increase. For example, if the real diameter of an object observed under the microscope os 20 micrometers and its image appears to be 2.0 millimeters, it is said to be magnified 1000 diameters or 1000 times or 1000X. When using the compound microscope, the total magnification of an object is determined by multiplying the magnification value of the ocular lens by the magnification value of the objective lens.
EXAMPLE: Ocular 10X times objective 45X = 450X (total magnification)
The relationship between the function of resolution and magnification is an important one. One can continue to magnify an object and reveal new detail and information only to a limit determined by the resolving power of the lens systems of the instrument. For example, recalling the resolution of the eye at .2 mm and the maximum resolution of the microscope at 0.1 micrometers, it is easy to calculate the maximum useful magnification of 1000X with a good optical system. it is possible to produce lenses that magnify greater than this but the images become "fuzzy" and no new information can be seen. Using magni- fication greater than the resolving power of the instrument can support results in "empty magnification". The lack of "crispness" in many photographs taken through a microscope occurs because of empty magnification.
276 "MICROSCOPE USE AND CARE" continued
Laboratory Activity: Investigations of a micro-scope ("little viewer") This exercise does not require the microscope, so place it aside to make a work area. You will be given two sandwich packets containing a piece of plastic wrap. Do not touch the surface of the plastic wrap with your fingers or you may change its interaction with water in this investigation. Note that the bottom inside cover of the paper wrap has a series of letter "e's" that can be seen through the plastic wrap. Lay the #1 sandwich on the desk top and fold back the top paper cover to the letters in the inside cover can be observed without disturbing the sandwich. Beginning with the second letter from the left, add a drop of water (using a droper pipet) to the plastic and look carefully at the letter "e" through the water drop. Compare the first dry letter to the second letter "e". Are there any differences? Repeat the procedure placing two drops of water over the third letter "e" and four drops over the fourth letter. What differences to you observe? Do all the letters appear the same size? Compare the size of the water "puddles" and the curvature of their surfaces by observing them from the top and from the side. Which letter appears to be magnified most? Which water puddle has the most depth, i.e., the greatest curve? What part of the microscope does the water represent? Remember that light is refracted (bent) as it passes from one medium into another. When passing through a convex surface (rounded outwardly), the rays are bent inward. When the light rays pass through a concave surface (rounded inwardly) they are bent outward. The objective of the microscope collects light rays coming from the object being studied. Are light rays coming from the letter "e" passing through a convex or concave surface? Using the above information, draw a lateral (side) view of the one-, two- and four-drop puddles that illustrates how refraction and magnification are related to the curvature of the water surface. How could you increase the magnification (distance of drop from letter "e") of the letter "e" using these same materials? Use the second sandwich to experiment.
277 This Activity is an Excerpt from the Unit: "FUNDAMENTALS OF THE RESPIRATORY AND EXCRETORY SYSTEM"
Objectives: 1) to identify the organs of the excretory system and their relative positions within the human; 2) to identify structures and learn their functions within the kidney; 3) to demonstrate an understanding of the nephron by assembling one.
Activities: 1) Locate and identify organs of the excretory system on human torso model. 2) Using pork kidneys and kidney model, identify structures in the kidney. 3) Using prepared slides of kidney tissue, locate ultrastructures. 4) Using a collection of items given, assemble a nephron*
*ITEMS: (2) 40 cm strands of 2 shades of red yarn knotted together at intervals (1) Thistle cup (1) 20 cm piece of glass tubing bent into U-shape (1) 15 cm convoluted tube (1) 25 cm convoluted tube
The assembled nephron should include the: afferent arteriole loop of Henle flomerulus distal convoluted tubule Bowman's capsule efferent arteriole proximal convoluted tubule peritubular capillaries
Suggest to the students that they assemble the filtration apparatus first, then place the vascular component.
278 These Activities are Excerpted from the Unit: "ANATOMY AND RESPONSES OF THE NERVOUS SYSTEM"
Objectives: 1) to understand the processes of simple versus choice reaction time; 2) to demonstrate the effect of learned reflexes on reaction time; 3) to demonstrate the effect of cold on reaction time; 4) to demonstrate left or right side dominance; 5) to demonstrate the importance of binocular vision for depth perception. 6) to investigate how fine a discrimination can be made in judging weights.
REACTION TIME: Procedure: Student holds thumb and forefinger of his preferred hand about 1 cm apart while the experimenter suspends a new dollar bill vertically between them, with the center of the bill level with the thumb and finger. Student is to grasp the bill as soon as the experimenter releases it. (Most students are unable to do so.) Now the student holds a meter stick between the thumb and index finger; then releases his grip and then as quickly as possible tries to grasp it again. The distance the stick travels before it is stopped is measured. Repeat several times. Average dis- tance the stick falls is a measure of reaction time. Compare your results with several other students. Repeat the test after chilling your hands in ice water. What effect does this have on the results? Why?
Simple and Choice Reaction Times: To show the differences between simple and choice reaction time, line up 10 students, single file, with each person placing his right hand on the shoulder of the student in front of him. The first person raises his arm so that it can be seen by all. Students are told that each person will be tapped on the shoulder and that he is to respond as quickly as possible by dropping his arm. Instructor then taps the right shoulder of the last person in the line, starting a stop-watch at the same time, and stopping it when the last arm in line drops. Time ten trials. Average the results and compute the mean time taken for each person to respond. This is the simple reaction time. To measure choice reaction time, subjects place both hands on the shoulders of the person in front. This time subject is to respond by tapping the shoulder opposite the one which received the stimulus, but the location of the stimulus will be unknown in advance and will vary from trial to trial. Make ten trials and compare results with those obtained for simple reaction time. Which reaction time is faster? What are the neural explanations for the differences in the two situations?
EYE DOMINANCE: Procedure: Cut a 3.5 cm diameter hole in a large sheet of cardboard. Using both eyes, sight a distant object through the 2.5 cm hole while the cardboard is held at arm's length. Hold the cardboard with both hands and move it gradually toward the face, keeping the distant object sighted. Determine which eye is used. Repeat several times. Do you always use the same eye to keep the object in sight? Is your dominant eye on the same side as your dominant hand? The dominant food-chewing side of the mouth? Make a record of eye, hand, and side-of-mouth dominance for your laboratory group. Does a consistant pattern emerge? With family?
279 DEPTH PERCEPTION: Procedure: Prop a large sheet of white cardboard (60 cm X 90 cm) about 5 meters in front of the class, and several feet from the front of the room. The top of the cardboard should be about 10 cm above eye level of students. Trial #1: Hold a strip of black paper 1 cm wide and 30 cm long, 10 cm behind the large white cardboard, with 5 cm showing above the card. Students are now to judge the distance between the card and the black strip under four conditions: 1) looking with one eye, holding the head still; 2) looking with one eye, moving the head back and forth; 3) looking with both eyes, holding the head still; and 4) looking with both eyes while moving the head back and forth.
Trial #2: Repeat the above procedure with the strip 30 cm from the card.
Trial #3: Repeat at 2.5 cm.
Now repeat Trials 1 through 3, but use a pencil instead of the black strip. Hold the pencil so that 5 cm shows above the card, use the same distances, but vary the order. Each student records his judgments (16 for black strip and 16 for the pencil trials). After the trials are completed, students are told the actual distances and asked to compute their errors. What is the average error for the entire class? Which conditions produced the poorest judgment? Which the best? Were there any differences between accuracies of judgment with the black strip and for the pencil? Why? (Discuss binocular vision)
JUDGING WEIGHTS: Procedure: To determine the limit of man's ability to judge whether two objects are of different weight -- Student closes his eyes and holds two 125 ml Erlenmeyer flasks, one in each hand. One is marked Control and the other Test. Each containg 50 grams water. Student reports if he considers them to weigh the same. Then the experimenter adds water to the test flask in 5 ml portions, each time handing the flasks back to the student but randomly mixing the right and left and each time asking the student to determine relative weights. When the test flask feels heavier, record the weight of added water. The difference between the weight of the test and control flask weights is the "difference threshold" for 50 g. Repeat this procedure, starting with 100, 200 and 500 g of water (using larger flasks). Be sure that each pair of flasks are of equal weight. How fine a discrimination can be made in judging weights? Does the difference threshold vary according to the initial weights used? Make a graph, plotting weight of control flask on the horizontal axis and difference threshold on the vertical axis. Do the results vary much from student to student? What are some variables that could affect the weight discrimination?
m w m
m
I w
WEIGHT OF CONTROL FLASK
280 The instructional periods of genetics fundamentals are followed by discussion of human genetics and problems concurrent to them. This attitudinal survey is a tool that is used as an aid for introspection and to stimulate discussion of ethics and genetic technology.
ETHICS AND GENETIC TECHNOLOGY AN ATTITUDINAL SURVEY
INSTRUCTIONS: The following statements concern ethical and moral decisions relative to genetic diseases. Circle A of you agree or agree more than disagree with the statement. Circle D if you disagree or disagree more than agree with the statement. Circle U if you have no tendency to either agree or disagree with the statement. Please respond as honestly as you can. Hemophilia is an inherited disease in which trauma produces excessive bleeding because there is a deficiency of one of the clotting factors. Death usually occurred in childhood until the recent institution of therapy consisting of frequent injections of highly con- centrated and purified clotting factor. Only males have this problem because the abnormal gene causing the deficiency is on the X chromosome. Before this new therapy was available, males usually died before they married and had children. Every daughter of a male with hemophilia will be a carrier of the gene and will pass it on to half her sons. The sons of hemophiliacs are not affected. The treatment of this disease currently costs $10,000 per year. Below are listed some actions that might be taken with regard to reproduction and treatment of men with this disease: 1. Males with hemophilia should be sterilized so they won't pass on the gene. A D U 2. Since the sex of offspring can be determined by amniocentesis early in pregnancy, all female offspring of hemophiliacs should be aborted. A D U 3. Female offspring of males with hemophilia and all females known to be carriers of the gene for hemophilia should be sterilized. A D U 4. The sex of the fetuses of known female carriers of hemophilia should be identified by amniocentesis and all male infants should be aborder. (There is currently no reliable test for hemophilia in the fetus). A D U 5. Since treatment is available, no intervention in reproduction should be allowed. A D U
6. The parents should always decide what action should be taken in any case. A D U 7. Since the treatment for this disease is so expensive that the state will usually have to pay for it, society (State or Federal Government) or insurance companies should decide what action should be taken. A D U With the new techniques in biochemistry, cell culture and chromosome analysis, it is often possible to detect inherited conditions soon after birth or in utero before 20 weeks of pregnancy. It may soon be possible to detect some inherited disorders that don't show up until early or even late adulthood.
281 Genetic screening tests have been used in some cases and might be used in others to detect these conditions. Please indicate in which of the following conditions or situations you feel genetic testing should be MANDATORY. 8. Screening tests for inherited disease should never be mandatory. A D U 9. Screening tests for inherited disease should sometimes be mandatory depending on the situation. A D U 10. If the screening test will detect an inherited metabolic disorder at birth affecting one in 15,000 newborns which if untreated may result in mental retardation, but which can be treated by a special diet. (Example: phenylketonuria) A D U 11. If the screening test will detect an inherited blood disorder affecting one in every 500-600 U.S. Blacks for which there is no effective treatment but whose Symptoms (intermittent pain, recurrence of infections) physicians feel can be ameliorated and whose lives can be prolonged from 25 to 30 years to 30 to 40 years. It costs about $4,000 to $5,000 per year to treat. (Example: sickle cell anemia) A D U 12. If the screening test will detect a complex inherited disease affecting one in every 2,500 whites, causing a debilitating disease of lungs and digestive tract. The usual life expectancy is for about 20 years. Doctors don't know whether or not life can be prolonged by medical therapy. Treatment costs about $4,000 to $5,000 per year. (Example: cyctic fibrosis) A D U 13. If the screening test will detect a nervous disorder that affects the individual from age 30 onwards in which there is gradual loss of control of hands, feet, chewing, swallowing and finally brain function. Persons usually die in mental institutions. No treatment is available. Each child of these individuals has a 50-50 chance of inheriting this condition and they are usually born before the person knows they have the disease. (Example: Huntington's Disease) A D U 14. If the screening test will detect sex chromosome abnormalities at birth which occur in about one in 500-600 infants (mostly in males) and these can be partially corrected by surgery and/or hormone treatment. (Example: XXY, Klinefelter's Syndrome, which in males produces slight breast development, small testes and sterility) A D U 15. If the screening test involves amniocentesis (taking out fluid and cells from around the fetus) in women over 38 years and detection of severe chromosomal disorders that are associated with birth defects and mental retardation. These cannot be treated and if found, fetuses would be aborted. A D U 16. If the screening test detects normal people who carry an inherited disease which has a 25% or 1 in 4 chance of being inherited by the children if two carriers marry. Fetuses with the disease can be detected in early pregnancy and aborted if the parents wish. There is no treatment for the disease which leads to mental deterioration and death by 3-4 years of age. A D U
282 17. If the screening test detected at birth an abnormality of fats in the blood that markedly increases the chance of a heart attack in men (and, to a lesser extent, women) before age 50 and in which the -effectiveness of a low fat diet, which would have to be followed throughout life, was unknown. (Example: type II Hyperlipopro- teinenia)
18. If the screening test detects normal people who carry a gene for an inherited blood disease which cannot be detected in early pregnancy by amniocentesis and which cannot be treated. If two carriers marry, each of their children will have a 25% or 1 in 4 chance of inheriting this disease. (Example: sickle cell anemia) A D U 19. Society should determine the type or kind of disease to be tested for in genetic screening. A D U 20. Society (government, institutions) should determine who should be screened for genetic diseases. A D U 21. Individuals should determine whether or not they wish to be screened. A D U 22. Every effort should be made to eradicate genetic disease. A D U 23. It is more important to control diseases that can be treated than to try to control hereditary diseases.
24. It is essential to pass on genetic information to relatives who might inherit or have children with the condition even if the person who has the disease objects strenuously. A D U 25. Before marrying, it is very important to know whether or not they are carriers of or have a family history of an inherited disease. A D U In each case below, legitimate rights are in conflict. Indicate your agreement or dis- agreement in each case with which rights should predominate. 26. The freedom of the individual predominates over his or her responsibility to society. A D U 27. The rights of the parents predominate over the rights of their children. A D U 28. The rights of the parents predominate over the rights of the fetus A D U 29. It is better to have no life at all if it is not of a reasonable quality. A D U 30. The rights of parents are more important than their obligation to future generations. A D U 31. The rights of persons with normal intelligence predominate over those who are mentally retarded. A D U
283 Those individuals who are willing to consider abortion under certain circumstances should indicate whether they agree, disagree or are uncertain about choosing to abort a fetus who has an inherited condition that would result in the following situations if he or she is born. 32. The child will die at birth or within a few days, no matter what measures are taken. (Example: anencephaly, in which the brain hemispheres don't develop) A D U 33. Development is normal for about six months after birth then there is retrogression of brain function and death by 3 years of age. (Example: Tey-Sach's disease) A D U 34. After birth there is a life expectancy of 18 or more years with complicated and expen- sive medical treatment with a normal but restricted life. (Example: cystic fibrosis) A D U 35. Life is normal until 35-40 years of age then Huntington's disease begins (a progres- sive loss of control of arms, legs, chewing, swallowing and brain function with bi- zarre movements. Death in a mental institution). A D U 36. Life expectancy is relatively normal but there is severe mental retardation requiring guardianship at home or in an institution.
37. Life expectancy is normal but there is some retardation in which the individual can be trained and work in a sheltered workshop.
38. Life expectancy is normal with normal intelligence but the individual is markedly incapacitated. (Example: paraplegic without control of urine or bowels as in spina bifida - failure of normal closure of the spinal column).
39. Life expectancy and intelligence are normal but the fetus is a sex not wanted by the parents. A D U 40. Life expectancy and intelligence are normal but fetus just not wanted by the parents. A D U 41. The recurrence risk for an hereditary condition is the chance that it will occur a second time if it has occurred once. It is sometimes used to indicate the chance that a condition will occur at all. What is the maximum (highest recurrence risk that you would be willing to take for a serious genetic condition in one of your children? (Circle your choice)
284 BMS 110: Biology From the Human Perspective Laboratory Topics Listing
*Unit 1. Methods of Science Unit 2. Scientific Measurements Unit 3. Microscope Use and Care Unit 4. Cell Observations Unit 5. The Procaryotic Pl an Unit 6. The Eucaryotic Plan Unit 7. Cell Reproduction Unit 8. Membrane Properties Unit 9. Enzyme Activity Unit 10. Gross Vertebrate Anatomy Unit 11. Digestive Systems Unit 12. Nutrition and Energy Unit 13. Respiratory/Excretory Unit 14. Metabolic Rates Unit 15. Circulatory System Unit 16. The Nervous System Unit 17. Reproduction/Meiosis Unit 18. Developmental Biology Unit 19. Mendelian Genetics Unit 20. Mendelian Genetics continued Unit 21. Genetic analyses Unit 22. Biodiversity Unit 23. Principles of Ecology Unit 24. Ecosystems and Energy Flow Unit 25. "Extinction"
Appendix A. Tools and Instruments
* The following activities were taken from Units 1, 2, 3, 13, 16, 20
285 General Biology for Elementary Education Laboratory Topics
1. Introduction: Plants and Animals in the Classroom Discussion of Semester's Activities Begin Library Survey and Resource Search 2. Animal "sets"/Missouri Animal Diversity, "Invent An Animal" (OBIS) ** "What Animal Am I?" (Sharing Nature with Children) 3. "Sassafrass" Ozarks Ecology (Multimedia production) or Hillcrest Field Trip: Habitats, "Litter Critters" (OBIS) 4. Community Interaction, "Nothing Lives Alone", "Microtrails" (Acclimatizing) Film: "More Than Trees" (Missouri Conservation) 5. Introduction to Microscopy and Cytology 6. Survey Kingdoms Monera and Protista/Library Survey due 7. Plant "sets"/Plant Diversity/Begin Plant Projects 8. Plant Morphology, "Tree Walk", "Invent A Plant" (OBIS) 9. Plant Physiology, "Desert Water Keepers" (OBIS), "Gift of Life" 10. Plant Reproduction and Growth, "Grocery Bag Botany" 11. "Functioning Human": Reception and Motor Responses 12. Human Circulatory and Respiratory Systems/Plant Projects due 13. Human Digestive System/Nutrition/Film, "Snack Facts" 14. "Urogenital System/Heredity: Why You Look Like You Whereas I Tend to Look Like Me" (Charlotte Pomerantz) 15. Animal Life Cycles: Comparative Development The Caterpillar and the Polliwoq (Jack Kent)
Goals: 1. Develop a positive attitude toward biological science. 2. Learn content sufficiently well to be comfortable teaching biology concepts. 3. Learn biology activities to support those concepts and that may be used in the ele- mentary claassroom.
** OBIS (Outdoor Biology Instructional Strategies)
286 General biology for teachers in elementary education is designed with the conventional lecture-lab format. The three hours/week of lecture are traditional but the three-hour lab is eclectic. There are three primary objectives for the lab: 1) to gain an understanding of the basic biological concepts; 2) to develop a file of resources available to teachers; and 3) to learn various activities useful in the elementary science classroom. Where possible, lab activities used to explore biological concepts are presented in a manner that can be adapted to the elementary classroom. No published manual meets these objectives, so each lab is designed to meet the particular need. A variety of outside activities are assigned to broaden the scope of the lab experience. The library survey requires that each student select five biological concepts, locate two different resources that can be used to teach them, and prepare a reference card for each. This activity initiates familiarity with the curriculum center and an exposure to available literature. As part of the resource search, the students must obtain resource materials from five off-campus sources such as government publications, state departments, county extension, manufactureres, service agencies, etc. in an area of their interest. Following discussion of scientific methods and measure, each pair of students is given a mini-experiment or investigation to complete. A scientific report and a brief oral presentation conclude that activity. Later, the students are given a container, soil, and packet containing three kinds of seeds to plant. They are asked to make observations and keep growth records. The project is complete when all three types of seed have germ- inated and at least one reaches fifteen centimeters in height. They are quite proud of this. Pets in an elementary classroom are quite common and sometimes disastrous. To enable students to gain first-hand knowledge of pets, their life cycles, requirements, etc., there are aquaria of fish (native) and turtles, terraria of salamanders and lizards, and cages of gerbils and guinea pigs in the classroom. On occasion there are also snakes, tadpoles and insects. The students are each assigned a week to care for the animals and water plants. One-time extra credit is available for designing and crafting a bulletin board illus- trating a biological concept. Creation of an attractive and informative display requires an understanding of the concept. Outdoor activities include identification of native trees on campus, fifteen spring (or fall) wildflowers and fifteen Missouri birds. A three-hour field trip to a local nature trail provides opportunity to study several plant communities -- cedar glade, creek area, oak hickory woods and prairie. This activity follows discussion of plant communities of the Ozarks area. The activities selected from Acclimatizing, Sharing Nature with Children and Outdoor Biological Instructional Strategies usually require little time to complete. The effect-
287 iveness for some of them lies in their brevity. Others are easily adapted to various levels of difficulty as well as time span. Some, especially OBIS, may be combined to compliment an entire unit. The lab is very eclectic depending upon the semester's students. Each planned activity (or most of them) may be completed during the semester. New ideas are explored and original themes modified. The schedule of topics is reversed from fall to spring to accommodate the seasons.
288 General Biology for Elementary Education Address List for Resource Material
Below is a list of some governmental agencies and private industries who offer free mater- ials on request. Many of these are appropriate for use in the elementary classroom. This list is given to you as a starting point and not at all intended to delineate the only resources available to you. As a courtesy, in making your requests, please be specific as to the topic and age level you are trying to reach.
American Bakers Association, 1700 Pennsylvania Avenue NW, Washington, DC 20006 American Dental Association, 211 East Chicago Avenue, Chicago, IL 60611 "Learning About Your Oral Health" American School Food Service Association, 4101 East Iliff Avenue, Denver, CO 80222 The American Humane Association, List of Educational Materials, 5351 South Roslyn Street, Englewood, CO 80111 American Humane Education Society, Department BA, 350 South Huntington Avenue, Boston, MA 02130 American Institute of Baking, 400 East Ontario Street, Chicago, IL 60611 American Medical Association, Order Department, P. 0. Box 821, Monroe, WI 53566 "Your Body and How It Works" Animal Welfare Institute, P. 0. Box 3650, Washington, DC 20007 "First Aid and Care of Small Animals" Arby's, Inc., Consumer Affairs, One Piedmont Center, 3565 Piedmont Road NE, Atlanta, GA 30305 "Eating Right Is Easy" Chevron Chemical Company, P. 0. Box 3744, San Francisco, CA 94119 A Child's Garden by Lou Czufin Consumer Information, Pueblo, CO 81009 Department of Health and Human Services, 5600 Fishers Lane, Rockville, MD 20857 Division of Health of Missouri, Broadway State Office Building, P. 0. Box 570, Jefferson City, MO 65102 "A Boy Grows Up" and "A Girl Grows Up" The Garden Club of America, 598 Madison Avenue, New York, NY 10022 "The World Around You - An Environmental Packet" Kellogg Company, Public Affairs Department, Battle Creek, MI 49016 Eli Lilly & Company, Public Relations Department, 307 East McCarty Street, Indianapolis, IN 46285 "Kidney in Action/The Fact of Life," "Children's Zoo/Biology at the Molecular Level" Oscar Mayer & Company, Consumer Affairs, P. 0. Box 7188, Madison, WI 53707 McDonald's Action Packs, Box 2594, Chicago, IL 60690 (Information for Action Packs) Missouri Department of Conservation, P. 0. Box 180, Jefferson City, MO 65132 Missouri Department of Natural Resources, Box 176, Jefferson City, MO 65101 "Environmental Education Resources" National Audubon Society, Education Division, 950 Third Avenue, New York, NY 10022 National Bureau of Standards, Metric Information, Washington, DC 20234 National Foundation/March of Dimes, Box 2000, White Plains, NY 10602 "Genetic Counseling"
289 National Peanut Council, 1000 16th Street NW, Suite 506, Washington, DC 20036 "Peanut Portfolio" National Wildlife Federation, 1412 - 16th Street NW, Washington, DC 20036 Environmental Discovery Units, Conservation Education Catalog Population Reference Bureau, Inc., 777 - 14th Street NW, Suite 800, Washington, DC 20005 Purina Cat Care Center, Checkerboard Square, St. Louis, MO 63188 "Handbook of Cat Care" Soil Conservancy Society of America, 7515 NE Ankeny Road, Ankeny, IA 50021 Sergeant's, P. 0. Box 25595, Richmond, VA 23260 U. S. Department of Agriculture, Box 385, Vandalia, OH 45377 "Great American Farm" U. S. Department of Agriculture, Forest Service, P. 0. Box 2417, Washington, DC 20013 U. S. Department of Agriculture, Publications, Office of Information, Washington, DC 20250 U. S. Department of Commerce, National Oceanic & Atmospheric Administration, Rockville, MD 20852 "Fish: Wet & Wild, Teachers Guide K-6" U. S. Department of Health and Human Services, Bureau of Health Education, HHS Publication No. (CDC) 80-8359 and 80-8382, Atlanta, GA 30333 "School Health Cirriculum Project" U. S. Government Printing Office, Superintendent of Documents, Washington, DC 20402 Marine Mammals of the Western Hemisphere (poster) #003-00106-8 "Soozie" #027-004-00024-5; Also, "Fun With The Environment (EPA)" University of Missouri Extension Service, University of Missouri, 1408 1-70 Drive SW, Columbia MO 65203 Wildlife Management Institute, 1000 Vermont Avenue, Suite 709 Wire Building, Washington, DC 20005 Xerox Education Publications, 1250 Fairwood Avenue, Columbus, OH 43206 Information for Science Unit Books "What Insect is That?" "The Body Machine - Parts & Functions" "Ecology: Man Explores Life"
RESOURCE MATERIALS: Bonus 25 points - must be biological in nature - five different cources - must have information card with: 1) topic; 2) name of publication; 3) source of publication; 4) cost, if any; 5) grade level; 6) classroom use. - Please use only one resource per agency, recently published material (last five years), and no commercial magazines, texts or old lesson plans.
290 LIBRARY SURVEY
The intention of this project is to make you aware of the variety and quantity of resource material that is available to you. It may also impress you with the amount and depth of content to be taught in the elementary classroom. Select five biological concepts. Locate two different resources which teach each of the concepts. These resources may be elementary science and health texts, OBIS activ- ities, or periodicals such as "Science and Children". Make a file card (5x7 preferably) for each of the ten resources. First, state the concept. (A scientific concept is a concise statement of a single principle, fact, or idea.) Then, make a complete bibliographic entry which includes author, title, publisher, date of publication, and page number. State the grade level and briefly summarize the activity. The activity should not exceed one hour of class time. To the best of your ability, evaluate the resource. Be brief. Consider the following questions: * Was it appropriate for the grade level? * Was the student actively involved? * Was the equipment easy to collect? available? simple to use? * Were the directions clear? * How well did the activity teach the concept? * Would you consider using this resource? Why or why not?
291 GENERAL BIOLOGY FOR ELEMENTARY EDUCATION "Mini" Investigations
REMOVAL OF SEED EMBRYOS MATERIALS: 12 bean seeds, 4 pots with soil, plastic wrap, paper toweling, petri dish, scalpel. DIRECTIONS: a) Place 12 bean seeds on wet paper towels in petri dish. Cover with plastic wrap. 1) When seed coats split and beans appear swollen (2-3 days), care- fully open a seed and look at the parts. Examine with a magnifying lens. Draw what you see. 2) Carefully dissect the plant embryo from 3 seeds. Carefully dissect the root embryo from 3 seeds. Carefully dissect the shoot embryo from 3 seeds. b) Plant 3 bean seeds in each pot. Provide a good growing environment for the seeds. Observe for one week. QUESTIONS: Identify the parts in the seed by using the text. Do all 4 sets show growth? Dig up the seeds. What part of the seed shows growth? Would seeds die if cotyledons are removed? Why or why not? What is necessary for a healthy seed?
A PICKLE TREE! MATERIALS: 1 cucumber 200 ml vinegar 15 ml salt 15 ml pickling spices 150 ml sugar 2 500-1111 jars DIRECTIONS: Wash the cucumber. Cut slices about 2 ml thick. Put ½ of them in each jar. Mix together the sugar, vinegar, salt, and pickling spices and put them in ONE jar with the cucumber. Mix it well. Put the lid on it. Put an equal volume of water into the second jar, mix well. Put the lid on it. Put both jars into the refrigerator for one week before tasting!
QUESTIONS: Describe the texture of the cucumbers in each jar. What physical changes occurred? What conditions are the same for both jars? What conditions are different? Is there a control for this investigation? What can be said about cell membranes in cucumbers (the permeabil ity). By what plant process does pickling take place? Why should uncanned pickles be stored in the refrigerator? Why will there never be a pickle tree?
292 THE EFFECTS OF GRAVITY ON PLANT GROWTH
MATERIALS: 10 bean seeds, 2 small wide mouth jars, paper toweling. DIRECTIONS: Soak the seeds overnight. Line the insides of both jars with a piece of wet paper towel, folded to fit. Fill the middle of each jar with crumpled towel, enough to hold the outside layer in place. Saturate the paper with water, pouring off the excess. Push 5 seeds between the glass and the towel (outside layer) about 3 cm from the top of each jar. Put the lid on each jar until the seeds sprout, then leave them open. Set ONE jar on its side and maintain this position throughout the experiment. Set the other jar upright and maintain that position throughout the experiment. Put both jars in moderate sunlight and BE SURE TO KEEP THE PAPER MOIST during the experiment. Check the jars each day to collect data. QUESTIONS: Is there a control for this experiment? If so, what? What is the variable for the experiment? If these jars were in a space capsule, describe the plant growth. Could the light source be a possible cause for this growth pattern? How could this be eliminated?
WHAT WILL THEY WEIGH?
MATERIALS: About 450 grams of green leaves with stems and big leaf-ribs removed. Cookie sheet and oven for drying leaves; large inexpensive piece of paper on which to chart the experiment for the class. Small note- size piece of paper for the class. Scales for weighing. DIRECTIONS: Collect and weigh the leaves. With a colored marker, chart the weight (in grams) on the bulletin board graph. Pass out note-size paper to each pupil. Ask the pupils to estimate the gram weight of the leaves when all the water has been removed. Collect the estimates and place them in a box or drawer. Explain to pupils that at the end of the experiment you will determine whose estimate came closest to the actual dry weight. Dry the leaves in a warm (66°C) oven. After two hours, weigh the leaves again. Chart the weight loss on the bulletin board. Announce the winner of the "Winning Weight Award". INFORMATION: Most plants require an amazing amount of water! People who study plant life say that for every pound of dry matter the plant produces, it requires 500 to 1,000 pounds of water. The plant does not retain all the water, because most of it is returned to the air as the plant respires. Just as animals return vapor to the air when they breathe, so do plants. Instead of a nose or mouth, plant leaves have tiny openings called "stomata" through which they respire.
293 AREA OF ROOT GROWTH
MATERIALS: Petri dish, paper towel, 3 bean and 3 corn seeds, permanent ink, thread, cm ruler. DIRECTIONS: Soak the seeds for 24 hours. Cut 2 layers of paper towel to fit the petri dish, wet it and put it into the dish. Place the soaked seeds on the paper, cover them with another layer of damp towel, replace the lid, and set the dish aside to germinate. Check the plate in 2 to 3 days to see that the seeds are still moist and if germination has occurred. When the roots of the seeds are about 2 cm, measure and mark them at 2 mm intervals. Use the cm ruler, and dip the thread into the ink to make the mark. Put the seeds back into the dish, cover them with a damp towel, and observe them for another four days. QUESTIONS: Where have the roots grown the most? How do you know this! How are the corn and beans alike in the way that they grow? What kind of plant tissue must be concentrated here?
HOW DOES WATER GET INTO A PLANT?
MATERIALS: Radish seeds, wet paper towel, petri dish, a magnifier. DIRECTIONS: Cut the paper towel to fit the dish. Put two layers of wet towel on the bottom of the dish. Arrange about 15 seeds on the wet towel. Cover the seeds with another damp paper towel. Place the cover on the plate and set aside at room temperature and light. Observe for several days, recording your observations. Use a dissecting microscope whenever possible, (or a hand lens). QUESTIONS: What do you notice about the roots? What are the small fuzzy projec- tions from each root called? What is the function of these structures? What happens to the roots of a plant when it is transplanted? What will happen if these structures are removed from the root? How far from the root tip do these structures extend?
EFFECT OF LIGHT ON GERMINATION OF SEEDS
MATERIALS: Bean seeds, two pots with garden soil. DIRECTIONS: Soak the seeds overnight. Plant three seeds in each pot of soil, add 125 ml water to each pot. Place one pot in lighted area. Place the other pot in a dark place away from light. Keep both pots at room temperature, water 50 ml every other day. QUESTIONS: What things are the same for each pot? What is different? Do you have a control? If so, what is it? If not, why not? What concept does this experiment illustrate? Why do we wait for spring to plant seeds? Could seeds germinate in a cave? What is germination?
294 DIRECTION OF ROOT GROWTH
MATERIALS: Corn seeds, damp paper towel, petri dish, cotton, masking tape. DIRECTIONS: Soak the seeds overnight. Place four corn seeds in a petri dish with each seed pointing toward the center. Cover the corn with a damp paper towel cut to fit; cover the towel with damp cotton until seeds are held securely in place. Tape the lid on dish and set dish on edge. Fix securely in position so that there is no rotation during the experiment. Observe the growth. QUESTIONS: What physical response is at work in this experiment? How do you account for any unexpected results in this experiment? If the dish was rotated on its edge, describe the position of the roots.
DO PLANTS GIVE OFF WATER?
MATERIALS: Two potted coleus plants, 2 quart jars, plastic bag or plastic wrap. DIRECTIONS: Add 125 ml water to the soil in the pots. Wrap the plastic around the entire pot, tying securely around the plant stem. Remove all leaves from one plant. Invert the glass jars over the plants. Place in the sun or under grow light in the lab. Observe. QUESTIONS: Why were the pots covered with plastic? Does the water get out of the plant? What is the evidence for this? What tissue is involved in water transport? If water leaves the plant, how does it? Why remove the leaves from one plant?
BEHOLD THE MOLD!
MATERIALS: Homemade wheat bread, commercially baked wheat bread, wet paper towel, 3 petri dishes. DIRECTIONS: Place 1 layer of wet paper towel in the bottom of each of the three petri dishes. Place a 5 cm square of homemade bread in 1 dish, and a 5 cm square of the commercially baked bread in the second dish. Leave the third dish with just the wet paper towel Put the lids on the dishes and set aside at room temperature and normal light for approximately three days. QUESTIONS: What things are the same for all three petri dishes? What is different? What is the purpose for the third dish? In which dish did mold grow? In which dish did the mold show first? the most? Why put wet paper towel in each dish? Where could the mold have come from? From this experiment, what inferences can be made about the effect of light on mold growth? the effect of temperature? the effect of moisture? the effect of time? the effect of the nutrients for mold growth?
295 SEEDS 'N SOIL
MATERIALS: 20 radish seeds, 4 containers each of gravel, sponge, sand and potting soil. DIRECTIONS: Soak the seeds overnight. Fill each container 2/3 full of potting material. Plant 5 seeds into each pot. Add 100 ml of water and set aside in a warm (not hot) sunny place. Water as needed, being sure that each one gets the same amount. QUESTIONS: In which pot did the seeds sprout first? second? etc. Did all of the seeds in each pot sprout? Did all the seeds grow at the same rate? What is the same for all 4 pots? What is different? Why plant five seeds instead of one? Are all of the seeds the same? How do you know? From this experiment, what can be inferred about the effect of water on radish growth? the temperature? the amount of light? about the potting material.
EFFECT OF HEAT ON GERMINATION OF SEEDS
MATERIALS: Bean seeds; 3 flower pots with soil. DIRECTIONS: Soak the seeds overnight. Plant 3 seeds in each of the pots of soil, add 125 ml water. Place one pot in a refrigerator (record the temperature). Place second pot in warm or hot spot (record the temperature). Keep the third pot at moderate room temperature (not in sun). Water each of the pots daily (the same amount of water for each one). QUESTIONS: What is the known variable? constants? control? What concept does this experiment illustrate? why soak the seeds overnight? Why plant more than one seed in each pot?
296 297 PLANT PHYSIOLOGY
There are many chemical reactions necessary to maintain life within the plant cell. The term metabolism has been used to describe the sum total of all these diverse chemical changes. As in other living cells, some chemical reactions are constructive and are termed anabolic. Some of these are photocynthesis and assimilation of nutrients for growth and repair. Destructive processes essential to survival are respiration and digestion. These are called catabolic reactions. Respiration Plant cells use energy to aid in building and maintaining protoplasmic structures and cell walls. Within each cell, energy is obtained by a slow and very controlled process of "burning". This oxidation reaction is called respiration. It combines oxygen with glucose and produces carbon dioxide, water, molecules of chemical energy (ATP) and heat energy. The process is represented by the following equation:
+ + + energy (ATP & heat)
Animal cells have the same basic cellular respiration. Their method of obtaining oxygen may often be different. The lower animals get oxygen by simple diffusion of gases from their environment across a moist membrane and into their system. Higher animals have a mechanical means of obtaining oxygen; that is, by breathing. Lenticels in stems and stomata in the leaves are openings which allow for the exchange of gas in plants. There are two guard cells which border and regulate the opening and closing of each stoma. When the crescent shaped cells are turgid, the stoma is open. When the cells are flaccid, the opening is closed. Heat energy produced by cellular respiration is considered to be wasted energy. How- ever, it can be very important commercially. If the heat is confined in an enclosed area, it can: a) aid growth of bacteria and fungus, b) raise temperature to above the death point and kill the seeds or plants, or c) cause spontaneous combustion, i.e., fires in hay barn! Two demonstrations illustrate evidence of respiration. The simple rise in tempera- ture within the vacuum containing living seeds indicates active respiration. What is the purpose of the other two containers? What will be the limiting factor in the thermos of living seeds. Another product of cellular respiration is carbon dioxide. Carbon dioxide in phenol red solution forms carbonic acid which changes the pH and turns the solution color from red to yellow. When the air from a closed bottle of germinating seeds is forced through the phenol red, a color change occurs. Air from a closed bottle of "killed" germinating seeds forced through phenol red solution remains red in color indicates no carbon dioxide present. How can you demonstrate the reaction of pehnol red to carbon dioxide? Photosynthesis Green plants have the ability to manufacture food from raw materials found in their environment. This process is called photosynthesis. It is the manufacture of sugar from carbon dioxide and water in the presence of chlorophyll and sunlight. A sobering thought is that the life of plants, animals and all of mankind depends on this activity. It is the most important chemical process known to man. The equation for this reaction is: chlorophyll 2 = radi ant energy 6CO + 6H2O C6H12O6 + 6O2
Photosynthesis proceeds in two series of reactions. In the "light" reaction, specific wavelengths of light energy are captured by chlorophyll and stored in chemical bonds of ATP and NADPH. Second, in the "dark" reaction, the energy in ATP and the reducing agent NADPH, is used to convert the carbon dioxide and water into organic molecules. In turn, those sugar molecules can be converted into cellulose molecules, other kinds of building materials, or hooked together and stored as starch.
298 A simple demonstration using a healthy variegated Coleus leaf can illustrate photo- synthesis activity by locating areas containing chlorophyll and where starch has been formed.
EVIDENCE OF PHOTOSYNTHESIS IN A COLEUS LEAF
1. Remove the leaf from the plant. Remove the black circle of paper from the leaf. Draw a picture of the leaf, noting the various areas of color. Turn the hot plate to medium heat. 2. Put the leaf into a 250 ml beaker containing about 100 ml water. Let it boil 4 or 5 minutes. Using tongs or forceps, remove the leaf and lay it flat in a petri dish. Draw a picture of the leaf. Why is the leaf limp? Have the areas of color changed? Why? What colors are still in the leaf? 3. Put the leaf into a 250 ml beaker containing about 75 ml of alcohol. Boil for 3 or 4 minutes. Using forceps, remove the leaf and lay it flat in a petri dish. Turn OFF the hot plate. Draw a picture of the leaf. What color is present now? Is chlorophyll water soluble? Can chlorophyll be in plant cells and not be visible? 4. Iodine has a brown color. When it is put on a starch, it turns a dark blue-black color. Therefore, it can be used as a reagent to detect the presence of starch. Using iodine from the dropper bottle, cover the leaf with iodine. Let it stand a couple of minutes, then examine the leaf for color again. Draw the leaf. Where is it darkest? What does that indicate? Was chlorophyll located uniformly in the leaf? What purpose was served by the black circle covering an area of the leaf? Does this experiment have a control? Be sure to clean up the area after the experiment!
299 Respiration
Diagram showing gas exchanges due to photosynthesis and respiration in a green leaf in the light. Carbon dioxide is used five to ten times as fast as it is produced.
Photosynthesis and Respiration Compared. The facts on photosynthesis and respir- ation can be presented in tabular form to contrast these two processes:
Photosynthesis Respiration 1. Occurs only in the green cells 1. Occurs in every active, living cell of plants of both plants and animals 2. Takes place only in the presence 2. Takes place during the life of the of light cell both in the light and the dark 3. Uses water and carbon dioxide 3. Uses food and oxygen 4. Releases oxygen 4. Releases water and carbon dioxide 5. Solar (radiant) energy is con- 5. Chemical energy is converted into verted into chemical energy heat and useful (ATP) energy 6. Results in an increase in weight 6. Results in a decrease in weight 7. Food is produced 7. Food is broken down
300 GENERAL BIOLOGY FOR ELEMENTARY EDUCATION
Review: Handouts from previous lab Slide survey of Animalia Kingdoms. Game*: "What Animal Am I?" Pin a picture or name of an animal onto the back of one of the persons in the group. Don't show him the picture. He then asks questions to discover his own identity. The other persons can answer only yes, no or maybe. Because we are studying properties of animals, when asking for information, use the terminology appropriate for classification. Example: Do I have radial symmetry? yes Do I have an external skeleton? no Am I warm blooded? no Am I an Echinoderm? yes (a starfish) Topic: Animal Adaptations Adaptation - any change or modification of the organism which aids in its survival and reproduction. Adaptations may enable the animal to survive short periods of stress such as hiber- nating in the winter and estivating in the summer. Or, anatomical and physiological modifications may result in the animal being better suited for survival on a daily basis. The owl with the keenest vision will catch the most mice and in so doing better his chances to live, reproduce and add his genes for keen vision to the next generation. Because the environment is continually changing, adaptation is a dynamic process. Those organisms which survive and reproduce add the most beneficial hereditary material to the gene pool, i.e., "Survival of the fittest." (or, survival of the best adapted.) Today we will study some adaptations and design a few of our own. Demonstration materials: How are these special? (Bird heads and feet from Ornithology lab) ** Activity: Using materials provided, create predator devices that can catch and pick up prey. What do you think? 1. Why are there so many different kinds of predator devices? 2. What would happen if every animal had the same predator device? 3. What adaptations do prey have to avoid being eaten by predators? 4. What would happen if the "unfit" were nurtured to survive and reproduce?
* Cornell, Joseph. Sharing Nature With Children ** Adaptation-Predator-Prey, OBIS I 301 GROCERY BAG BOTANY
Grocery Bag Botany is exerpt from the plant reproduction lab which follows a brief study of plant morphology. It is a fun way to conclude the botany section of labs. It can also be challenging. When used in second and third grade science, we first discuss whether the produce is fruit or vegetable in common use, then determine whether the items are fruit or vegetable in the botanical sense. We review the organs of plants and their primary functions. The students then determine which plant organ is represented by each item. (Keep everything inside a large paper grocery bag and present one item at a time.) Select a mixture of common and exotic items (more of the common for the elementary grades and more of the exotic for the college students): leaf lettuce and spinach (leaves); artichoke, onions, garlic (fleshy leaves); brussel sprouts, cabbage (leaf buds); beets, carrots, sweet potatoes (roots); celery, rhubarb (petioles); cauliflower, broccoli (flowers); strawberry, tomato, cucumber (seeds and ovary); corn on cob, shelled peas, walnuts, coconut (seeds); irish potato, asparagus (stems); green pepper, apple, orange, squash (ovary). Peanuts in the hull are interesting be- cause of their underground development. Be sure to include a mushroom--it creates discussion (and an egg for laughs). Most college students seem to enjoy this exercise as an informal quiz. However, questions may be more specific. This activity may be as involved or as brief as you want to make it.
302 Brown Paper People
This short activity is used to introduce the series of labs review- ing human systems. The first system of study is the skeletal. The only materials required initially are 6'x3' lengths of brown paper, pencils with erasers, and felt marking pens or crayons. A large picture of a skeleton or actual skeleton is needed to complete the activity. The disarticulated skeletons are optional. At SMSU these are borrowed from Anatomy lab. Working in groups of two or three, one student must lie down on the paper (dorsal down), in anatomical position. Using marking pens, the other student(s) carefully draw(s) his outline. The prone student gets up and the two or three of them procede to sketch in the skeleton as they perceive it to be. Their only resource being recall and palpating their own skeleton where possible. Be sure to have them sign their drawings. (Allow about 15 minutes. ) As the students are finishing their drawings, set out two or three boxes of disarticulated skeletons. Select two or three drawings and have the students (now in larger groups) attempt to assemble the bones in correct fashion within the human outline. (Allow about 10 minutes. ) Uncover the articulated skeleton and as a class discuss which bone goes where and why. Each group shows their work or if room permits I hang them on the walls. After lab, the paper people are rolled up and saved for another system. Cutouts of organs of the digestive, respiratory, repro- ductive and urinary system can be placed using glue sticks. On the reverse side another outline is made and the circulatory system drawn in using red and blue crayons or markers. While this is a very elementary activity, it serves several purposes. There is active student involvement first as small groups, then larger groups, and ultimately as a class effort. Lying on the floor in one's stocking feet while your classmates draw around you tends to soften defensive/"stand offish" attitudes. Pooling efforts of recall and creativity in making that brown paper person assume a personality make a shared experience. Finally, displaying all the originality of forms makes the students aware of their need to learn about the human system(s).
303
Chapter 14
Laboratory Safety Principles
A. Radioactive Materials-Jerry Staiger
B. Toxic, Reactive, Carcinogenic, and Teratogenic Chemicals-Keith Carlson
C. Infectious Agents-Jim Lauer
D. Fire and Physical Hazards-Ray Arntson
Note: This workshop consisted of four separate subtitles and presentations.
305
Radioactive Materials
Jerome W. Staiger
Radiation Protection Officer Department of Environmental Health and Safety 410 Church St. S. E. University of Minnesota Minneapolis, MN 55455
Jerome (Jerry) W. Staiger received a B.S. degree from the University of Wisconsin- River Falls (physics) and an M.S. degree from Rutgers University, New Brunswick, New Jersey (radiological health). He has worked as a professional health physicist in the Department of Environmental Health and Safety, University of Minnesota since 1966. For the past three years he has been Radiation Protection Officer for the University of Minnesota. As director of the Radiation Protection Program for the University of Minnesota, he has responsibility for radiation safety review and inspection of a wide scope of medical and research uses of ionizing radiation.
307
PROCEDURES FOR RADIOISOTOPE USE AREAS
All users of radioisotopes must comply with the following procedures and ensure that persons under their direction adhere to these rules.
Identification of Use Areas
Each door to an area where radioisotopes are used or stored must be posted with a "CAUTION RADIOACTIVE MATERIAL" sign, and such areas must be maintained as controlled areas, with doors closed and locked when the area is unattended. For further information on controlled areas, see Appendix F. The name and phone number of the approved user must be on the caution sign, along with the phone number of the Radiation Protection Program. The signs must not be removed from any room except by a Radiation Protection Program staff member following a radiation protection closeout survey.
Each radioisotope storage area must be conspicuously posted with a "CAUTION RADIOACTIVE MATERIAL" sign. Any vessel which contains stock radioactive material must also be labeled with a "CAUTION RADIOACTIVE MATERIAL" label such as radiation caution tape. These signs and labels must state the kinds of radioisotopes and the quantity being stored.
Any area where persons could be exposed to 5 mrem/hour to the whole body or where the exposure could exceed 100 mrem in five consecutive days must be posted with a "CAUTION RADIATION AREA" sign.
Contact the Radiation Protection Program for information on the availability of caution signs for various purposes.
Shielding of Radioactive Sources
Appropriate shielding must be provided so that the radiation exposure rate from radioisotope sources is less than 2.5 mR/hour in any controlled area and less than 0.25 mR/hour in any uncontrolled area. Radiation exposure rates should be maintained as low as is reasonably achievable within these limits. Contact the Radiation Protection Program regarding shielding materials and techniques.
309 Radioactive Gases, Dusts and Aerosols
All procedures involving radioactive gases, dusts or aerosols or any experiment that might release airborne radioactive contamination must be conducted in a well ventilated hood or other enclosure that has been approved by a Radiation Protection Program staff member. Specifications for such enclosures may be obtained from the Radiation Protection Program.
Radioactive gases must be stored in gas-tight containers that are kept in approved ventilated areas.
Sealed Radiation Sources
Sealed radiation sources must be handled with tongs or other remote handling devices. (This procedure is not required when handling very low activity check sources.) The dose rate at the hand and body location should be determined before such sources are used or moved. As required under Nuclear Regulatory Commission license conditions, alpha sources will be leak tested by a Radiation Protection Program staff member every three months, and beta and/or gamma sources every six months. Damaged or lost sources must be reported to the Radiation Protection Program immediately.
Periodic Surveys of Radioisotope Facilities
Each laboratory using radioactive materials (except labora- tories using only very low activity sealed sources) must have available a radiation monitoring instrument capable of detecting low levels of contamination. Smear surveys of laboratory use, storage and waste disposal areas must be performed on a routine basis. A minimum frequency of once per month when radioisotopes are in use and once per quarter when the laboratory is used only for storage is required. Permanent records of the results of these surveys must be maintained. For quarterly report requirements, see Appendix D.
The preferred method for evaluating removable radioactive contamination is to wipe the surface to be evaluated with a filter paper such as Whatman 1 (4.25 cm). A surface of approximately 100 square centimeters should be wiped and the paper counted in an appropriately calibrated liquid scintillation counter or other counting instrument capable of detecting the type of radionuclide being used. Liquid scintillation counting is the preferred method for detecting contamination from low energy beta emitting radioisotopes 14 3 such as C and H. It is also a good method for evaluating possible contamination from all other beta emitting radioisotopes.
310 According to Nuclear Regulatory Commission license require- ments, all smear survey results obtained by individual users must be reported in disintegrations per minute (DPM) per 100 square centimeters, rather than in counts per minute (CPM).
In order to obtain these results, it will be necessary for the user to indicate on the survey form the gross CPM per 100 square centimeters, the background CPM, the fractional efficiency of the counting instrument for each radioisotope analyzed, and the particular counting system used, i.e., liquid scintillation, auto-gamma, etc. The fractional efficiency (E) can best be determined by counting a known activity of the radioisotope and dividing the count rate (CPM) by the disintegration rate of the known activity (DPM), i.e., E = CPM/DPM. The disintegration rate (DPM) is obtained by multiplying the activity in microcuries (µCi) by 2.22 x 106 DPM/µCi.
A Geiger-Mueller (G-M) portable radiation survey instrument is often used to detect contamination; however, this instrument is not effective in detecting low energy beta or alpha emitters. Contact the Radiation Protection Program for assistance in determining the appropriate method for evaluating possible contamination in individual laboratories before purchasing radiation survey instruments.
The Radiation Protection Program provides a service of periodically performing radiation protection surveys of all radioisotope laboratories. As part of this survey, a Radiation Protection Program representative will review the records of surveys made by the approved user. A report of these surveys will be sent to the approved user. These surveys do not substitute for the routine surveys performed by the approved user.
Laboratory Equipment Maintenance and Disposal
Radioisotope laboratory equipment intended for disposal or requiring maintenance must not be removed from the radioisotope laboratory until surveyed by the approved user and demonstrated to be free of contamination. If repairs must be made of contaminated equipment, the user must contact a representative of the Radiation Protection Program who will directly supervise the work. No contaminated glassware may be taken to the glass shop for repair or modification unless approved by a Radiation Protection Program staff member.
311 Use of Radioisotopes in Animals
Before a permit is granted for use of radioisotopes in animals, a Radiation Protection Program representative will review the animal care procedures with the applicant. The applicant must have adequate animal care facilities and must make provisions for collection and storage of animal carcasses and associated waste.
Animals that present a potential for airborne radioactive contamination must be housed in a properly ventilated radio- isotope hood or room.
Animal carcasses and associated wastes must be packaged in accordance with the procedures outlined on pages XI-7,8.
Cages used for housing animals treated with radioisotopes must be labeled with a radiation caution tag that lists the type and quantity of radioisotopes in each animal and the date of administration.
Absorbent paper used to collect animal urine and feces must be changed frequently to reduce the potential for airborne release of radioisotopes. The contaminated absorbent paper must be disposed of in a radioactive waste container located in the animal care area.
Routine contamination smear surveys must be made of the cages and room where such animals are housed, and areas that indicate removable contamination must be immediately decontaminated.
Cages that are to be cleaned in a cage wash facility must first be surveyed for contamination, and contaminated areas must be cleaned to less than 2200 DPM per 100 square centimeters. A tag which instructs the cage washer operator not to recycle the wash water must be attached to the cages.
If animals containing radioisotopes are to be cared for in general animal care facilities, the applicant must provide training to personnel who will care for the animals following the procedures outlined in Appendix O. If any assistance is needed in the training of animal care personnel or the monitoring of use facilities, contact the Radiation Protection Program.
For further information on the use of radioisotopes in animals, see Appendix O.
312 Laboratory Design and Plan Review
All plans for radioisotopes laboratory modification and new laboratory construction must be reviewed and approved by a Radiation Protection Program representative. Effective and safe use of radioisotopes requires special design of facilities and equipment. The laboratory should be designed to reduce the possibility of contamination and to be easily decontaminated if accidental contamination should occur.
General Laboratory Considerations
Good housekeeping should be observed at all times to control contamination and to facilitate cleaning in laboratories where radioisotopes are used.
Eating, drinking, smoking, and cosmetic application are prohibited in the radioisotope laboratory. Never store food of any kind in a radioisotope laboratory or in any refrigerator or freezer in a radioisotope laboratory. University policy on this regulation is given in Appendix T.
Mouth pipetting is prohibited in all radioisotope laboratory areas. Use syringes or other remote pipetters. Contact the Radiation Protection Program for information on such equipment. See Appendix T for the policy statement on this regulation.
Wear laboratory coats, disposable gloves, and when necessary, eye protection when working with radioactive materials. Wash hands thoroughly after removal of gloves.
To prevent the spread of contamination, use radioisotope trays lined with absorbent paper when working with radio- isotopes. Contact the Radiation Protection Program regarding the availability of these trays.
If there is any possibility of an airborne hazard, all work with radioisotopes must be carried out in an adequately ventilated hood.
For further information, see Appendix F, "Guidelines for Safe Use of Radioisotopes".
Radioisotope Shipments
Radioisotope shipments must be opened in a hood if there is any possibility of airborne contamination.
313 When opening a shipment, wear disposable gloves and examine the contents to determine possible damage or leakage. If damage or leakage is noticed, contact the Radiation Protection Program immediately.
A smear survey of the shipping box must be conducted. If the smear survey indicates contamination, dispose of the shipping box in the solid radioactive waste container. If a smear survey indicates no contamination, the box may be discarded as non-radioactive waste provided all radiation labels and markings have been removed from the box.
Monitoring
Radiation survey instruments shall be used to monitor clothing, hands, shoes and the laboratory area. A G-M survey instrument can be used to make contamination surveys for most types of radioisotopes; however, for 3H, 14C, or other low energy beta emitting radioisotopes, it is necessary to make a smear survey of laboratory surfaces, including the floor, using a piece of filter paper such as Whatman 1 (4.25 cm diameter). This paper is then placed in a counting vial and counted in a liquid scintillation counter. The smear survey is the preferred method for evaluating contamination for all beta emitting radioisotopes. It is important that frequent surveys be made to reduce the possibile spread of radioactive contamination. See Appendix D for minimum frequency requirements. Records of these surveys must be maintained by the approved user. Contact the Radiation Protection Program for more details on instrumentation.
Occupationally exposed pregnant women should maintain radiation exposure to within 500 mrem during the nine month gestation period according to Nuclear Regulatory Commission regulation guide 8.13. For further information, see Appendix S.
Personnel monitoring devices must be worn when the radi- ation exposure potential warrants their use. A Radiation Protection Program representative will evaluate the need for film badges in areas where potential external radiation exposure exists.
Storage
Provisions must be made for the proper storage of radioisotopes in the laboratory. Sufficient shielding must be provided to maintain radiation exposure levels within permissible limits. Radioisotopes must be stored in containers that prevent spillage and contamination. See Appendix G for permissible dose limits.
314 Transportation
When radioisotopes are transported through an uncontrolled area, they must be packaged in a leak-proof, nonbreakable container to prevent release of the radioactive material. When radioactive liquids are in a glass container, the container must be kept in a second, non-breakable vessel that will hold the radioactive contents in the event that the glass container is broken.
Wastes
Radioactive wastes must be disposed of only in approved, approp- riately marked containers. Contact the Radiation Protection Program to request disposal service. Do not mix acids and organic solvents as waste. Records must be kept of the radioisotopes and activities in each waste container. These records will be reviewed by Radiation Protection Program staff members. For more detailed information on radioactive waste, see section XI.
Information on Specific Radioisotopes
General laboratory procedures for radioisotope use have been given above. More specific information on radioisotopes commonly used at the University can be found in the following appendices:
Appendix U: Some Commonly Used Radioisotopes and Their Relative Radiotoxicities.
Appendix J: Information and Procedures for Use of 3H and 14C.
Appendix K: Handling and Survey Procedures for High Energy Beta Emitting Radioisotopes.
125 131 Appendix L: Precautions in the Use of I and I.
315
APPENDIX F
GUIDELINES FOR THE SAFE USE OF RADIOISOTOPES
1. Observe good housekeeping at all times to prevent contamination and to facilitate cleaning.
2. When pipetting radioactive materials, use syringes, remote pipetters, rubber bulbs, or other mechanical devices. DO NOT PIPETTE BY MOUTH.
3. DO NOT EAT, DRINK OR SMOKE IN THE LABORATORY. Do not store food of any kind in the laboratory or in a refrigerator or freezer where radioactive materials are stored.
4. Wear laboratory coats and disposable gloves when working with radioactive mate- rials. Wash hands thoroughly after removing gloves.
5. When working with radioisotopes, use stainless steel or fiberglass trays lined with absorbent paper. Contact the Radiation Protection Program for information about these trays.
6. Wear film badges when the radiation hazard warrants such use. The Radiation Protection Program will evaluate the need for film badges in areas where potential external radiation hazards exist.
7. If there is any possibility of an airborne hazard, do all work in a hood.
8. Transport and store all radioisotopes in a manner that will prevent breakage and spills. When a radioactive liquid is in a glass container, keep the container in a second unbreakable container that would hold the radioactive contents if the glass container were to break.
9. Provide for proper storage of radioisotopes in the laboratory, with sufficient shielding to maintain a safe radiation level.
10. Post proper radiation caution signs on the doors of laboratories in which radio- isotopes are used and indicate use and storage areas within the laboratory. Contact the Radiation Protection Program for caution signs.
11. Use a radiation survey instrument to monitor clothing, hands, shoes and the general laboratory area. For 3H and 14C, smears should be taken on laboratory surfaces, including the floor, using a piece of absorbent filter paper. This paper may then be placed in a counting vial and counted in a liquid scintillation counter. It is important that weekly radiation surveys be made to reduce the hazard from radioactive contamination. The laboratory director must keep records of these surveys. Contact the Radiation Protection Program for more details on instrumentation.
12. When opening a shipment, examine the contents to determine if there has been any damage or leaking. Report damaged or leaking shipments to the Radiation Protection Program immediately.
13. If there is any possibility of airborne radiation hazard, open radioisotope shipments in a hood. This is especially important with shipments of particulate matter or materials with high radiotoxicity.
317 14. If a smear survey indicates contamination, dispose of shipping boxes in a solid radioactive waste container. If a smear survey indicates no contamination, the box may be discarded as non-radioactive waste, provided ALL radiation labels and markings have been removed.
15. Handle sealed radiation sources, regardless of activity, with tongs. Determine the dose rate at the hand and body location before sealed radiation sources are used or moved. Alpha sources must be leak tested by the Radiation Protection Program every 3 months, beta and gamma sources every 6 months. Report damaged or lost sources immediately to the Radiation Protection Program.
16. Do not remove contaminated tools, glassware, or equipment from the laboratory. Store away from non-contaminated equipment. Glassware used with radioisotopes may not be taken to the glassblower without prior approval by the Radiation Protection Program.
17. Dispose of radioactive wastes in appropriately marked approved radioactive waste containers. Do not mix acids and organic solvents as waste. Contact the Radiation Protection Program for waste disposal service.
For further information on any aspect of .radiation safety, contact the Radiation Protection Program, W140 Boynton Health Service, Department of Environmental Health and Safety, 626-6002.
318 APPENDIX G
EXPOSURE TO INDIVIDUALS TO RADIATION IN RESTRICTED AREAS MAXIMUM PERMISSIBLE DOSE PER CALENDAR QUARTER
Rems/Qtr Rems/Yr
1 . Whole body; head and trunk; active blood-forming organs; lens of eye; or gonads ...... 1.25 5 2 . Hands and forearms; feet and ankles ...... 18.75 75 3 . Skin of whole body ...... 7.5 30 4 . Other organs ...... 3.75 15 5 . Thyroid ...... 7.5 30 6 . Individual in the general public ...... 0.5 7 . Student ...... 0.1 8 . Occupationally exposed pregnant women ...... 0.5 rem/ 9 mo . gestation
319
APPENDIX J
INFORMATION AND PROCEDURES FOR USE OF 3H AND 14C
3 14 3 14 1. Both H and C emit low energy beta radiation ( H - 0.018 MeV, C - 0.156 MeV), and therefore present no appreciable external radiation exposure hazard. Personnel working with 3H or 14C do not require a film badge monitor.
2. Shipments of 3H or 14C should be opened in a hood and inspected for damage and 14 contamination before transfer to a storage area. Most C shipments can be stored safely in a laboratory freezer or refrigerator. All 3H stock vials and labeled samples should be stored in a glass containment vessel with a metal foil seal. Tritium readily diffuses through plastic containers due to the small molecular size. Proper storage in a glass container will minimize diffusion of the tritium and minimize contamination to surrounding areas.
3 14 3 14 If the material labeled with H or C is volatile, such as NaB H4, 3H2O, or CO2, it should be stored and used in a well-ventilated hood. Before such volatile compounds are used, a "dry-run" of the experiment must be performed under the supervision of a member of the Radiation Protection Program. For further infor- mation regarding these procedures, contact the Radiation Protection Program at 626-6002.
3. Because of their low beta energies, 3H and 14C cannot be readily detected with a portable G-M survey instrument. Therefore, in order to evaluate possible contami- nation of the laboratory, it is necessary to perform contamination smear surveys of radioisotope use areas at least once per month using dry filter paper smears such as Whatman No. 1 filter paper, 4.25 cm diameter. The smears should be counted in a liquid scintillation counting system and the results of each survey recorded on the quarterly report form. Areas showing removable contamination greater than 250 disintegrations per minute (DPM) above background per 100 square centimeters should be decontaminated and resurveyed. (100 square centimeters is approximately equivalent to a smear swipe 24 inches in length.)
3 3 14 4. Individuals who receive H2O, NB H4, or CO2 in quantities of 100 millicuries or more, or those who receive tritium-labeled nucleotides in quantities of 10 millicuries or more, must have a urine analysis performed within one week of experimental use. Shipments of tritiated compounds of these quantities or greater will be stored by the Radiation Protection Program in the Boynton Health Service building until the indivi- dual user is ready to use the tritium and requests its delivery. A urine collection container will be provided with the shipment. The individual who uses the tritiated material must collect a urine sample approximately 24 hours after the use. The urine sample should then be delivered to W131 Boynton Health Service for analysis by the Radiation Protection Program.
5. If any spills of radioactive materials or any other emergency conditions arise, notify the Radiation Protection Program immediately at 626-6002.
321
APPENDIX K
HANDLING AND SURVEY PROCEDURES FOR HIGH ENERGY BETA EMITTING RADIOISOTOPES
I. Materials Required for Stock Solution Transfer
- Tray lined with absorbent toweling. - Cylindrical lucite stock vial shield. - Lucite barrier (body shield) to provide eye and body protection. - Remote handling devices (e.g., tongs, clamps, forceps, hemostats). - Gloves, surgical and disposable types. - Lab coat. - Film badge and TLD ring (wear ring under protective surgical glove with TLD ring label toward palm of hand). - Portable G-M survey instrument, preferably with pancake style probe for periodic check of hands, handling materials and transport containers to assure contamination control.
II. Protocol for Transfer and Handling of High Energy Beta Emitting Radioisotopes
1. Survey stock solution transfer station (i.e., hood or bench top) prior to bringing stock material to this area. This will assure that the area is free of contami- nation and will prevent possible cross contamination of individual experiments.
2. Double gloves must be worn during all radioisotope handling procedures. The outer glove should be monitored frequently and, if found to be contaminated, the glove should be removed and properly disposed of.
3. CAUTION - Because of the extremely high external exposure rate, the stock vial solution should never be held in the hands. Always use remote handling devices.
Exposure rates from 1 mCi of 32P over 1 square centimeter of skin: 2000 rads/hr at surface. 200 rads/hr at 1 centimeter. 22 rads/hr at 10 centimeters.
In 1 ml of water the surface dose rate for 1 mCi of 32P is 780 rads/hr or 13 rads/min. Because of these very high exposure rates, the handling of uncovered vessels (open, unshielded top) presents a serious potential for excessive and unnecessary radiation dose to the hands and face. Never place hands or any other part of the body over an open, unshielded vessel containing large activities of 32P in relatively small volumes of liquid.
4. All stock transfer procedures should be conducted in a hood if possible. Open the shipping container in the hood, using a remote handling device to transfer the stock vial to a lucite stock shield. Monitor the empty box with a G-M survey instrument. Remove any radioactive labels and markings before disposal.
323 5. A likely source of contamination results from opening the stock vial. Caution must be used in removing the vial cover and/or septum with the remote handling device. The tool used to remove the cover must then be treated as contaminated. A small beaker with absorbent gauze in the bottom should be used to isolate the vial cover, as well as the handling device.
6. Care must be taken when dispensing stock solution from the stock vial. Pipette tips or other transfer devices will be highly contaminated and are a major source of contamination of the work station and the individual's hands. A disposable waste receptacle (preferably a one-gallon plastic jug, Chemical Storehouse stock #CX12760), should be conveniently located behind the shielded work station for disposal of these and other contaminated items.
7. Cover each of the sample tubes with a cap or other suitable seal. Hands should be monitored following this procedure to determine if the gloves have become contaminated.
8. Seal the stock solution vial using the handling device initially used to open the vial. Dispose of outer gloves and monitor the surgical gloves prior to putting on a new pair of disposable gloves.
9. Transfer the stock solution in the lucite shield to your permanent storage lo- cation.
10. If the sample tubes are to be transferred to another work station prior to con- ducting the experimental procedure, this area must be equipped and shielded in the same manner as the stock transfer station (as indicated above.) A properly designed shield (lucite block with sample holes) must be used to carry the samples from the stock station area to the work station to prevent unnecessary radiation exposure and accidental spillage. If other laboratory personnel use the areas near the radioisotope work station, lucite side and back shields may be necessary.
11. During the experimental procedures, all items which contact the radioisotope solutions will be contaminated and must be properly handled and/or disposed of as radioactive waste. Hands and handling devices must be monitored frequently to check for possible contamination. Do not leave the work station before removing the outer pair of disposable gloves and monitoring the surgical gloves.
12. If the sample tubes are to be centrifuged, these must be placed in a larger containment vessel which has absorbent gauze placed in the bottom prior to centrifugation. This practice will reduce the possibility of contamination spread in the event that the sample tube is broken.
13. Upon the completion of the experiment, all radioactive waste must be transferred to the proper waste receptacle. The work area must be thoroughly surveyed (both G-M and smear survey) and decontaminated.
If additional information or assistance is needed in determining appropriate precautions in the use of high energy beta emitters, contact the Radiation Protection Program at 626-6002.
324 APPENDIX L
PRECAUTIONS IN THE USE OF 125I / 131I
Radioiodine, when used in volatile forms, presents the potential for exposure of the thyroid gland to levels of ionizing radiation in excess of permissible limits. To minimize personnel exposure, it is important that the following procedures be strictly adhered to.
BEFORE THE LABELING
1. Persons who plan to use radioiodine in labeling procedures must contact the Radiation Protection Program (626-6002) to obtain authorization for use.
2. All persons associated with the labeling must view the radiation protection training tapes and complete a test questionnaire which will be provided with the tapes.
3. Persons authorized to handle radioiodine must review and be familiar with the precautions and procedures recommended by the Radiation Protection Program.
4. Appropriate personnel monitoring devices must be obtained. These devices are available from the Radiation Protection Program. If 1.0 mCi or more of 125I or 131I is used, a TLD ring must be worn in addition to a film badge. The ring must be worn on the index finger under both pairs of gloves, with the ring label turned toward the palm of the hand.
5. A baseline thyroid scan is required. To schedule a scan contact the Radiation Protection Program at 626-6002.
6. A "dry-run" procedure (no radioactive material used) must be conducted. A member of the Radiation Protection Program staff will be present to make suggestions concerning appropriate precautions and procedures. Call 626-6002 to make arrangements for a dry-run.
7. Upon completion of these requirements, call 626-6002 to schedule a date and time for using the labeling facilities.
GENERAL CONSIDERATIONS
1. Shipments of sodium iodide and other volatile iodinated compounds must be addressed for delivery to 118 Boynton Health Service, 410 Church Street SE, Minneapolis, MN, ATTN: approved user's name. Such materials will be held by the Radiation Protection Program until the scheduled radioiodine labeling.
2. All uses of volatile radioiodine materials, in radioiodine labeling procedures or other operations where volatile radioiodine may be released, are restricted to facilities approved by the Radiation Protection Program. The facility on the Minneapolis campus is the Bond Laboratory, Boynton Health Service. The approved facility on the St. Paul campus is located in Room 338 Animal Science/ Veterinary Medicine building.
325 Breathing zone and environmental air samples will be taken during the procedure. Copies of air sampling reports, with any relevant comments, will be sent to the approved user under whose permit the radioiodination was done. To assure that personnel radiation exposure and environmental radioiodine releases are maintained within permissible limits, the Radiation Protection Program has developed investigation action levels which correspond to 10% of the MPC-hr. thyroid uptake, and 20% or the quarterly environmental release limit. If any individual exceeds these levels, it will be necessary for an investigation to be made to determine what caused the limit to be exceeded, and to designate corrective action necessary to reduce future levels. Persons exceeding the limits will not be allowed to conduct additional procedures using volatile radioiodine until corrective action has been completed.
4. All persons present during radioiodine labeling or during the handling of other volatile radioiodine must have a thyroid scan within one week of the procedure.
PROCEDURES FOR HANDLING RADIOIODINE
1. Absolutely NO mouth pipetting, smoking, eating, drinking, food or beverage storage or cosmetic application is allowed in radioisotope laboratories.
2. Sheet lead (1/16 inch) which provides adequate shielding for 125I should be used to shield labeling columns, stock vials, collection containers, etc.
3. To prevent contamination of the hood, work surfaces must be completely covered with absorbent pads (Hospital Supplies, inventory #CX41710).
4. A laboratory coat and long protective disposable gloves must be worn to protect skin surfaces from volatile radioiodine releases within the hood. The laboratory coat and disposable gloves must be surveyed frequently, and if contaminated, must be removed and disposed or decontaminated.
5. Disposable gloves must be worn when handling radioiodine, and double gloving is required. It is recommended that the bottom glove be latex (Chemical Storehouse, inventory nos. CX40932 - CX40938, according to size).
6. To reduce the possibility of contaminating other areas of the laboratory, the outer gloves should be changed frequently during the procedure. The other gloves must be removed and the hands surveyed before leaving the hood for any reason. All equipment must also be surveyed for contamination before removal from the hood.
7. An appropriate radiation survey instrument (e.g., pancake style G-M, thin crystal gamma scintillation detector), must be available in the laboratory and should be used to survey hands, clothing, and work surfaces during and after labeling procedures to assure that contamination is not present.
8. Contamination smear surveys must be made of the hood and the lab area immedi- ately after each iodination. Contaminated areas should be decontaminated to less than 100 disintegrations per minute per 100 cm2 above background. Results of these surveys must be included with quarterly reports and a copy must be kept by the approved user.
326 LABELING TECHNIOUES
1. To minimize the release of volatile radioiodine, a closed system should be maintained using charcoal traps, syringes, and septum-sealed vials.
a) Before the radioiodine compound is extracted from the stock vial through the septum, or if the stock vial is to be used as the reaction vessel, the air above the solution in the vial should first be purged through a charcoal trap to equalize the pressure in the vial. A syringe, which has the barrel filled with granular activated charcoal and both ends plugged with cotton or glass wool, should be inserted through the septum to vent the vial. With another syringe, several replacement volumes of air should be injected into the vial.
b) When the stock vial of iodine is used as the reaction vessel, all reactants should be placed in the vial through the septum with the use of an appropriate syringe. Hamilton-type or disposable tuberculine syringes have proven to work satisfactorily.
c) If only a portion of the stock iodine is to be used, that aliquot is to be removed from the stock vial through the septum and placed in another septum-sealed vial to be used as the reaction vial. All reactants should then be added or removed, with syringes, through the septum of this reaction vial.
2. Maintaining the pH of the radioiodine solution above 8.0 will also minimize the release of volatile radioiodine.
RADIOIODINE WASTE
1. Solid Waste Contaminated with Radioiodine
a) All vials, tubes, syringes, disposable pipettes, etc., which may contain volatile radioiodine should be rinsed with 0.1N sodium thiosulfate solution. Then they must be capped or otherwise sealed and placed in a sealed disposable container (Chemical Storehouse, stock # CX12760) before removal from the hood.
b) Other solid waste (gloves, absorbent toweling, etc.) contaminated with radioiodine must be bagged while in the hood and then placed in the solid combustible waste container.
c) Sodium thiosulfate solution (see liquid waste section, below) should be used to rinse reusable pipettes, syringes, etc., to combine with any volatile radioiodine present. The rinse solution must then be disposed of as liquid radioiodine waste. (See liquid waste section below.)
d) If a sephadex type column is used for fractionating, the column should be sealed at both ends and if possible placed in a sealed disposable container.
327 Liquid Radioiodine Waste
a) All liquid radioiodine waste must be stored and transferred in the hood.
b) 0.1N sodium thiosulfate solution must he added to all liquid radioiodine waste.
c) Liquid radioiodine waste with sodium thiosulfate added must be transferred in the hood to a disposable plastic container (Chemical Storehouse stock # CX12760) which is half-filled with granular absorbent (Hi-Dry Granular Absorbent, General Storehouse inventory # GC20830).
d) When the half-filled container of granular absorbent becomes uniformly moistened with liquid radioiodine waste and sodium thiosulfate solution, do NOT add any further liquid. Fill the remaining half of the container with dry absorbent and place the cover tightly on the container. The sealed container should then be placed in the solid non-combustible radioactive waste container. Do not saturate the absorbent above the 1/2 level of the plastic container. Department of Transportation (DOT) regulations require that a layer of dry absorbent remain on the top of the plastic container to assure that there is no unabsorbed liquid.
e) Do not treat liquid radioiodine with sodium hypochlorite or introduce it into liquid radioactive waste containers which contain chlorinated oxidizing agents such as sodium hypochlorite. Chlorinated oxidizing agents will interact with radioiodine solutions in such a way that volatile radioiodine will be generated and released. Sodium hypochlorite solutions are commonly
used to deactivate biohazardous waste materials. , A possible substitute which does not present the problem of generating volatile radioiodine is the use of formalin solutions. If it is necessary to use sodium hypochlorite solutions, separate radioactive waste containers must be used and must be clearly labeled to maintain separation of radioiodine and sodium hypochlorite solutions.
If you have any questions regarding the safe use of volatile radioiodine, contact the Radiation Protection Program at 626-6002.
COMMENTS ON CLEANING UP SPILLS OF RADIOACTIVE MATERIALS
If a spill occurs or special problems arise, IMMEDIATELY contact the Radiation Protection Program at 626-6002 for assistance.
1. Confine the spill to as small an area as possible.
2. Restrict access to the spill area. Persons who may have been contaminated should be monitored immediately. Contaminated clothing should be removed and decontaminate or disposed of in the solid radioactive waste container. Skin surfaces should be washed and resurveyed.
3. The spill should be cleaned up as soon as possible. Do not spread the contamination to an area larger than the original spill. Always clean a spill area by starting at the perimeter and proceeding toward the center of the contaminated area. NOTE: Appropriate protective clothing, gloves, shoe covers and respiratory protection are to be worn by personnel who perform spill clean- u p.
328 SPILLS (cont.)
4. If radioiodine compounds are spilled, moisten absorbent pads with 0.1N thiosulfate solution and carefully place the pads over the spill area. NOTE: Appropriate protective clothing, gloves, shoe covers and respiratory protection are to be worn by personnel who perform spill clean-up.
5. Spilled liquids should be absorbed using disposable towels or granular absorbent. The contaminated towels or absorbent should then be placed in the solid radioactive waste container.
6. The spill area must be decontaminated to less than 100 disintegrations per minute per 100 cm2 above background. A smear survey of the spill area must be performed to verify that the area has been decontaminated.
7. A report of the incident, including the decontamination survey results, must be sent to the Radiation Protection Program.
329
APPENDIX O
RADIATION PROTECTION INSTRUCTIONS FOR ANIMAL CARETAKERS
1. The door to the animal facility must be posted with a radiation caution sign which lists the name of the approved user and his or her telephone number.
2. The animal cages must be posted with an approved University of Minnesota "Caution Radioactive Material" sign, and/or "Radiation Area" sign as needed. The name of the approved user, along with the type and quantity of radioisotope administered to the animals must be listed on the caution sign.
3. Laboratory coats, disposable gloves and other required protective equipment must be worn when caring for animals and during cage cleaning procedures.
4. Personnel radiation monitors may be required in some animal care situations. The Radiation Protection Program will assess the need for such monitors.
5. The Radiation Protection Program will assess the need for evaluation of external radiation exposure levels in the animal care facility, and will perform this required monitoring.
6. The approved user is required to perform routine contamination smear surveys of the animal care facility. A minimum of one survey per month is required. A copy of these survey results must be forwarded to the Radiation Protection Program with the approved user's quarterly report.
7. Animals that have been irradiated by x-ray or external radiation from sealed sources of gamma rays will not present a radiation hazard.
8. If the radioisotopes will be excreted in the urine or feces, absorbent material in a tray must be provided below or within each cage. The absorbent material must be changed periodically and disposed of as radioactive waste. If dogs or other large animals will excrete radioisotopes in the urine or feces, a metabolic cage must be used, and the excrement must be collected and properly stored prior to pickup as radioactive waste. Appropriate containers must be provided in the animal care facility.
9. Prior to washing animal cages, a contamination smear survey must be made by the approved user to assure that contamination levels are reduced to less than 2200 DPM/100 square centimeters (DPM = CPM/fractional efficiency). After reducing contamination below this level, the cages may be washed in a sink designated and labeled for washing items that have low-level contamination. If cages are to be washed in a cage washing facility, the Radiation Protection Program must be contacted (626-6002) and specific wash procedures will be outlined.
10. Animals or portions of animal carcasses containing radioisotopes must be properly disposed of in accordance with the requirements of the Radiation Protection Program. Radioactive animals or portions of animal carcasses must be NOT be placed in radioactive waste containers in the laboratory. This type of waste may need to be refrigerated or frozen depending on lab location and size of animal. The following procedures must be used in packaging and disposal of radioactive animals: (over)
331 Animal Disposal (cont.)
- Place the animal(s) and/or parts of the carcass in sealed transparent or translucent plastic bags (double bag), seal the bags and label with radiation caution tape. Do not use colored plastic bags because this prevents easy inspection of the contents. A label must indicate the radioisotope(s), the activity of each radioisotope in millicuries, and the number and type of animals contained in the bag.
- DO NOT PLACE PADS, TOWELING, ETC. IN THE BAG WITH THE ANIMAL. Do not individually bag or wrap small animal carcasses when more than one animal is placed in the disposal bag. Collect animal bedding, disposable toweling, gauze, pads, etc. in a separate transparent plastic bag and place in the solid radioactive waste can.
- Absorb and remove free-standing blood present in larger animals by using absorbent pads, gauze, etc. and dispose in a sealed bag in the solid radioactive waste can.
- In areas in which same-day pickup from the laboratory cannot be provided, it will be necessary for the approved user to temporarily store the waste in a laboratory freezer to prevent biodegradation.
- Animals which contain 14C and/or 3H must be collected and packaged separately from other radioactive animal waste.
- Animals which contain 131I, 125I, 331P, 51Cr, or other radioisotopes with a half-life of 60 days or less must be packaged separately by isotope.
- Vials or test tubes which contain small amounts of tissue or blood products require special packaging and preparation prior to disposal. Contact the Radiation Protection Program at 626-6002 for instructions.
- For disposal of the animals, call the Radiation Protection Program at 626-6819 between the hours of 9:00-10:30 a.m., or 1:30-3:00 p.m. to request pickup from the laboratory. Be prepared to provide the location of the waste, the weight of the waste, and the isotopes and activity present in the waste.
11. In case of radiation emergencies such as spillage of contaminated waste, contact the Radiation Protection Program in the Department of Environmental Health and Safety (626-6002).
332 APPENDIX U
SOME COMMONLY USED RADIOISOTOPES AND THEIR RELATIVE RADIOTOXICITIES
I. Low Hazard Radioisotopes. (The level of intermediate activity for laboratory use in this group is 1-30 millicuries.)
Radioisotope Half -Life Type of Ionizing Energy of Radiation Emitted Radiation
1. H-3 (tritium) 12 years beta 0.014 MeV 2. C-14 5730 years beta 0.15 MeV
II. Medium Hazard Radioisotopes. (The level of intermediate activity for laboratory use in this group is 100 microcuries - 3 millicuries.) Type of Ionizing Energy of mR/hr-mCi Radioisotope Half- Life Radiation Emitted Radiation at 1 meter
15 hours beta 1.39 MeV 1.84 gamma 2.75 MeV 12.4 hours beta 3.5 MeV 0.15 gamma 1.5 MeV 64 hours gamma 0.19 MeV 0.04 14.3 days beta 1.7 MeV 87 days beta 0.167 MeV 3 x 105years beta 0.714 MeV 45 days beta 0.46 MeV 0.64 gamma 1.10 MeV 18.6 days beta 1.78 MeV 0.05 gamma 1.08 MeV 9. Sr-89 50 days beta 1.46 MeV 10. Au-198 2.7 days beta 0.96 MeV 0.23 gamma 0.41 MeV 11. Hg-203 46 days beta 0.21 MeV 0.13 gamma 0.28 MeV 12. Cr-51 27.8 days gamma 0.32 MeV 0.018 13. P-33 25.2 days beta 0.248 MeV
333
III. High Hazard Radioisotopes. (The level of intermediate activity for laboratory use in this group is 10 microcuries - 300 microcuries.)
Radioisotope Half -Life Type of Ionizing Enernv of mR/hr-mCi Radiation Emitted Radiation at 1 meter
1. Na-22 2.6 years positron 0.54 MeV 1.20 gamma 1.27 MeV 2. Ca-45 164 days beta 0.254 MeV 3. Co-60 5.24 years beta 0.312 MeV 1.32 gamma 1.17, 1.33 MeV 4. Sr-90 28.4 years beta 0.545 MeV 5. I-131 8 days beta 0.6 MeV 0.22 gamma 0.364 MeV 6. I-125 60 days gamma 0.035 MeV 0.07 7. Cs-137 30 years beta 0.514 MeV 0.33 gamma 0.667 MeV
IV. Very High Hazard Radioisotopes (Intermediate laboratory level - 1-10 microcuries). Energy of mR/hr-mCi Radioisotope Half -Life Type of Ionizing Radiation Emitted Radiation at 1 meter 1. Pb-210 22 years beta 0.017 MeV gamma 0.0465 MeV 2. Po-210 138 days alpha 5.3 MeV 3. Ra-226 1620 years alpha 4.7 MeV
335
Toxic, Reactive, Carcinogenic, and Teratogenic Chemicals
Keith Carlson
Industrial Hygiene Officer Department of Environmental Health and Safety Unviersity of Minnesota Minneapolis, MN 55455
Keith Carlson received his B.A. degree (interdepartmental major) in 1974 and his Master's degree (environmental health) in 1978 from the University of Minnesota. He is presently the Industrial Hygiene Officer in the Department of Environmental Health and Safety at the University of Minnesota He is a member of the American Industrial Hygiene Association and a member of the American Conference of Governmental Industrial Hygienists
337
Laboratory Safety
A. Toxicology Background
1. Modes of Entry 2. Dose 3. Exposure Time 4. Chronic/Acute effects
B. Regulatory Agencies
OSHA EPA
C. MSDS
D. Classification of Chemicals/Hazards
E. Fume Hoods
339 I. GENERAL PRINCIPLES
Every laboratory worker should observe the following rules:
1. Know the safety rules and procedures that apply to the work that is being done. Determine the potential hazards (e.g., physical, chemical, biological) and appropriate safety precautions before beginning any new operation.
2. Know the location of and how to use the emergency equipment in your area, as well as how to obtain additional help in an emergency, and be familar with emergency procedures.
3. Know the types of protective equipment available and use the proper type for each job.
4. Be alert to unsafe conditions and actions and call attention to them so that corrections can be made as soon as possible. Someone else's accident can be as dangerous to you as any you might have.
5. Avoid consuming food or beverages or smoking in areas where chemicals are being used or stored.
6. Avoid hazards to the environment by following accepted waste disposal procedures. Chemical reactions may require traps or scrubbing devices to prevent the escape of toxic. substances.
7. Be certain all chemicals are correctly and clearly labeled. Post warning signs when unusual hazards, such as radiation, laser operations, flammable materials, biological hazards, or other special problems exist.
8. Remain out of the area of a fire or personal injury unless it is your responsibility to help meet the emergency. Curious bystanders interfere with rescue and emergency personnel and endanger themselves.
9. Avoid distracting or startling any other worker. Practical jokes or horseplay cannot be tolerated at any time.
10. Use equipment only for its designed purpose.
11. Position and clamp reaction apparatus thoughtfully in order to permit manipulation without the need to move the apparatus until the entire reaction is completed. Combine reagents in appropriate order, and avoid adding solids to hot liquids.
12. Think, act, and encourage safety until it becomes a habit.
II. Health and Hygiene
Laboratory workers should observe the following health practices:
1. Wear appropriate eye protection at all times.
340 2. Use protective apparel, including face shields, gloves, and other special clothing or foot wear as needed.
3. Confine long hair and loose clothing when in the laboratory
4. Do not use mouth suction to pipet chemicals or to start a siphon; a pipet bulb or an aspirator should be used to provide vacuum.
5. Avoid exposure to gases, vapors, and aerosols. Use appropriate safety equipment whenever such exposure is likely.
6. Wash well before leaving the laboratory area. However, avoid the use of solvents for washing the skin. They remove the natural protective oils from the skin and can cause irritation and inflammation. In some cases, washing with a solvent might facilitate absorption of a toxic material.
III. FOOD HANDLING
Contamination of food, drink, and smoking materials is a potential route for exposure to toxic substances. Food should be stored, handled, and consumed in an area free of hazardous substances.
1. Well-defined areas should be established for storage and consumption of food and beverages. No food should be stored or consumed outside of this area.
2. Areas where food is permitted should be prominently marked and a warning sign (e.g., EATING AREA -- NO CHEMICALS) posted. No chemicals or chemical equipment should be allowed in such areas.
3. Consumption of food or beverages and smoking should not be permitted in areas where laboratory operations are being carried out.
4. Glassware or utensils that have been used for laboratory operations should never be used to prepare or consume food or beverages. Laboratory refrigerators, ice chests, cold rooms, and such should not be used for food storage; separate equipment should be dedicated to that use and prominently labeled.
IV. HOUSEKEEPING
There is a definite relationship between safety performance and orderliness in the laboratory. When housekeeping standards fall, safety performance inevitably deteriorates. The work area should be kept clean, and chemicals and equipment should be properly labeled and stored.
1. Work areas should be kept clean and free from obstructions. Cleanup should follow the completion of any operation or at the end of each day.
2. Waste should be deposited in appropriate receptacles.
341 3. Spilled chemicals should be cleaned up immediately and disposed of properly. Disposal procedures should be established and all laboratory personnel should be informed of them; the effects of other laboratory accidents should also be cleaned up promptly.
4. Unlabeled containers and chemical wastes should be disposed of promptly, by using appropriate procedures. Such materials, as well as chemicals that are no longer needed, should not accumulate in the laboratory.
5. Floors should be cleaned regularly; accumulated dust, chromatography adsorbents, and other assorted chemicals pose respiratory hazards.
6. Stairways and hallways should not be used as storage areas.
7. Access to exits, emergency equipment, controls, and such should never be blocked.
8. Equipment and chemicals should be stored proplery; clutter should be minimized.
342 Infectious Agents
James Lauer
Biosafety Officer Department of Environmental Health and Safety University of Minnesota Minneapolis, MN 55455
James Lauer received his B.A. in Biology from St. Cloud State University in 1969 and his M.P.H. degree in Environmental Health from the University of Minnesota in 1976. He is currently the Biosafety Officer in the Department of Environmental Health and Safety at the University of Minnesota.
343
Introduction
Research involving biological agents capable of causing disease in man has been conducted throughout the past century. In the last few decades, research with known or potentially hazardous agents has been greatly intensified. This intensification is primarily due to rapid advancements in disciplines such as Genetics, Cell Biology, Immunology, Biochemistry, Virology, Medicine (especially transplan- tation), and Animal and Plant Parasitology. During this century of research, there has been only a few incidents of spread of disease outside of the laboratory and the number of laboratory-acquired di- seases have been relatively few. Unfortunately, such a record is not comforting for those suffering disease complications or death. This is especially true when one considered that the factors responsible for these disease incidents, were, in most situations, controllable.
Today's presentation will examine biohazard containment, or in other words, controls which prevent the spread of biohazardous agents. The implementation of such controls necessitates an understanding of the Infectious Disease Process.
Infectious Disease Process
The Infectious Disease Process can be defined in simple terms as the interreactions between the biological agents, the environment and the host. This interreaction has six essential components. These components are depicted as a circular linked chain (see handout) and are labeled as follows:
Etiological Agent (causitive agent) Reservoir (typical habitat of agent: man, animal, soil, etc.) Escape from Reservoir (typical mode(s) by which agents escape from reservoir Transmission (typical mode(s) of transmission between reservoir and host) Entry into Host (typical mode(s) of entry into host) Host Susceptibility (host prediliction to agents disease capacities)
The disease process depends on the continuity of the chain. If even one of the links is broken, the disease process is interrupted. To understand how the disease process works, some specific examples will be examined.
Smallpox (variola major, minor, etc.) Hepatitis Type B Salmonella L.D.B.
One of the best controls against biohazardous agents is by de- creasing the host susceptibility through vaccination. However, this type of control is only possible for a handfull of agents. Physical controls are also used to interrupt the disease process.
345 Physical Controls
Physical controls are directed at the escape and transmission component of the disease process. They separate the agent from the host by a physical barrier. Secondary physical barriers separate the laboratory environment from the outside environment. Certain laboratory design criteria for such control depends on the risk classification of the agent.
High Containment Facility Special Design Facility
A more important type of physical control is known as primary physical barriers. These barriers separate the agent from the person during the actual work. Such barriers include:
BSC LFB SC Safety blender Safety trunnion cup for centrifugation Gloves and gowns Etc.
Microbiological Technique
Another type of control, and in my opinion, the most important, is what is referred to as good microbiological technique. Without such good techniques, the other controls are practically useless, The lab- oratory techniques, which include good personnel hygiene practices, are as follows:
Use pipetting aids The appropriate use of gloves and gowns Avoid the use of syringes and needles Good handwashing practices are essentiail Hand habits Flaming and transfer techniques Smoking and the storage or consumption of food or beverages must be prohibited Etc.
Case Studies Emphasizing the Importance of Controls
1. Hepatitis Type B in Hospital 2. Smallpox in England
346 347 348 349 CLEAN-UP OF BIOHAZARDOUS SPILLS
I. Biohazard Spill in a Laminar Flow Biological Safety Cabinet*
In the LFBSC, chemical decontamination procedures should be initiated
once while the cabinet continues to operate to prevent escape of
contaminants from the cabinet.
A. Spray or wipe walls, work surfaces, and equipment with 2%
Wescodyne** (or other appropriate disinfectant detergent). A
disinfectant detergent has the advantage of detergent activity
which is important because extraneous organic substances frequently
interfere with the reaction between a microbe and microbiocidal
agent. Operator should wear gloves during this procedure.
B. Flood top tray, drain pans, and catch basins below work surface
with disinfectant and allow to stand 10-15 minutes.
C. Dump excess disinfectant from tray and drain pans into cabinet
base. Lift out tray and removable exhaust grille work. Wipe off
top and bottom (underside) surfaces with disinfectant sponge or
cloth. Replace in position. Gloves, cloth or sponge should be
discarded in an autoclave pan and autoclaved.
D. Drain disinfectant from cabinet base into appropriate container
and autoclave according to standard procedures.
* Procedure adapted from "Laminar Flow Biological Safety Cabinets," A Training Manual for Biomedical Investigators, National Cancer Institute.
** West Chemical Products, Inc. 16-42 West Street, Long Island City, New York. 11101
350 II. Biohazard Spill Outside a Laninar Flow Biological Safety Cabinet
A. Holding your breath, leave the room immediately and close the door.
B. Warn others not to enter the contaminated area.
C. Remove and containerize contaminated garments for autoclaving and
thoroughly wash hands and face.
D. Wait 30 minutes to allow dissipation of aerosals created by the spill.
E. Don a long-sleeve gown, mask, and rubber gloves before reentering the
room. (For a high risk agent, a jumpsuit with tight-fitting wrists
and use of a respirator should be considered.)
F. Pour a germicidal solution (5% Wescodyne or 5% Hypochlorite are
recommended) around the spill and allow to flow into the spill.
Paper towels soaked with the germicide may be used to cover the
area. To minimize reaerosolization, avoid pouring the germicidal
solution directly onto the spill.
G. Let stand 20 minutes to allow adequate disinfectant contact time.
H. Using an autoclavable dust pan and squeegee, transfer all contami-
nated materials (paper towels, glass, liquid, gloves, etc. ) into
a deep autoclave pan. Cover the pan with aluminum foil or other
suitable cover and autoclave according to standard directions.
I. The dust pan and squeegee should be placed in an autoclavable bag and autoclaved according to standard directions. Contact of
reusable items with non-autoclavable plastic bags should be
avoided--separation of the plastic after autoclaving can be
very difficult.
351 III. Radioactive biohazard Spill Outside a Laminar Flow Biological Safety
Cabinet
In the unlikely event that a biohazardous spill also involves a
radiation hazard, the clean-up procedure may have to be modified,
depending on an evaluation of the risk assessment of relative
biological and radiological hazard.
Laboratories handling radioactive substances must have the
services of a designated radiation protection officer available for
consultation.
The following procedure indicates suggested variations from the
biohazard spill procedure (above) that should be considered whena
radioactive biohazard spill occurs outside a safety cabinet.*
A. Holding your breath, leave the room immediately and close the door.
B. Warn others not to enter the contaminated area.
C. Remove and containerize contaminated garments for autoclaving and
thoroughly wash hands and face.
*Personnel should be surveyed for radiation hazard before and
after clothing change and wash-up.
D. Wait thirty minutes to allow dissipation of aerosols created by
the spill.
*Before clean-up procedures begin, a radiation protection officer
should survey the spill for external radiation hazard to determine
the relative degree of risk.
* Changes in procedure have been starred and underlined.
352 E. Don a long-sleeve gown, mask, and rubber gloves before reentering
the room. (For a high risk agent, a jumpsuit with tight-fitting
sleeves and use of a respirator should be considered.)
F. Pour a germicidal solution (5% Wescodyne or 5% Hypochlorite are
recommended) around the spill and allow to flow into the spill.
Paper towels soaked with the germicide may be used to cover the area.
To minimize reaerosolization, avoid pouring the germicidal solution
directly onto the spill.
G. Let stand 20 minutes to allow adequate disinfectant contact tine.
H. *In most cases, the spill will involve 14C or 3H, which present
no external hazard. However, if more energetic beta or gamma
emitters are involved, care must be taken to prevent hand and body
radiation exposure. The radiation protection officer must make this
determination before the clean-up operation is begun.
If the radiation protection officer approves, the biohazard
handling procedure may begin: Using an autoclavable dust pan and
squeegee, transfer all contaminated materials (paper towels, glass,
liquid, gloves, etc.) into a deep autoclave pan. Cover the pan
with aluminum foil or other suitable cover and autoclave according
to standard directions.
*If the radiation protection officer determines that the radiation
hazard is too great, the material must not be autoclaved. In
that case, sufficient germicidal solution to immerse the contents
should be added to the waste container. The cover should be sealed
353 with waterproof tape, and the container stored and handled for
disposal as radioactive waste. Radioactive and biohazard warning
symbols should be affixed to the waste container.
I. If autoclaving has been approved, the dust pan and squeegee should
be placed in an autoclavable bag and autoclaved according to standard
directions. Contact of reusable items with plastic bags should be
avoided--separation of the plastic after autoclaving can be difficult.
*A final radioactive survey should be made of the spill area, dust
pan, and squeegee with a Geiger counter, or a smear should be taken
and counted in a liquid scintillation counter. Twice (2X) background
count indicates the need for further decontamination.
354 Fire and Physical Hazards
Ray Arntson
Safety Officer Department of Environmental Health and Safety University of Minneosta Minneapolis, MN 55455
Raymond E. Arntson received his B.S. and M.P.H. degrees from the University of Minnesota in the area of Occupational Environmental Health. He has worked in the safety and health field within the United States Navy and the insurance industry. His present position is Safety Officer in the Department of Environmental Health and Safety an the University of Minneosta, with a focus on occupational safety and fire prevention.
355
Laboratory Fire Safety
I. The Fire Problem of Laboratories
A. Factors of Origin
B. Factors of Spread
C. General Concepts
II. Methods of Fire Control
A. Hazard Classification
B. Physical Design and Construction
C. Fire Protection and Alarm
D. Laboratory Ventilation and Fume Hoods
E. Flammable and Combustible Liquids Control
F. Compressed Gas and Fixed Pi ping Systems
III. Laboratory Operations
A. Assign Responsible Individual
B. Assess Operation and Apparatus
C. Provide Hazard Identification
357
Italic = New or Revised January 1, 1987 Safety Guidelines Department of Environmental Health and Safety
Design and Construction of Laboratories Using Chemicals
I. Laboratories must be designed to conform with one of the following National Fire Protection Association (NFPA) standards.
A. NFPA 45 - Standard on Fire Protection for Laboratories Using Chemicals
B. NFPA 99, Chapter 7 - Laboratories in Health Related Institutions
II. Laboratories must also conform with NFPA 101, Code for Safety to Life From Fire In Buildings and Structures. Depending upon the building in which the laboratory will be located and the nature of the work performed in the laboratory, a laboratory will be considered to be within one of the following occupancies:
A. Business
B. Health Care
C. Industrial
III. Compartmentation of each laboratory unit must be achieved by providing it with at least:
A. One-hour fire separation from adjacent laboratories, or Other areas.
B. Self-closing fire doors with at least 3/4 hour ratinq (NFPA 101, 6-2.2.5)
C. Class A interior finishes
D. Class I floor finishes
359 IV. Access to two or more exits must be provided within each laboratory. Aisles servinq a single work area must be a minimum of 36" wide. Double aisles must be a minimum of 60" wide. Avoid aisles longer than 20 feet. Arrange furniture for easy access to an exit from any point in the laboratory.
V. Doors in laboratories where hazardous materials are used must swing in the direction of exit.
VI. Faucets must be provided with vacuum breakers, or a special laboratory water supply must be provided as required by the stat? plumbing code.
VII. In a lab equipped with a fume hood, a safety shower and an eyewash must be provided. In other laboratories using chemicals at least an eyewash will be required. If it is feasible, water supply should be controlled to a temperature between 60º and 95º. Refer to the Department of Environmental and Safety Recommended Practices for the Installation of Emergency Eyewashes and Safety Showers.
VIII. The laboratory user must be consulted to determine the quantities of flammable and combustible liquids which will be present in the laboratory. These materials must be stored according to the University of Minnesota Standard for the Quantity of Flammable and Combustible Liquids in University of Minnesota Laboratories and NFPA 30, the Flammable and Combustible Liquids Code.
IX. Storage and supply systems for compressed and liquified gases shall comply with requirements of the NFPA and ANSI. Standards which should be consulted include:
(a) NFPA 50, Standard for Bulk Oxygen Systems at Consumer Sites
(b) NFPA 50A, Standard for Gaseous Hydrogen Systems at Consumer Sites
(c) NFPA 50B, Liquefied Hydrogen Systems at Consumer Sites
(d) NFPA 5 1, Design and Installation of Oxygen-Fuel Gas Systems for Cutting and Welding
360 (e) NFPA 56F, Standard for Nonflammable Medical Gas Systems
(f) NFPA 54, National Fuel Gas Code
(g) NFPA 58, Standard for the Storage and Handling of Liquefied Petroleum Gases
(h) NFPA 99, Standard for Health Care Facilities Chapter 3 - Use of Inhalation Anesthetics (flammable and non-flammable) Chapter 4 - Use of Inhalation Anesthetics in Ambulatory Care Facilities Chapter 5 - Respiratory Therapy
(i) ANSl B31.1.0, Power Piping (including Addenda B31.1.0a, B31.1.1.0c, and B31.1.1.0d)
(j) ANSl B31.2, Fuel Gas Piping (k) ANSI B31.3, Petroleum Refinery Piping
X. Systems for other gases shall comply with manufacturers designs and specifications. The Handbook of Compressed Gases by the Compressed Gas Association and the Matheson Gas Data Book by Matheson Gas Products may be consulted.
XI. When a central supply of flammable, combustible or oxidizing gases is provided, shut-off valves in accessible locations must be provided; they must be outside of the areas in which the gases are used. These shut-offs are in addition to those at the points of supply and use. They may be located in either a distant area of the lab or in the corridor if security is not a problem..
XII. Controls for air, gas and other utilities must be color coded and labeled (different control handle configurations are desirable).
361 XI I I. Fire extinguishers must be provided to meet the requirements of NFPA 10. A five-pound multipurpose dry chemical fire extinguisher must be provided for each laboratory unit. The extinguisher must be mounted near an exit from the unit.
XIV. Laboratories using chemicals must be under negative pressure, with respect to adjacent areas. No recirculation of exhaust air from laboratories is permitted.
XV. Laboratory fume hoods must meet the Department of Environmental Hood Design Criteria.
XVI. Electrical systems must meet the requirements of NFPA 70, the National Electric Code. Outlets must be provided for fixed appliances and one duplex must be provided per each 3 to 5 feet of bench length.
The following items are preferred design criteria for chemical laboratories and may be required when appropriate:
1. Metal laboratory furniture with stainless or synthetic stone benchtop.
2. Wall cabinets with a continuous enclosed front plane to the ceiling.
3. Glass waste lines.
4. A glassware cleaning sink at least 12" deep
5. An emergency equipment room, cabinet or kit, including an air supply mask, chemical neutralizer, liquid absorbents and protective clothing at a minimum.
362 Italics=New or Revised January 1, 1987 Safety Standard Department of Environmental Health and Safety
General Purpose Fume Hoods and Additional Requirements For Radioisotope and Perchloric Acid Hoods
General purpose fume hoods are provided for operations using flammables or toxic chemicals. A multipurpose hood design which will provide adequate safety for changes in use that may occur during the useful life of the hood is required. Compliance with the following design criteria will support user safety. These criteria are supplemental to SAMA Standard LF 10-1980, University of Minnesota Construction Standards, and to more specific material and equipment specifications.
Fume Hood Construction and Design
1. Non-combustible construction is required.
2. It shall be of air foil design with the bottom foil raised to provide a one-inch clear opening between the foil and the work top.
3. Provide a vertical sliding safety glass sash operable with one band from any point on the bottom.
4. On 14' and larger hoods, 15" wide horizontal sliding safety shield shall be provided, supported to resist pressure displacement. It shall be suspended on bearings or slide in an easily cleanable channel.
363 5. An air by-pass designed to prevent hood face velocity from exceeding 200 feet per minute shall be provided.
6. Electrical outlets shall be located outside of the fume hood interior.
7. Utility (gas, water, vacuum, etc.) controls must be located on the exterior of the hood with utility outlets mounted on the interior side wall. Controls shall be identified.
8. A liquid-tight work surface to contain at least 3/8" liquid depth shall be identified.
9. A baffle system that allows air to be drawn evenly to the top, middle, bottom of the hood and so arranged that is possible to adjust the flow of the air but not shut it down completely shall be provided. * The top baffle must have a opening width limited to no more than 3/4 inch.
10. One fume hood base cabinet per laboratory (two cabinets are acceptable in large laboratories) may be used as a storage cabinet for flammable liquids. Cabinet top, bottom, sides and doors shall not be less than 18 gauge steel, double-walled construction with a 1 1/2" air space between the walls. * The doors shall be equipped with a 3 point latch system. All joints shall be welded or screwed to provide rigid enclosures. A liquid tight pan capable of containing a 2" depth of liquid shall be provided. The cabinet shall be ventilated at a minimum rate of 5 CFM with a stainless steel duct penetrating behind the baffle at least 1" above the work surface. Flame arrestors shall be provided on cabinet vents. Make up air supply for the cabinet shall be taken from the pipe space behind the cabinet; supply vents shall not be placed on the front or side of the cabinet. The exterior exposed surfaces of the storage cabinet shall be painted with yellow paint and must be labeled with at least 1" high black letters with 0.25" stroke "FLAMMABLE STORAGE" or "FLAMMABLE LIQUIDS"
364 11. Acid storage base cabinets shall be constructed to resist corrosion. Ventilation shall be provided in the cabinet at the rate of 5 CFM. Vent pipes shall extend above the work surface and behind the baffle in the same manner as with flammable storage cabinets although flame arrestors are not required. Vents may be provided in the cabinet door. Cabinets shall be provided with a liquid tight pan, capable of holding a 2" depth of liquid. Cabinets shall be labeled in at at least 1" letters with 0.25" stroke "ACID STORAGE."
12. Drying base cabinets shall not be installed under fume hoods.
13. Interior lighting shal l be vapor sealed and covered with a safety glass lens. Illumination levels at the working surface shall be at least 80 foot-candles.
14. Fume hoods shall, whenever feasible,be located in the distal corners of a lab and away from high traffic areas. This is to avoid locations in a traffic pat tern which will cause conflicting convection currents and to avoid blocking an exit if there is fume hood fire.
Filter Enclosures*
15. When a HEPA f liter enclosure is provided, it must provide for' easy bag-in, bag-out of filters so there is no exposure to maintenance staff. The filter enclosure shall be located in such a position as to allow easy installation and removal of the filters.
16. On hoods with filter enclosures, an airflow indicator shall be provided in a clearly visible location to indicate pressure drop across the filter.
365 17. Filter enclosure:
a. The filter enclosure shall be of stainless steel construction. b. It shall be designed to provide a tight seal between the enclosure and the filter frame (use levered cams on four corners of filter frame and gasketed filter frames to prevent air leakage). c. The filter enclosure must provide for installatioin of a 11 1 /2" x 24" x 24 or 5 3/4" x 24" x 24" HEPA filter, and a 2" x 24" x 24" pre-filter. d. The filter access door(s) shall be gasketed to prevent air leakage. e. The duct between the hood and the filter shall be of stainless steel construction.
18. If the hood is installed with a HEPA filter enclosure, the open face velocity shall be 150 fpm to allow for filter loading. If the hood is not installed with a filter, the face velocity shall be 100 fpm but reserve capacity for a filter must be engineered into the fan design.
* Note: Enclosure must be equal to Mine Safety Equipment Company Ultra-Lok Series U.
Exhaust and Ductwork Requirements
1 9. Systems shall generally be installed in accordance with the r equirements of NFPA 91-1983, Standard for the Installation of Blower and Exhaust Systems.
20. The fire ratingof laboratory units and other building fire compartments must be protected. Stainless steel and sheet metal ductwork will usually be considered to provide one hour fire separation. When more than one hour separation or the use of other ductwork materials are proposed, construction of 3 suitable fire rating must also be proposed to protect the system.
366 21. General purpose fume hoods shall be individually ducted - except that up to four hoods, located in the same room, may be connected to a common exhaust duct leading from that room to an exhaust fan. If more than one hood is connected to an exhaust duct, a balanced undampered drop must be engineered. Fume hoods provided with HEPA filters enclosures shall always be individually ducted
22. Fire dampers or other restrictions shall not be placed in any chemical fume exhaust duct.
23. Fume hood exhaust systems shall function independently of the general building HVAC system. Fume hood exhaust volumes shall not be modulated or controlled to balance air requirements for air conditioningor heating
24. Associated equipment in the same room, such as flammable liquid storage cabinets, biological safety cabinets and atomic absorption units, should beprovided with an independant exhaust system. However, after review by the Department of Environmental Health and Safety, associated equipment might be permitted to be ducted into the fume hood. On hoods with HEPA filter enclosures, associated equipment must be connected between the hood and the enclosure.
25. Clearly visible airflow indicators shall be installed on new laboratory hoods or on existing fume hoods when they are modified
26. Fume hood exhaust ductwork shall be operated with negative static pressure in the ductwork through all spaces within the building. Fume hood exhaust fans shall be located on the roof of the building or In a ventilated equipment room just below the roof of the building.
Fan discharge ducts shall discharge vertically through the roof and terminate at least seven feet above the roof. Seal the discharge duct airtight.
367 27. Stack design and discharge velocity shall distribute contaminants outside the eddy current envelope of the building. On structures with roof areas at more that none level, discharge ducts within 30 feet of a higher level shall terminate at a point at least seven feet above the elevation of the higher level.
28. Air velocity on the suction of the fan shall be a minimum of 1,000 fpm and should not exceed 2,000 fpm under any conditions. High duct velocity results in high noise levels, excessive leakage and high power consumption. An optimum velocity of 1,200 fpm is recommended. Ductwork should be round to assure uniform air flow, rectangular ductwork will be acceptable only when condition require its use.
Fume Hood Operation
29. Fume hoods shall run continuously. On-off control to be by Physical Plant only.
30. Open-face velocity average shall be 100 fpm + or - 10 fpm. Individual face velocities shall not exceed 20% of the open- face velocity average when readings are taken in the center of several square grids measured in the plane of the face opening (see SAMA Standard LF 10-1980 for the recommended sampling grid! Open-face" for two-speed fume hoods shall be the maximum possible opening while the "open-face" for reduced capacity fume hood shall be the 18" sash lock level (see number 32.b below).
31. When calculating exhaust volumes required for a fume hood, the 15" wide horizontal sliding safety shield shall not be taken into consideration for any hood less than 4 feet wide because they are often removed by users: On a reduced volume fume hood, the safety shield shall not be taken into consideration for any hood less than 6 feet wide. This standard may result in higher face velocities.
368 32. To conserve energy, fume hoods may be provided with two - speed fans of a reduced capacity fume hood exhaust system.
a. Two-speed system
1 ) Two-speed fans, when provided, shall function at low speed when the sash is closed or within 2" of the working surface. Activation device shall be fail-safe so that if the device fails the fan will operate at high speed.
2) High speed and low speed signal lights shall be provided and labeled to indicate their function.
3) Provide an easily readable sign: "Conserve Energy, Close Hood Sash When Possible to Activate Lower Fan Speed."
b. Reduced capacity fume hood exhaust system
1) The vertical sliding safety glass sash shall havea positive steel mechanical latch to prevent opening above 18" without operator intervention. Latch shall be operable with one hand and allow unobstructed closing of sash from any position.
2) A flashing red warning light shall be installed to operate whenever vertical sash is above 18". The warning light shall be adjacent to latch control and be observable by hood operator.
3) An easily readable sign shall be provided adjacent to the sash height warning light: "Caution: Face Velocity Reduced - Position Sash Below 18" for Continued Use".
369 Supply Air Requirements
33. Auxiliary air supply hoods are not desirableunder usual circumstances at the University. Before proposing an auxiliary air supply hood, contact the Department of Environmental Health and Safety
34. Room cross drafts shall be avoided. Whenever possible air should be provide in a diffuse manner from behind an operator.
Additional Requirements for a Radioisotope Fume Hood
1. A raised cup sink shall be provided. It shall be raised (1/4- 5/16") above the work surface but shall be lower by (1/16") than the raised margins of the work surface. The front lip of the work surface shall be raised at least 1/2 inch.
2. The interior lining and baffles of the fume hood shall be constructed of stainless steel.
3. The work surface shall be capable of supporting up to 200 pounds/foot2 of shielding material.
4. Corners shall be smooth and seamless and radiused 1 /2" except at the top.
5. An absolute filter enclosure may be required Contact the Department of Environmental Health and Safety for licensure requirements.
Additional Requirements for a Perchloric Acid Fume Hoods
1. Hoods and exhaust duct work shall be constructed of acid resistant, non -reactive, impervious materials:
2. Duct work shall take the shortest and straightest path to the outside Positive drainage shall be provided back to the hood.
370 3. A water spray system shall be provided to wash down the entire exhaust system from the hood interior behind the baffle, through the fan, up to the roof line . The hood work surface shall be watertight with a minimum depression of 1/2" at the front and sides. An integral trough shall be provided at the rear of the hood to collect wash down water. The baffle shall be removable for cleaning.
4. Washdown must be easily initiated by the user. Initiation of the washdown cycle must start a minimum 1/2 hour washdown. Since the washdown of a contaminated hood requires up to 24 hours of continuous washing, manual control of the cycles' duration must also be possible.
5. Provide an easily readable placard stating "Wash down after use, do not use perchloric acid with incompatible materials -- e.g., acetic acid, bismuth and its alloys, alcohol, paper, wood, grease or oils".
371 Italics=New or Revised January 1, 1987 Safety Standard Department of Environmental Health and Safety
Installlation of Emergency Eyewashes and Safety Showers
These recommendations apply to the installation of eyewashes and safety showers in new buildings or renovation projects at the University of Minnesota. Eyewashes and safety showers are effective means of flushing corrosive materials out of the eyes or off the body. Safety showers can also be used to extinguish clothing fires.
General Considerations
1. Emergency eyewashes and safety showers are not a substitute for proper protective devices. To protect against flying particles and splashing injurious liquids, persons shall wear eye and face protection and protective clothing.
2. Eyewashes and safety showers shall be located in areas where the eyes or body may be exposed to corrosive chemicals, e.g. laboratories, battery operations, corrosive dip tanks.
3. Eyewashes and safety showers shall be located so that the maximum distance from the hazard does not exceed 1 00 feet and so that they can be reached within 10 seconds.
4. Eyewashes and safety showers should be located in the normal path of egress. For example, in a laboratory, the location should be near a corridor door.
5. Water supplied to eyewashes and safety showers should be tempered. A temperature of 60º-95º is considered optimum.
372 6. Only potable water shall be used to supply eyewashes and safety showers.
7. The activation devices of eyewashes and safety showers must be uniform throughout a building.
8. All eyewashes and safety showers must be uniformly marked throughout a building.
Eyewash/Face Washes
1. Eyewashes should be of a type that provide a curtain of water over the entire facial area. Streams of water should be simultaneously released from two sides to clean foreign particles or liquids from both the eyes and facial area.
2. Eyewashes shall have a flow rate of at least 3 gallons per minute at a pressure of 30 psi supply to fixture.
3. The eyewash control should be of the paddle type with dimensions of approximately 4 x 4". The control should require no more than 10 ounces of force for activation. The valve should be designed to remain activated until intentionally shut off.
4. The recommended distance from the floor to the eyewash jets is 33- 45 inches.
5. All eyewash fixtures are required by the State Plumbing Code to be drained to sewer. Safety Showers
1. Safety showers shall be of deluge type with a continuous flow valve. Such a valve would require a second pull of the cord or ring, for example, to deactivate the shower. The valve shall remain activated until intentionally shut off.
2. Safety showers may be ceiling or wall mounted, and they may be installed in combination with an eyewash fixture. The supply lines and connections of combination units shall not create a contact hazard for persons using the eyewash.
373 3. Safety showers should provide a head discharge of at least 30 gallons per minute at a pressure of 30 psi.
4. The recommended distance from the floor to the shower is 82-96"
5. Activation of the shower may be by wall cord, ring and chain or pull bar. The location of the activating device should be designed so that it will not be in the way of normal operations. This is to prevent accidental discharge.
6. The provision of a floor drain is a desirable option for a safety shower.
374 QUANTITY OF FLAMMABLE AND COMBUSTIBLE LIQUIDS October 12, 1983 IN UNIVERSITY OF MINNESOTA LABORATORIES
This standard shall apply to University laboratory facilities including instructional, research, health care and general purpose, to regulate the quantity of flammable and combustible liquids within the laboratory. It addresses quantities in use as well as those in storage in approved storage cabinets. It does not address storage in specifically constructed flammable liquids storage rooms.
For use within this standard, the following definitions shall apply.
Flammable Liquid: A liquid having a flashpoint below 100ºF and having a vapor pressure not exceeding 40 pounds per square inch at 100ºF shall be known as a Class I liquid. Class I liquids shall be sub-divided as follows:
Class IA shall include those having flashpoints below 73ºF and having a boiling point below 100ºF.
Class IB shall include those having flashpoints below 73ºF and having boiling point at or above 100°F.
Class IC shall include those having flashpoints at or above 73ºF and below 100ºF.
Combustible Liquid: A liquid having a flashpoint at or above 100ºF. Com- bustible liquids shall be sub-divided as follows:
Class II shall include those having flashpoints at or above 100º F and below 140ºF.
Class IIIA liquids shall include those having flashpoints at or above 140ºF and below 200°F.
Class IIIB liquids shall include those having flashpoints at or above 200°F.
Laboratory Unit: A enclosed space used for experiments or tests. Laboratory units may or may not include offices, laboratories, and other contiguous rooms maintained for or used by laboratory personnel, and corridors within the unit. It may contain one or more separate laboratory work areas. It may be an entire building. It must, however, be separated from other building areas by appro- priate fire resistive construction having at least a one hour fire resistive rating. Requirements
1. Maximum quantities of flammable and combustible liquids in research and general purpose laboratory units shall be in accordance with the following table.
375 Maximum Quantities of Flammable and Combustible Liquids in Laboratory Units Outside of Approved Flammable Liquid Storage Rooms
Excluding Quantities in Storage Cabinets and Safety Cans
Maximum Quantity per Maximum Quantity per*** Liquid Class 100 sq. ft. of Lab Unit Lab Unit
I* 2 gallons 75 gallons
I, II and IIIA** 4 gallons 100 gallons
Including Quantities in Storage Cabinets and Storage Cans
I* 4 gallons 150 gallons
I,II and IIIA** 8 gallons 200 gallons
Notes: *Class I liquid maximum shall be inclusive of Class IA, IB and IC collectively.
**The Maximum quantities of Class I liquids shall not exceed the quantities specified for Class I liquids alone.
***The more restrictive quantity, based on either the Maximum quantity per 100 sq. ft. of Lab Unit or Maximum Quantity per Lab Unit, shall apply in all cases.
376 2. Maximum quantities of flammable and combustible liquids in instruc- tional laboratory units shall not exceed 50% of that allowable in research and general purpose laboratories.
3. Maximumquantities of flammable and combustible liquids in use, out- side of approved storage cabinets in health care laboratories, shall not exceed 2 gallons per 1,000 square feet of laboratory unit. The total capacity of all storage cabinets, within health care laboratories, shall not exceed 12 gallons per 1,000 square feet of laboratory unit.
4. Maximum allowable container size for use in all laboratories shall be in accordance with the following table.
Maximum Allowable Container Size for Use in Laboratories Using Chemicals
Flammable Liquids Combustible Liquids Container Type 1A 1B 1C II IIIA
Glass 1 pt* 1 qt* 1 gal 1 gal 5 gal
Metal (other than DOT Drums) or Approved Plastic 1 gal 5gal** 5 gal** 5 gal** 5 gal Safety Cans 2 gal 5 gal** 5 gal** 5 gal** 5 gal
Notes: *Glass containers, as large as one gallon, may be use if needed, and if the required purity would be adversely affected by storage in a metal or an approved plastic container, or if the liquid would cause excessive corrosion, or degradation of metal or approved plastic container.
**In instruction laboratory work areas, no container for Class I or Class II liquids shall exceed a capacity of one gallon, except that safety cans may be of two-gallon capacity.
5. Flammable or combustible liquids shall not be stored in ordinary refrigerators. Storage of flammable or combustible liquids in well- sealed containers is permissible in approved flammable materials storage refrigerators or in refrigerators approved for Class I, Division I, Group C and D. The outside of doors to laboratory re- frigerators shall be labeled to denote whether or not they are acceptable for storage of flammable or combustible liquids.
377 6. Incompatible materials shall be segregated to prevent accidental contact with one another.
7. Flammable and combustible liquids stored in the open in, the labora- tory work area shall be kept to the minimum necessary for the work being conducted.
8. All containers used for storage of flammable and combustible liquids shall be labeled as to content in accordance with good laboratory practice.
378 Use and Storage of Compressed Gas Cylinders in University of Minnesota Buildings This Standard applies to all compressed gas cylinders in University buildings. Special uses such as administration, of combustible anesthetics, are the responsibility of persons associated with such applications. Requirements 1. Cylinders shall have the name of the chemical appearing in legible form on the cylinder. A color code is not satisfactory designation. 2. Cylinders shall be securely held in an upright position. String, wire or similar makeshift materials are not acceptable. 3. Cylinders shall be located so they are not exposed to direct flame or heat in excess of 125 degrees Fahrenheit. 4. Cylinders not in use shall have the valve protective cop securely in place. (Lecture bottles are an exception. ) 5. Cylinders shall be moved only in suitable hand carts. 6. Cylinders containing flammable gas shall be used and stored in a ventilated area. (10 air changes par hour, minimum.) No other gases or chemicals shall be stored in the same area. 7. Cylinders containing toxic or corrosive gases shall be used and stored in well-ventilated areas. 8. Cylinders containing corrosive gas shall be returned to the supplier no later than six months from time of first use. Cylinder and regulator valves shall be opened and closed frequently during the period of use. 9. Cylinders discharging into liquids or closed systems containing other chemicals shall have a trap, check valve, of vacuum breaker between cylinder and system or liquid. 10. Systems mixing two or more gases shall be provided with necessary control or check valves to prevent contamination of the separate gas sources. 11. Closed systems, or any arrangement that might accidentally become a closed system to which a cylinder is attached, shall have a safety relief valve set at a relief pressure that will prevent damage to any part of the equipment. 12. Cylinder valves and regulators shall have outlets and inlets res- pectively for the specific gas as designated in the American Standard of Compressed Gas Association Pamphlet V-1 or for flush-type cylinder valves according to Compressed Gaa Association Pamphlet V-1. 13. Use of adaptors between cylinder valve and regulator should be dis- couraged, but if used, they should be only a type listed in Appendix of Compressed Gas Association Pamphlet V-1. 14. Emergency plans shall be developed to insure control or safe removal of leaking cylinders. Properly maintained protective clothing and equipment for safe entry into an area, contaminated by compressed gases in use, shall be available in the immediate area.
379 Extension Cords in University Buildings (Revision-2) April 1980 Extension Cord Construction 1. The flexible cord shall contain a ground wire and shall be type S, hard usage cord; for cords used with heating appliances, type HS cord is required. 2. Plugs and connectors shall be the grounding type, single-service (multiple outlet cube connectors or adaptors not allowed), allow for inspection of wire connections and have a device to prevent tension transfer to terminal connections. 3. The wire in the flexible cord servicing a current draw of over 7 amps shall be No. 14 or larger wire.
Extension Cord Use and Maintenance 1. Extension cords shall not be used as a substitute for permanent wiring. Multiple strip outlets, fused for wire size of connecting cord are allowed on a temporary basis until permanent wiring is installed.
2. Extension cords shall not be used on fixed or semi-fixed equipment (refrigerators, centrifuges, etc.) or equipment drawing more than 15 amps. 3. Extension cords shall not run through, behind or in walls, ceilings or floors; or run in or through ventilation or other ducts; or suspended over pipes, fixtures or other metal objects. They cannot be used as a substitute for permanent wiring. 4. Extension cords shall not be placed under carpets, under doors, or other locations that subject the cord to wear, abrasion or other damage. 5. Extension cords with broken wires or damaged insulation must be removed from service; splicing or taping is not allowed. 6. Extension cords shall not be used where hazardous atmospheres exist or may exist due to presence of flammable gases or vapors or explosive dusts. 7. Extension cords shall not be placed across aisles or corridor floors or located so as to produce a tripping hazard. 8. The combined length of the appliance cord and extension cord used on very portable equipment such as floor scrubbers, projectors and tools shall not exceed 105 feet. 9. Long cords shall not be left in a coiled or semi-coiled condition when in use.
General Considerations
1. Replacing a very short appliance cord with a longer one is generally more desirable than using an extension cord. 2. Consider the proximity of electrical outlets when locating furniture. 3. Select a cord with proper insulating materials if there will be exposures to moisture, oil or other chemicals. 4. Check frequently for damaged insulation and poor connections at the plug and connector.
380 381 Chemical Label Hazard Signals
Chemicals packaged by Chemical Storehouse and several departments have new color-coded labels to help laboratory and service personnel see at a glance the hazards to health and safety if the chemical is spilled or mishandled.
Fire Hazard 4 Extreme DIOXANE (I .4-DIETHYLENE DIOXIDE) 3 Severe WARNING! Flammable-Vapor harmful 2 Moderate to form explosive peroxides 1 Minor especially when anhydrous 0 None Keep away from heat and open flame. Blue Keep container closed. Health Hazard Use only with adequate ventilation. Avoid prolonged breathing of vapor, or skin 4 Extreme contact. 3 Severe Do nor allow to evaporate to near dryness. 2 Moderate FLASH POINT 12º C. Addition of water or appropriate reducing (54º F.) agents will lessen peroxide formation. 1 Minor 0 None Instability Hazard 4 Extreme Readily explosive under normal conditions. 3 Severe Explosive if strongly initiated, heated, or water added. 2 Moderate Normally unstable, or violently reactive with water. 1 Minor Unstable at elevated temperatures, or reacts with water. 0 None Normally stable.
Special Precautions
Transportation and handling of chemicals in hazard grades 3 and 4 require special precautions to prevent breakage, spills, or exposure to fire. Ventilation and fire-protected storage are required for safe use.
Spills of grade 3 or 4 chemicals require prompt action. Keep people out of the danger area and take action to neutralize, absorb, emulsify or ventilate the spill. Prevent fire by keeping flames, sparks and other ignition sources away from flammable vapors and gases.
Flash points are shown for flammable chemicals--flash point is the lowest temperature at which the chemical will give off enough vapor for ignition to cause a flash fire or explosion.
Chemicals which are highly flammable or only slightly soluble in water are marked "Do Not Dispose of in Building Drains." Such chemicals should be disposed of by calling Plant Services at 273-3625 for free weekly pickup.
Departments that are interested in applying this hazard labeling system to reagent and other chemicals may purchase Pamphlet 325M of the National Fire Protection Association or contact the University Health Service, Division of Environmental Health and Safety.
382 Chapter 15
Plant Growth Responses to a Resource Gradient
Mary Lynn Cowan
Department of Ecology and Behavioral Biology Univesity of Minnesota 109 Zoology Building 318 Church St. S. E. Minneapolis, MN 55455
Mary Lynn Cowan received her B.S. in environmental science from the University of Wisconsin-Green Bay (1979) and her M.S. in plant ecology from the University of Minnesota (1986), where she studied old-field succession. She has worked as a naturalist and environmental educator throughout the Midwest, and is currently working on a Ph.D. in science education at the University of Minnesota. Her current research interests are in the areas of elementary science and science learning at informal education centers. She plans to conduct her dissertation research in West Germany.
383
PLANT GROWTH RESPONSES TO A RESOURCE GRADIENT Mary Lynn Cowan, Dept. of Ecology and Behavioral Biology, University of Minnesota, 318 Church St. S.E., Mpls, MN 55455
OBJECTIVES
1. To demonstrate and measure the growth responses of plants grown along a nitrogen gradient and/or light gradient.
2. To compare the growth responses of different plant species when grown along a nitrogen gradient and/or light gradient.
INTRODUCTION
Nitrogen is an essential mineral nutrient for plants. It is part of many plant molecules, including proteins, chlorophyll, and nucleic acids. Nitrogen deficiency is the most common mineral deficiency in plants. Two forms of nitrogen, nitrate (NO3-) and ammonium (NH4+), are absorbed by plants from the soil. Nitrogen-deficient plants generally exhibit chlorosis (deficiency of chlorophyll) and their leaves turn yellow.
Nitrogen in gaseous form (N2) composes 78 percent by volume of the atmosphere. N2 moves in and out of the leaves through the stomates, but plants cannot use nitrogen in this form. Some organisms are able to "fix" atmospheric nitrogen (N2): they are able to take nitrogen from the air in the form of N2 and reduce it to NH4+ (ammonium), which is a form of nitrogen that can be used by plants. These specialized organisms include some species of soil bacteria, cyanobacteria, and bacteria that live symbiotically with the roots of higher plants. Many plants in the legume or pea family (Fabaceae) have root nodules where the symbiotic bacteria live.
Succession is a gradual and orderly process of ecosystem development brought about by changes in species populations: it is the replacement of populations in a habitat through a regular progression. It often culminates in a "climax community" that is characteristic of a particular geographic region. Succession occurs when a new habitat is created as a result of some process. Primary and secondary succession are the two types of succession. Primary succession occurs in a newly formed habitat, such as an area covered by volcanic ash or lava, an area exposed as a glacier recedes, or a sand dune that is exposed as the water level in a lake deceases. In secondary succession a new habitat is formed, but the area was previously covered by plant growth. Examples of secondary succession include abandoned farm fields and clear-cut forests. Generally, in primary succession there is little or no soil and, therefore, very low availability of soil nutrients. In secondary succession, the soil nutrient availibility varies from very low to relatively high, depending on the soil type and the use (or abuse) of the land before clearing. Either type of succession follows a general pattern. The earliest plants in the sequences are usually short grasses
385 and forbs. These are followed by taller grasses and forbs, and then by woody plants -- shrubs and often trees. In most of eastern North America, the "climax community" is a forest of some type.
Many theories on the mechanisms of ecological succession have been proposed. One of these theories, the resource competition theory, deals with the effects of plant competition for resources. Competition is the use of a resource by one individual that reduces the availability of that resource to other individuals. All plants require below-ground resources (soil nutrients and water) and above-ground resources (light). Below-ground resources are acquired by the roots, while above- ground resources are acquired primarily by the leaves and stems of a plant. In the resource competition theory of plant succession, there is a trade-off between the acquisition of below- and above-ground resources. When below-ground resources are limited, the plants that are most effective at acquiring these limited resources will out-compete the others. Large roots or an expansive root system are two ways that plants can maximize their uptake of limited soil resources. When the density of plants in an area increases, the above-ground resource, light, becomes limiting. Those plants which allocate more biomass to above-ground structures will then be better competitors for the limited amount of light.
In many plant communities, low availability of nitrogen limits plant growth. This is especially true in early stages of primary (and sometimes secondary) succession. Some of the non- legumes that have root. nodules for N2 fixation are pioneer species that are found in early successional sites where the soil is very low in nitrogen. Therefore, they are better competitors for nitrogen because the bacteria in their root nodules fix N 2 from the atmosphere, and possibly because they are also able to absorb the small amounts of NH4+ and NO3- in the soil.
As the early successional field gets "older", soil nutrients increase as decaying vegetation, bacteria, and other organisms add nitrogen to the soil. When the density of plants increases, the availibility of light to individual plants deceases. The resource competition theory of plant succession states that information on nutrient-limited growth rates should make it possible to predict the outcome of interspecific Competition. This theory predicts that plants dominant in younger, more nitrogen-poor fields should have a lower requirement for nitrogen, i.e., a greater ability to grow at low nitrogen levels, than plants dominant in later-successional, more nitrogen-rich fields. The plants growing in the later successional fields are better competitors for the above-ground limiting resource, light. In the later-successional fields, the soils are nitrogen-rich and the plants are not inhibited by low availibility of nitrogen in the soil. But the plants are now more crowded and shade each other. Therefore, the dominant plants will be better light competitors, i.e., those that can grow in the shade of others, and attain a greater height at maturity. A study of several
386 plants dominant along the successional gradient should reveal the following characteristics: early-successional plants - good nitrogen competitors, poor light competitors; mid-successional plants - average nitrogen and light competitors; later- successional plants - poor nitrogen competitors, good light competitors. (See Figure 1.)
Nitrogen Light
Figure 1 - Relative Growth Rate as a function of nitrogen (1a) and light (1b) where species A is an early successional plant and species B is a late successional plant.
In this experiment, you will be studying the growth responses of two plants grown at seven different nitrogen levels. Growth characteristics of the better nitrogen competitors, i.e., those dominant in early succession, should include: (1) higher relative growth rates (RGR), (2) higher root:shoot ratios, (3) shorter height at maturity, and (4) earlier reproduction. Because of the length of the experiment, you will probably not be able to make comparisons using the last two characteristics (height and time of first reproduction).
Relative growth rates are used to compare the growth of two or more species during a specific time period. Relative growth rates can be calculated using the following equation:
RGR = 1n (W4/ W1)
t4 - t1 where W4 = total plant weight at final harvest; W1 = seed weight; t4 = day of final harvest: and t1 = day of planting. You may also wish to determine RGR's during other periods of growth, e.g., between day 1 (seed weight) and day 14, between day 14 and day 70, etc. The early-successional plants should exhibit higher relative growth rates than the late-successional plants because they are able to grow more quickly under conditions of low nitrogen.
Root:shoot ratios can be calculated by dividing the total below-ground biomass per pot by the total above-ground biomass per pot. Because nitrogen is a resource acquired through the roots, plants adapted to low nitrogen soils should maximize their ability to absorb whatever nitrogen is available by allocating more of their biomass to roots, i.e., by having higher root:shoot ratios.
387 MATERIALS
1. Soils - In the nitrogen gradient experiment, you will use seven soil mixtures that have been pre-mixed by the instructor or lab assistant. Each soil mixture is clearly labeled. Nutrients other than nitrogen have been added. In the light gradient experiment, a soil rich in nutrients is used. Additional fertilizers have been added if necessary.
2. Pots and trays - Each group will need 56 pots and trays.
3. Coarse gravel.
4. Seeds - The instructor will tell you the amount (in weight or volume) of seeds you need to use. This will vary from species to species and from year to year because of variations in viability. You will also need exactly 100 seeds of each species to determine the weight of an individual seed. (See step 3 under Methods.)
5. Soil moisture meter - A soil moisture meter measures the percentage of water in the soil. 6. Materials for data collection - These include: data sheets, ruler, paper bags, markers,
7. Harvesting equipment - This includes: sink or basin with running water (preferably warm), sieve, basin to collect soil (not necessary if harvested outdoors), paper bags, scissors, drying oven, balance
METHODS (The schedule below may be altered by your instructor.)
Day 1 -
1. Label each pot with nitrogen level, species, replicate. E.g., N1 - A - 1; N1 - A - 2; etc., where Nl = lowest nitrogen level, A = species A, and 1 (or 2) = replicate 1 (or 2), etc.
2. Put a layer of coarse gravel in the bottom of each pot. Add the soils. Add sufficient water to the pots so that some water drains out the bottom. Let the soils drain for 1 day.
3. Count out exactly 100 seeds of species A and weigh the seeds. Divide this weight by 100 to get the average weight of one seed of species A. Record this information. Repeat for species B.
Day 2 -
4. Add the pre-determined amount of seed to the top of each pot
388 and cover the seeds with a 5-10 mm layer of coarse sand. In an experiment using a nitrogen gradient, the soil mixtures at the different nitrogen levels may be different colors and, therefore, heat up differently. The uniform color of the sand removes this variability.
Days 3 - 70 -
5. Damping off is a disease of planted seeds or young seedlings caused by fungi and resulting in the death of the plants. To reduce the possibility of damping off, do not thoroughly water again until the seeds have germinated and are at least 1 cm tall. If the soil appears dry, mist the surface.
6. Weed and water as necessary. In this experiment, we want to insure that the plants are not water-limited. The seven soil mixtures have different percentages of sand, silt, clay, and organic matter. Since the range of available water varies in soils with different compositions, you must use the soil moisture meter to determine the percentage of water in each pot. Your instructor should have information on the lowest and highest readings required for each soil level. In this experiment, sandy soils may retain water for a longer period. Also, more water is available for use by the plant in sandy soils because the sand particles do not bind to the water as much as clay and silt particles do. (See a soils science text for more information.)
Days 14, 28, 42, and 56 -
7. Collect and record data on the height of individual plants in each pot.
Day 14 -
8. Harvest one replicate of each species at each nitrogen level (a total of 14 pots). Use the following procedure to harvest the plants from one pot: remove the plants from the pot, knock off any loose soil, place the plants in a sieve under a water supply, and wash the soil off of the roots. Separate the above- and below-ground portions of the plants, and place them into labeled bags. Repeat for the other 13 pots.
9. Dry the plants at 50º C until all of the moisture is removed from plants. The amount of time required to dry the plants will depend on the size and type of the plant. Small plants (under 10 cm tall) will only require one or two days of drying.
10. Weigh the plants during the next lab period (Day 21).
Day 42 -
11. Repeat steps 8 - 10. (Harvest one replicate of each species at each nitrogen level.)
Day 70 -
389 12. Repeat steps 8 - 10. (Harvest the remaining plants -- 28 pots total. )
For lab report:
Plot any or all of the following: growth rate as a function of nitrogen level; height as a function of nitrogen or light level; weight as a function of nitrogen or light level; root:shoot ratio as a function of nitrogen or light level.
QUESTIONS
1. Define the following terms: a. succession (including the difference between primary and secondary succession) b. competition
2. Which of your species had the higher relative growth rate at the lowest nitrogen level? at the highest nitrogen level? Is this what you expected? Why or why not?
3. How do the root:shoot ratios of the two species compare?
4. Give some general growth characteristics of plants that you would expect to find in: (a) a farm field that has recently been abandoned, (b) a 50-year old tall grass prairie, (c) the area within a 1-mile radius of Mt. St. Helens today.
REFERENCES
Begon, M., J.L. Harper, C.R. Townsend. 1986. Ecology -- Individuals, Populations, and Communities. Sinauer Associates, Sunderland, Massachusetts.
Tilman, D. 1982. Resource Competition and Community -Structure.--- Princeton University Press, Princeton, N.J.
Tilman, D. 1986. Nitrogen-limited growth in plants from different successional stages. Ecology 67:555-563.
Cowan, M.L. 1986. Growth responses of old-field plants to a nitrogen gradient. M.S. thesis, University of Minnesota.
390 INSTRUCTOR'S PREPARATION MANUAL
INTRODUCTION
This lab, as written, is designed for an introductory or advanced ecology course, but it may be appropriate for an introductory plant science course, or a plant physiology or agronomy course. The biggest drawback for its use in an introductory biology course is the fact that the experiment takes at least 10 weeks to complete, and this would be a lot of time devoted to a plant- and/or ecology-related topic in an introductory biology course. Also, some preliminary information on plants and basic ecological principles, while not necessary, would be helpful.
This lab is written as a greenhouse or growth chamber exercise. If you teach a summer course and have the appropriate outdoor space, you may wish to expand this experiment by using larger pots and conducting the experiment outdoors.
MATERIALS
1. Soils - For the soil nutrient gradient experiment, you will need 2 soils (one low and one high in nitrogen) or 3 soils (one each of low, medium, and high nitrogen availability). These soils may be: 1) topsoil from a nearby field - the quality of this can vary greatly; 2) a black loam or potting soil which you purchase - this will probably have a relatively high nitrogen content; 3) coarse beach sand - this will probably be very low in nitrogen. (See below for soil analysis and construction of nitrogen gradient.) For the light gradient experiment, it is best to use a soil with a high nutrient content to ensure that the plants are not limited by any resource absorbed through the roots.
2. Fertilizer - In the nitrogen gradient experiment, prior to planting, add sufficient amounts of all macro- and micronutrients, except nitrogen, to the soils. In the light gradient experiment, prior to planting, add sufficient amounts of all nutrients that may be limiting based on the soil analysis (see below).
3. Pots - In the greenhouse or growth chamber, use small plastic pots (7 to 15 cm in diameter) and grow one to several plants per pot. Small plastic trays, such as meat trays, should be placed under each pot to collect water that seeps through the pot after watering. This is to insure that there is no transfer of nutrients from a pot of high to low availability. Instead of round or square plastic pots, you can use small,
391 cone-shaped containers (cone-tainers). These are suitable if you are planning to grow one plant per container. You can buy holders for the cone-tainers. Cone-tainers are available from some greenhouse and nursery supply centers, or you can order them directly from the distributor (cone-tainer Nursery, 1500 North Maple, Canby, Oregon 97013, 503-266-3333) If the experiment is to be done in the field, large plastic pots (Zarn pots or poly-pots) should be used. The larger pots will allow more space for the rapid root growth. Pots that are 30 cm in diameter and 30 cm deep weigh approximately 23 kg (50 #) when filled with soil. Use smaller pots if you wish to harvest plants before the end of the growing season. All of the above pots are available through garden supply centers.
4. Coarse gravel or two layers of cheesecloth to line the pot, to allow for drainage and to prevent soil loss.
5. Soil moisture meter - (available from Forestry Supplies) A soil moisture meter measures the percentage of water in the soil. The range of available water varies in soils with different proportions of clay, silt, sand, and organic matter. When conducting a nutrient gradient experiment, make sure you have a soil moisture meter that can be calibrated so that you can adjust it to measure the percentages of available water in all of your soil mixtures.
6. Seeds - You will need at least two species of herbaceous plants for this experiment. It may be interesting to have each group of students use a different set of two species. If possible, use species commonly found in your area. The strongest interspecific differences will be seen if you choose species which a) have very different growth forms (e.g., rosettes v. grasses), or b) are dominant during different stages of ecological succession ("weedy" colonizers v. "climax" species). Since the experiment will be relatively short if it is part of a course taught during the regular school year, you may wish to use annual species. However, perennial species can be used successfully if you refrain from using those that produce tubers or large tap roots during their early growth. (See below for information on seed collection and viability.)
7. Materials for data collection - These include: data sheets, ruler, paper bags, markers,
8. Harvesting equipment - This includes: sink or basin with running water (preferably warm), sieve, basin to collect soil (not necessary if harvested outdoors), paper bags, scissors, drying oven, balance
392 PRELIMINARY ANALYSES AND PROCEDURES
A. Soil -analysis---- - Collect 10 small samples (20-30 ml) of each soil type that you plan to use. Mix all the samples from one soil type together in a paper bag. Label the bag and send it to a soils lab for analysis. Soils analysis can be done as part of class project in a soils class, or by the soils department at your school, or by a soil analysis service run by your state. Have the soils tested for percentage of organic matter and all macronutrients (N, P, K, Mg, Ca, S). It is probably not necessary to have analysis done on the micronutrients unless you have reason to believe that one of these may be lacking. (A soil scientist in your area can advise you in this matter.) Avoid soils with a high percentage of organic matter (> 20%) because these soils will decompose relatively quickly and release more nutrients, including N. This will cause complications in your analyses because a soil mixture with more of the soil that is high in organic matter may have a different total nitrogen value than originally intended. (Note: Regular potting soil that you buy for use at home has a high percentage of peat moss, and therefore, is high in organic matter. Greenhouse supply personnel can assist you in finding soil that has a lower organic material content.)
B. Calibrating the soil -moisture----- meter- -
A few soil science definitions may be useful before explaining this next section. When a soil is at "field capacity", it essentially contains the maximum amount of water that can be held in its pore spaces. When the soil moisture content is so low that a plant remains wilted both night and day, the soil has reached the stage that is called the "wilting coefficient". "Available water" is the range of soil moisture conditions where there the soil moisture content is above the wilting coefficient and below the field capacity. Because of the variation in water holding abilities in different soils, you will need to calibrate the meter so that your meter readings cover the range of available water in all of the soils. For example, the range of "available water" in a typical sandy soil is 3 to 9 per cent by weight; in a sandy loam the range is 7 to 15 per cent. See a soil science textbook for more information on water availability in different soils. (Brady, N.C. 1974. The Nature and Properties of -Soils,-- 8th Edition. MacMillan Publishing Co., Inc., New York, NY. )
Once you determine the range of available water for each of your soil mixtures (they will probably be fairly close), you will need to calibrate the soil moisture meter. A soil moisture meter will give you a reading from 0 to 10 with the low value relating to little or no water in the soil. The procedure for determining
393 the percentages of water per dry weight soil in each of your soils follows:
1. Take a small sample (30-50 ml) of each soil. 2. Soak the soils until you think they have reached field capacity. Set aside for approximately one hour. 3. Take a meter reading. Record. 4. Weigh the sample. Record. (Make sure you have weighed the container beforehand so you can subtract that weight in your calculations.) 5. Place the containers in a drying oven for 30 - 60 minutes. 6. Remove the containers from the drying oven, then repeat steps 3 - 5. 7. Keep repeating steps 3 - 6 until the soils are very dry. (This should take about 3-4 hours.) 8. Using the last reading for dry weight, calculate the percentage of water in each soil sample for each reading. Match this percentage with the reading on the meter and determine the proper meter readings for the range of available water for each soil.
C. Constructing a nitrogen -gradient-- - Materials: soils, scale, cement surface for mixing, shovels, large heavy-duty plastic bags or other containers for storing soil mixtures
Procedure: Determine the range of nitrogen availability that you will use. Your highest value should be at least as high as the richest soils in your area. Determine the number of nitrogen levels you will use. The weight and height of plants should be expected to increase along the nitrogen gradient, but eventually the response will level off. The shape of the response will be a Monod growth curve (or Michaelis-Menten curve). To get adequate data for statistical analysis of the results, you should use at least 6 nitrogen levels. Using the soil nitrogen analysis data, mix soils in different proportions to obtain sufficient soil for each nitrogen level. If this experiment is done in a greenhouse or growth chamber, you will need relatively small amounts of soil for each nitrogen level. To reduce variability, mix a sufficient amount of soil for each nutrient level at one time. Under most circumstances, the mixing can be done on a cement surface (greenhouse floor or parking lot) using shovels. Add fertilizers at this time. The example below shows the percentages (by weight) of three soils in each soil mixture in a gradient using seven nitrogen levels (Cowan, 1986).
394 NITROGEN LEVEL BLACK LOAM TOP SOIL SAND (5000 ppm N) (350 ppm N) (25 ppm N) 1 - 125 ppm 2 - 250 ppm
3 - 375 ppm 4 - 575 ppm 5 - 825 ppm 6 - 1275 ppm 7 - 1825 ppm
D. Seed --Collection-- - Seeds may be collected at a local field site or purchased from a nursery supply center. Throughout the Midwest, prairie grasses and forbs can be purchased from nurseries that specialize in native species. While purchased seed requires much less work on your part, if you wish to make predictions about responses of plants at your field site, it is best to collect seed from your site. In the upper Midwest, mature seeds can be collected from most species in August and September. Some species, such as the "cool season" grasses, have mature seeds earlier in the season. Collect the seeds by stripping them from the stem or cutting off the entire flowering stalk. Store in a cool, dry place in paper bags. Some seeds need to be stratified (stored at cold temperatures to simulate winter conditions). These seeds can be stored in a refrigerator. Some seeds have an impermeable seed coat and need to be scarified. This can be accomplished by placing the seeds in a solution of sulfuric acid for 1-5 minutes, or by rubbing the seeds on sandpaper. Consult an agronomist or botanist about the correct procedure for the particular species you are using. Some plant labs have equipment for scarifying seeds and for cleaning seeds (removing stems and other extraneous plant parts).
E. Seed -Viability--- - If you purchase seed, it normally has been tested for viability, and this value is given. If you collect your own seed, you should do germination trials to insure sufficient germination during the experiment. First, you should clean the seed by removing stems, etc. Next, count out 50 or 100 seeds, weigh them, and determine their volume. Place these seeds on
395 moist paper toweling in a petri dish and cover them. Observe daily and record the number of seeds that germinate. Do not let the seeds mold or dry out. Some seeds will germinate in 3-5 days, others will take several weeks. It may be best to use species that germinate in approximately the same amount of time when conducting a short-term experiment. Decide how many seeds you want to grow in each pot. Determine the amount of seed (by weight or volume) you will need to use to achieve this density. If you are only planning to grow one plant per pot, it is best to plant extra seeds and thin the plants after germination.
F. Setting up a light gradient -
Determine the range of light availability using a light meter and decide the number of light levels you will use. If the experiment is conducted in a greenhouse or growth chamber, determine which area receives the greatest amount of light throughout the day. Use this area as the highest light level. To make lower light levels, use cheesecloth, shading material available through garden centers, or some other material that can be layered to cut out light. Depending on your set-up, you may want to place the shading material above and/or on the sides within each light level. Allow space between each light level to take into account shading from adjacent levels.
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