LABORATORY SET-UP AND OPERATIONAL …testbankcart.com/wp-content/uploads/2017/01/Downloadable-Solution...Laboratory Set-Up and Operational Suggestions 9 ... and the nature of scientific

Laboratory Set-Up and Operational Suggestions 9

LABORATORY SET-UP AND OPERATIONAL SUGGESTIONS

These suggestions for laboratory preparation and operation are provided to save you time and to help your labs runmore smoothly. Generally, only those portions of the labs that merit special attention are discussed.

PART I FUNDAMENTALS

Exercise 1. Orientation

This initial exercise is designed to help students “get off on the right foot” in the laboratory. It emphasizes how stu-dents should prepare for a laboratory session and how they can work most effectively in the laboratory. It is advisableto verbally stress laboratory safety and housekeeping. Activities are designed to help students (1) understand biologicalterms, (2) make simple measurements and calculations using the metric system, and (3) make careful observations ofbiological specimens.

The first laboratory session is the best time for you to emphasize your own preferences for (1) how students shouldprepare for the lab session and (2) behaviors that enhance success in the laboratory. Stressing these points now will saveyou time later in the course. You should emphasize what you expect from students and what they can expect from you.One of the major hurdles in teaching students to prepare an effective laboratory is to convince students to come to thelab prepared for each lab session. You will need an effective strategy to establish this behavior.

The exercise emphasizes (1) structure and meaning of biological terms, (2) use of the metric system, (3) makingcareful observations, and the nature of scientific inquiry with an emphasis on the scientific method. Some students mayneed help in making the metric system conversions, so you may wish to work an example or two on the board. Also, stu-dents may need to be assured that there is no “one correct way” to make the dichotomous key for the leaves.

Students are guided step by step through the scientific method, which may seem rather laborious at first. After theyunderstand and can use the process, there is no need for them to write down the mental steps, as long as they are“thought through.”

Exercise 2. The Microscope

This exercise is designed to help students develop basic microscopy skills. Most students think that they know howto use a microscope, but few really do. It is worth the effort to check out the skills of each student during the lab becauseit will save time later on for both you and the students.

If your microscopes are different than those described, identify the differences to your students. If the high-powerobjectives of your microscopes are 43x or 45x, students will need to take this into consideration when calculating the(1) total magnification and (2) diameter of field. They also need to recognize that when the manual requests observa-tions at total magnifications of 400x, this means 430x or 450x for them.

Care of the Microscope

Emphasize the care of the microscope that you want students to follow if it is different from that described in thelaboratory manual.

Focusing

Students need to be reminded (1) to always start focusing with a low-power objective, usually the 10x objective,and then to switch to the high-power objective if it is to be used, rather than starting with the high-power objective and(2) to use only the fine-focusing knob when using the high-power objective.

Depth of Field

It is a good idea to emphasize that prepared slides (1) are to be handled by the label or edges to avoid fingerprintson the cover glass and (2) are to be cleaned with soap, water, and lens paper.

Some students have difficulty with the concept of depth of field and in determining the three-dimensional shape ofobjects from the two-dimensional images observed. Observations of fly-wing spines are a big help for such students.Their understanding of depth of field and their focusing skills will be “tested” later in the exercise when observing theslide of crossed human hairs. How easily they determine which hair is uppermost identifies their understanding of depthof field. A little practice here saves time later.

An explanation of “optical midsection” is usually required when students are trying to focus through the depth ofthe blond hair.

10 Laboratory Set-Up and Operational Suggestions

PART II CELL BIOLOGY

Exercise 3. The Cell

The background information presents (1) the distinguishing characteristics of prokaryotic and eukaryotic cells and(2) the similarities and differences in the ultrastructure of plant and animal cells. Students need to be reminded that theycan only see the very large organelles with their microscopes. Labeling the cell diagrams helps students learn cellularcomponents.

Moneran Cells

The major point of students’ study of moneran cells is to observe their small size and the inability to see cellularcomponents in contrast to what they will observe in eukaryotic cells later in the exercise. Selected slides may be set upas a demonstration, if you wish.

Mixture of Prokaryotic and Eukaryotic Cells

The purpose here is to challenge students to distinguish bacterial and yeast cells in a mixture. The mixture shouldbe rather concentrated, so that cells of each type will be in each drop sampled by students. Prepare the mixture by mix-ing a half packet of baker’s yeast in 50 ml of 1% glucose solution. After an hour, add a broth culture of Bacillus subtilisor a similar bacterium. Mix by stirring.

Onion Epidermal Cells

Cut a few red onion scales into small strips about 1–2 cm long and place them in water in a fingerbowl for easydispensing. These smaller pieces make it easier for students to remove the epidermis. The only difficulty students mayencounter is removing too many cell layers when removing the red epidermis.

Elodea Cells

Students may need to be reminded to select a light green portion of the Elodea leaf for study of the cell structuresince too many chloroplasts make the finding of a nucleus more difficult. Some may need help in distinguishing surfaceand optical midsection views of an Elodea cell and focusing through the depth of a leaf to locate the two cell layerscomposing the leaf. Students typically have trouble ignoring portions of the image that are not in focus and sometimesoverlook the three-dimensional shape of cells.

Human Epithelial Cells

Toothpicks used to scrape epithelial cells from the lining of the mouth and the glass slides and cover glasses used withhuman epithelial cells should be placed in biohazard bags and autoclaved prior to discarding as an infection-control measure.If you wish to save the slides, have students place them in a separate biohazard bag; then they can be autoclaved before wash-ing. Or, if you prefer, the slides may be placed in a disinfectant solution of Amphyl (0.5%) or household bleach (10%) for atleast 30 min instead of autoclaving. The lab table should be washed with Amphyl or bleach at the end of the lab session.

Exercise 4. Chemical Aspects

This exercise enables a basic understanding of chemistry used in the exercises that follow.

Acids, Bases, and pH

You may wish to include sample solutions of your choice for pH determination.In Assignment 3, steps 3–7, it is important to adjust the pH of distilled water to pH 7.0 with 1.0% NaOH, so that both

the distilled water and buffer solution have the same starting pH for the experiment. Distilled water exposed to atmospher-ic for a time will have a pH less than 7. This experiment works well to impress students about the function of buffers.

Identifying Biological Molecules

You may wish to include materials of your choice to test for the presence of carbohydrates, fats, and proteins. Gen-erally, students have no problem with this section once they understand the test procedures.

Starch Preparation

Prepare the 0.1% soluble starch solution by mixing the starch into distilled water. Heat the mixture, but not to boil-ing, while stirring until the starch goes into solution. Store the starch solution in a stoppered flask in the refrigerator

CO2


until needed. Allow time for it to return to room temperature before use. Test a sample of the starch solution before thelaboratory to be sure that it gives a positive reaction with iodine solution.

Benedict’s Solution

Place Benedict’s solution in light-proof dropping bottles to prevent deterioration. It is best to use a fresh supply foreach exercise where it is used.

Exercise 5. Diffusion and Osmosis

This exercise requires that students utilize their time well. If time is limited, the observation of Brownian move-ment and the experiment on diffusion and molecular weight may easily be set up as demonstrations. A brief review ofthe scientific method may be useful since it is emphasized in the experiments.

This exercise uses “drops” and “droppers” in dispensing solutions. If you prefer to have students dispense solu-tions with a pipette, 1 ml equals 1 dropper.

Brownian Movement

Students need to be cautioned not to place too much carmine powder on the slide. Only a few barely visible parti-cles are needed.

Prepare the water–detergent solution by adding 3 drops of liquid detergent to 100 ml water.

Starch Preparation

Prepare the 0.1% soluble starch solution by mixing the starch in distilled water. Heat the mixture, but not to boil-ing, while stirring until the starch goes into solution. Store the starch solution in a stoppered flask in the refrigeratoruntil needed, but allow time for it to return to room temperature before use. It is important to test a sample of the starchsolution before the laboratory session to be sure that it gives a positive reaction with iodine solution.

Osmosis

In setting up the osmometer, place the 2-hole rubber stoppers on the glass tubing in an inverted position, that is, in-sert the glass tubing into the small end of the stoppers. The cellulose sacs can then be tied on with string so that they willnot slip off the end of the stoppers. Fill the sacs with the medicine droppers through the other hole of the stopper. Whenfilled, insert the dropper snugly into the hole to prevent leakage, and rinse the exterior of the sac to remove any spilledsucrose solution.

The osmometer will last an entire week if the water in the beakers is changed daily and the cellulose sacs are ele-vated out of the water when not in use.

In the section on Osmosis and Living Cells, the experiment with celery sticks requires at least an hour for good re-sults. Students need to plan on this, or you can set it up as a demonstration prior to the lab session. In the experiment onplasmolysis in Elodea cells, it is wise to remind students to rotate the high-power objective out of the way when addingthe salt solution to the slide at the edge of the cover glass. This will minimize the chance of getting salt solution on theobjectives and stage. However, after completion of the observations, the objectives and stage should be washed anddried in case any salt solution got on them.

Exercise 6. Enzymes

This experiment acquaints students with the nature of catalase, an enzyme present in cells of all aerobic organisms.Generally, they have no problems with the experiments. The key to success is (1) following the directions and (2) clean-liness to prevent unwanted contamination by enzyme extracts. Washing the mortar and pestle after each extract prepa-ration and the test tubes after each experiment is crucial.

This exercise uses “drops” and “droppers” in dispensing solutions. If you prefer to have students dispense solu-tions with a pipette, 1 ml equals 1 dropper.

In Assignment 4, it is crucial that the enzyme extract and hydrogen peroxide be heated separately at the appropri-ate temperatures for 10 min before the contents are mixed.

Assignment 5 allows students to think through the problem and to design their own experiments to evaluate the ef-fect of pH on catalase activity. The experiment in Assignment 4 provides a suitable model for them. The key here is thatthe extract containing catalase must be exposed to each pH tested for 3–5 min before hydrogen peroxide is added.


Exercise 7. Photosynthesis

This exercise works best if students work in groups of 2–4 and share the duties and data.The fairy primrose (Primula malacoides) is the best plant to use for the initial series of experiments involving

starch tests on leaves. It is usually available from nurseries in the fall in frost-free regions and in the spring in colder cli-mates. They are easily grown from seeds if facilities are available. Tomato plants work fairly well, but their leaves donot keep their shape after boiling.

Caution students about the flammable nature of alcohol. Use hot plates, not Bunsen burners, and a water bath forchlorophyll extraction from the leaves. The use of boiling chips will reduce any splatter of alcohol or water.

Provide students with light-proof dropping bottles of Benedict’s solution and iodine solution

Carbon Dioxide and Photosynthesis

If you have a number of laboratory sections to prepare in sequence, keep a supply of well-watered plants in dark-ness and a similar number exposed to light in a green house or prep room. These may be moved into the laboratory asreplacements for the plants giving negative and positive reactions to the starch test, respectively, as their leaves get de-pleted by students. This procedure avoids the hassle of preparing several experimental setups that test the effect of car-bon dioxide on photosynthesis and moving in an entire new setup for each lab session.

Light and Photosynthesis

If you prefer not to use light shields on leaves in this experiment, just use leaves from (a) plants exposed to lightand (b) plants kept in darkness for 24 h prior to the laboratory session.

Chlorophyll and Photosynthesis

Small Coleus plants with green and white variegated leaves are best for this experiment. No other plant seems towork as well. They are easily grown from cuttings. If you can’t grow your own, they may be obtained from biologicalsupply houses year around and from local nurseries in the spring.

Plant the green and albino corn seed two weeks prior to the laboratory session in order to have seedlings for the lab.

Chloroplast Pigments

Students enjoy the separation of chloroplast pigments by paper chromatography. Caution students about thevolatile and flammable nature of the chloroplast and developing solvent solutions. These materials must not be nearopen flames. It is best to place stoppered Erlenmeyer flasks of both the chloroplast pigment solution and the developingsolvent in the fume hood where students can obtain these liquids. The dispersal of these liquids to students and theirsubsequent collection in a waste jar should be done in a fume hood. If a fume hood is not available, use a well-ventilat-ed portion of the room away from open flames.

Chloroplast Pigment Solution

Prepare the chloroplast pigment solution under a fume hood as follows:

1. Shred several fresh or frozen spinach leaves with scissors and place them in a blender containing 200 ml acetone.

2. Blend at high speed for 2 min.

3. Filter through cheesecloth, collecting the filtrate. Discard the residue.

4. Filter the filtrate again using fast-filtering filter paper.

5. Evaporate the filtrate under a fume hood to yield 50 ml of the concentrated solution.

6. Store in a stoppered flask in the refrigerator until used.

Developing Solvent

The developing solvent consists of 22 parts petroleum ether to 3 parts acetone. To make 1000 ml, use 880 ml of pe-troleum ether and 120 ml of acetone. This mixture gives good chromatographic results.

Light Quantity and the Rate of Photosynthesis

It is important that students read and understand the concept and procedures of this experiment before starting it.They must also be able to read the fluid level in the inverted pipette, so they should understand the graduations on thepipette before starting the experiment.

1I2 + KI2.


If you are using a spot lamp, use either a 300-W or 150-W spot bulb. A 300-W will give faster results but increas-es the chance that a student may get injured by touching a very hot bulb. Using a 150-W bulb or a fluorescent lamp mayrequire shorter light-to-plant distances. The use of a fluorescent lamp eliminates the need for a heat filter.

The most likely source of difficulty is getting a good production of oxygen bubbles from the cut end of the Elodeashoot. The diagonal cut of the stalk is best made with a single-edged razor blade. After the shoot is submerged in waterin an inverted position in the test tube, it must be checked for good bubble production before continuing. Often severalcuts must be made to get a good bubble production. If bubble production is limited, the time at each light quantity maybe increased or the distance from the light source may be decreased to compensate for it.

The Color of Light and the Rate of Photosynthesis

This section provides an opportunity for students to design and conduct an experiment to assess this problem.Students should use a short distance between light source and Elodea shoot to get a good bubble production. Coloredlight filters are available from biological supply houses. It is best to place the filter next to the Elodea shoot to preventunwanted light contamination from other sources. Never place the filter next to the spot lamp since it may catch fire.

Exercise 8. Cellular Respiration and Fermentation

This exercise requires good preparation and proper utilization of time by students. They should work in groups of2–4 and share the duties and data.

Dilution of stock bromthymol blue to yield a 0.004% solution is necessary to increase its sensitivity so that it willyield rapid color changes in the experiments.

Cellular Respiration and Carbon Dioxide Production

Germinating pea seeds should be 3–6 days old. Crickets work well because they are easily kept out of the bromthy-mol blue solution by the segments of glass tubing.

Students should set up the experiment in Assignment 3 while waiting for Experiment 2, in Assignment 2, to devel-op. It is important to place the upper surface of the leaf against the test tube since most stomata are on the lower surfaceof the leaf. A fluorescent lamp is best for illumination. If a laboratory lamp is used, a heat filter is needed to prevent ex-posing the leaves to excessive heat.

Cellular Respiration and Heat Production

Set up the vacuum bottles at least 12 h prior to the laboratory session to get good results on heat production. A wadof cotton may be used as a stopper to support a thermometer instead of a cork stopper. This allows gas exchange andstill retains sufficient heat to obtain the desired results.

Rate of Cellular Respiration

The respirometer for this experiment is made from a test tube, a 1-hole rubber stopper, and a 1-ml pipette. Thepipettes should be bent and inserted into the 1-hole stoppers for each student group before the lab. Make the 90º bendbetween the 0.0 ml mark and the base of the pipette. If the pipette is plastic, very little heat is required to bend it, andplastic, disposable pipettes work well. Insert the base of the pipette into the 1-hole rubber stopper so that it is flush withthe inner surface of the stopper.

It is crucial that the inside of the test tube be completely dry prior to adding the soda lime. Students must know howto read the graduations on the pipette before setting up the experiment. They may also need assistance in the calculations.

This experiment may be extended to determine the effect of temperature on the rate of cellular respiration by deter-mining the rate of cellular respiration at 40° C and 10° C, in addition to room temperature, if you are not doing sections6 and 8. A beaker of water at the desired temperature is a suitable water bath.

Temperature and the Rate of Cellular Respiration

The experiment using vertebrates requires a respirometer similar to that shown in Figure 8.5. If you wish, smalldessicators may be used as the chambers instead of wide-mouthed jars, but obtaining a good seal requires petroleumjelly and a bit of care. Good technique is required in manipulating the respirometer and reading the manometer. A fewprocedures that will increase the success rate are as follows:

1. Ample time must be provided for temperature equilibration before starting the experiment.

2. The vent tubes must not be closed until after the stoppers are snugly inserted into the jars. Failure to do so may in-jure the test animals. It is essential to double-back the ends of the vent tubes, before securing the tube clamps, toassure closure.


3. Precaution must be taken (i.e. use of ample cotton under the screen and a little luck) to prevent the soda lime fromgetting wet by urine. If urine (or water) contacts the soda lime, pressure will build up in the experimental chamber.The only solution is to replace the soda lime and start over.

Fermentation in Yeast

Prepare the yeast suspension by adding one packet of granulated baker’s yeast to 100 ml of a 1% glucose solution.Keep at room temperature for about an hour before the laboratory session.

Prepare the soluble starch solution by mixing the starch in distilled water. Heat the mixture, but not to boiling,while stirring until the starch goes into solution. Store in a stoppered flask. All solutions should be at room temperatureprior to use.

You may need to demonstrate the use of a pipette pump and how to clamp the tubing before removing the pipettepump. The plastic tubing must be long enough to fold and clamp off with ease.

Independent Inquiry

Set the parameters for the student investigations, and provide the materials that you wish students to use.

Exercise 9. Cell Division

Students have no problems with this exercise. Many of the background questions can be answered prior to the lab-oratory session so that lab time is devoted to observations. It is helpful to remind students that they may need to exam-ine several slides in order to find each of the mitotic phases.

Use of chromosome simulation kits or pop beads enhances students’ understanding of the differences between mi-tosis and meiosis.

PART III DIVERSITY OF ORGANISMS

The next seven exercises expose students to the diversity of organisms. If you use a classification system that is dif-ferent than the one presented here, inform students of the differences as they work through these exercises. It is also agood time to point out the dynamic nature of taxonomy and systematics.

Color plates (inserted in this section) of selected organisms provide structural details that are often difficult for stu-dents to observe in the laboratory.

Life cycles and reproductive patterns are included in these diversity exercises since they are important distinctivefeatures of each organismic group.

The laboratory report for each exercise on diversity contains a section for the identification of “unknown organ-isms”—a sort of mini-lab practicum. The use of “unknowns” is a very effective way to stimulate student learning, andit provides an easy way for both instructor and students to evaluate students’ understanding of the key recognitioncharacteristics of each organismic group.

Exercise 10. Prokaryotes and Protists

Take a few moments to discuss the dynamic nature of taxonomy and the rationale for the use of the three domains.If a microscope video system is available, it may be used very effectively in this exercise to augment students’ observa-tions using student microscopes.

Bacteria

Bacteria are always smaller than what students imagine. Remind students to use reduced illumination for betterviewing. Refer students to Plates 1.1 to 1.3.

If a microscope video system is not available, set up the demonstration slides under oil immersion (1000x). It is help-ful to set up the living flagellated bacteria using a hanging-drop slide and either dark-field or phase-contrast illumination.

Demonstration slides of living Gloeocapsa and Oscillatoria set up under oil immersion objectives of demonstra-tion microscopes help students to see cellular structure and the absence of a nucleus. Use of a microscope video systemis very effective here. Refer students to Plate 1.4.

Preparation of Agar Plates to Test Antibiotics

1. Prepare a flask of brain–heart infusion agar.

2. Pour about 10–15 ml of the agar into each Petri dish. Replace the cover and allow time for cooling and solidification.


3. Dip a sterile, cotton-tipped swab into a culture of E. coli B and swab the entire surface of the plate. Use a new swabfor each plate. Replace the lid.

4. Use the same technique to inoculate plates with S. epidermidis.

5. Use flamed forceps or a disc dispenser to add the antibiotic discs to the agar, spacing the discs equally from eachother. (Antibiotic discs may be purchased from Wards, P.O. Box 92912, Rochester, NY 14692-9012.)

6. Press the discs slightly into the agar with flamed forceps. Replace the lid.

7. Incubate the plates in an inverted position for 24 h. After incubation, it is a good idea to tape the Petri dishes closedso students cannot open them.

Protozoa

Plates 2.1 to 3.5 are very helpful to students as they study these organisms microscopically.Be sure to inform students how to obtain Pelomyxa with a medicine dropper from the bottom of the culture jar and

place it on a slide. Without instruction, students usually withdraw too much water into the dropper with the organismsat the top of the water column, so they seldom get the organisms on the slide.

Plasmodium vivax

This sporozoan is difficult for students to see because of its small size even at 1000x. A microscope video systemand Plate 3.5 are helpful.

Plantlike Protists

Refer students to Plates 4.1 to 4.5 when studying these microscopic specimens. It is helpful to set up a slide ofEuglena, with Protoslo mixed in, under a demonstration microscope at 1000x (oil immersion) so students can better ob-serve the cellular structure. Phase-contrast is helpful.

Light and Euglena Distribution

Prepare the demonstration showing the effect of light on the distribution of Euglena as follows:

1. Tape a circle of filter paper to the bottom (outside) of a Petri dish.

2. Attach a semicircle of black construction paper to the lid by a “hinge” of masking tape at the edge of the lid so thatit covers half the lid as a light shield. Place a small piece of masking tape at the front edge (opposite the hinge) ofthe paper that can serve as a tab for lightly attaching the front edge to the lid and for lifting the paper to make ob-servations. The dish may be taped to the top of a counter or table so that it cannot be moved, if you wish.

3. Add a Euglena culture to the Petri plate, and replace the lid.

4. Use ambient light, not a laboratory lamp, for illumination since Euglena avoids bright light. It takes 2–3 h forEuglena to orient to the light.

Fungus-like Protists

Students should compare their observations of amoeboid and sporangia stages of Physarum with Plates 5.1 and 5.2.

Reproduction in Protists

It is essential for students to understand meiosis and Figure 10.7 before starting their observations. Mix the mating typesof Paramecium bursarium at least an hour before the lab session to allow conjugants to form by the time the lab begins.

Exercise 11. Green, Brown, and Red Algae

If a microscope video system is available, it may be used very effectively in this exercise. Plates 7.1 to 7.5 and 8.1to 8.5 also help students interpret their observations.

Green Algae

Setting up a demonstration slide of Chlamydomonas under an oil immersion objective helps students to perceivethe structure of this small green alga. Students never expect it to be so small. Carteria is a similar, larger alga that is asuitable substitute for Chlamydomonas, although it possesses four flagella instead of two.


Light and Chlamydomonas Distribution

Prepare the demonstration showing the effect of light on the distribution of Chlamydomonas as follows:

1. Tape a circle of filter paper to the bottom (outside) of a Petri dish.

2. Attach a semicircle of black construction paper to the lid by a “hinge” of masking tape at the edge of the lid so thatit covers half the lid as a light shield. Place a small piece of masking tape at the front edge (opposite the hinge) ofthe paper that can serve as a tab for lightly attaching the front edge to the lid and for lifting the paper to make ob-servations. The dish may be taped to the top of a counter or table so that it cannot be moved, if you wish.

3. Add a Chlamydomonas culture to the Petri plate and replace the lid.

4. Use ambient light, not a laboratory lamp, for illumination since Chlamydomonas avoids bright light. It takes 2 to 3h for Chlamydomonas to orient to the light.

To determine whether Chlamydomonas prefers bright or dim light, students can prepare a setup similar to the oneabove, but use white typing paper instead of black construction paper. Use a fluorescent lamp at a distance of 3 ft as thelight source. Allow 2 h for orientation of Chlamydomonas.

Slide Preparations

A hanging-drop slide is best for a demonstration setup of Volvox at 40x or 100x to give students a good apprecia-tion of the motility and size of this alga.

Students may need to be reminded to use a very small amount of Spirogyra when preparing the water-mount slide.They tend to use too large a clump, which prevents good observations.

Brown and Red Algae

If a marine aquarium is available, living brown and red algae can be easily demonstrated in it. If not, fresh speci-mens may be obtained from a biological supply house and maintained for a week or so if kept moist with sea water andin the refrigerator, when not in use. Preserved specimens that have been washed and placed in water for observation arealso quite suitable.

Reproduction in Green Algae

Fundamental reproductive patterns, including alternation of generations, are introduced here. Students should un-derstand Figures 11.6 and 11.7 before making their microscopic observations.

Exercise 12. Fungi

The use of a microscope video system and the reference to Plates 5.3 to 6.4 are helpful in this exercise.

Preparation of a Water–Detergent Solution

A water–detergent solution is used in making wet-mount slides of fungi because it aids the separation of hyphae forbetter observations. Prepare it by adding 3 drops liquid detergent to 100 ml water.

Preparation of Rhizopus Culture on Bread

1. Be sure to use bread without preservatives for the culture of Rhizopus.

2. Place a piece of bread in a Petri dish and autoclave it at 100º C for 10 min.

3. Innoculate the bread by transferring spores from a Rhizopus culture tube with a flamed loop.

4. Sprinkle lightly with water.

5. Incubate at room temperature for 4–7 days.

Preparation for Mold Cultures on Agar

1. Pour 10–15 ml of Sabouraud’s agar into a Petri dish and replace the cover. Allow time for cooling and solidification.

2. Innoculate with spores from a culture tube with a flamed loop.

3. Incubate at room temperature for 4–7 days.

Reproduction in Black Bread Mold

Prepare the demonstration of Rhizopus zygospores by using a flamed loop to innoculate a sterile plate ofSabouraud’s agar with spores from and mating types. Streaks of the two mating types should be about 0.5 to0.75 cm apart across the diameter of the plate. Allow about 7 days incubation at room temperature.

-+


Hyphae from Germinating Spores

The structure of hyphae are best observed by germinating spores on slides of Sabouraud’s agar. Prepare the slidesas follows:

1. Place 1–2 dozen sterile slides on clean paper towels.

2. Use a pipette to place a few drops of Sabouraud’s agar on each slide. A thin, even coating is all that is needed.

3. After the agar has cooled, add spores to the agar film. Innoculate half of the slides with Rhizopus and the other halfwith Penicillium. Add a cover glass to each slide.

4. Place the slides vertically in a Coplin jar to maintain humidity and incubate at room temperature for 2–3 days.

5. After the hyphae have grown sufficiently, refrigerate until used.

Wet-Mount Slides

In making the wet-mount slide of Rhizopus, students often use a piece of bread that is too large. They should use apiece about 3 mm across and tease the bread from the hyphae with dissecting needles under a dissecting microscope.After adding the cover glass, apply a little pressure to it with the handle of a dissecting needle, which will flatten out thematerial for better viewing with a compound microscope.

Basidiomycetes

Very thin longitudinal sections of stipe are needed if hyphae are to be clearly observed. It is better to prepare verythin sections of mushroom stipe and place them in water in a Syracuse dish for student use rather than have studentsmake their own sections. Students tend to prepare sections that are too thick.

Exercise 13. Terrestrial Plants

This exercise focuses on recognition characteristics and reproductive patterns and processes in mosses, ferns,conifers, and flowering plants. The anatomical structure of flowering plants is in a separate exercise, Exercise 31. It iscrucial that students understand the basic nature of alternation of generations before starting this exercise.

Instructors are encouraged to use as much fresh and living material as possible, but much of the study comes from pre-pared slides by necessity. Depending on the depth of the coverage you wish, the exercise may require two lab sessions.

Exercise 14. Simple Animals

Major criteria for classifying animals are provided at the start of the exercise to give students an overview of majoranimal characteristics. It is important that students understand gametic sexual reproduction, Figure 14.3, in this exercisesince it is characteristic of all animals.

Sponges

To best view whole and longitudinally sectioned preserved Grantia under a dissecting microscope, advise studentsto use the black surface of the stage plate to provide contrast. The specimens should be in a Syracuse dish and coveredwith water.

A prepared slide of Grantia, l.s. is set up under a demonstration compound microscope because the slides are oftenin rather poor condition, the cells are very small, and students have difficulty in locating the component parts.

Cnidarians

The feeding activity of Hydra intrigues students. It is best demonstrated by placing all of the Hydra in a culture jarfor 25 students in a 1000 ml beaker of pond water. Do not use tap water. Add 1–2 sprigs of Elodea to serve as sites for at-tachment of the Hydra. Prepare this setup 2–3 days before the laboratory session to allow the Hydra to adapt and disbursethemselves throughout the container. Add a few small-sized Daphnia during the laboratory session to demonstrate thefeeding response. Feeding can be stimulated by adding a few drops of 0.01% glutathione solution, but this is usually notnecessary. Students are fascinated by this demonstration.

Flatworms

Students can best observe a live planarian by placing it in a small amount of pond water in a watch glass. The watersurface should be only about 2 cm in diameter for easy viewing with a dissecting microscope. The white surface of thebase plate should be uppermost. Using a watch glass overcomes the thigmotaxis behavior of planaria, which is a prob-lem when they are placed in other types of dishes.


The feeding behavior can be demonstrated by placing a very small piece of liver in the center of a small Petri dishcontaining 2–3 planaria. Culture water in the dish should be about 2 mm deep.

The preference for dark areas is easily shown by taping black construction paper over half the lid of a Petri dish con-taining about 25 planaria in pond water. The paper is attached by a “hinge” of masking tape so that the paper may be lift-ed for observations without moving the lid or dish. A small tab of masking tape attached to the paper opposite to the“hinge” is used to attach the front edge of the paper to the lid as well as to lift the paper for viewing. Use ambient light ora fluorescent lamp for illumination. Do not use an incandescent laboratory lamp, since heat is damaging to the planaria.

A fingerbowl is best for demonstrating the response of planaria to a water current. The water may be gently stirredin a circular motion with a magnetic stirrer or a glass rod at a very slow speed to demonstrate the response. Use 10–20planaria per dish.

Roundworms

Students perform their first dissection in this exercise. A brief discussion of the purpose of dissection is helpful.Some students tend to want to “cut things up” rather than separate organs and tissues to expose structures for observa-tion. Students perform the Ascaris dissection best using only scissors, probe, forceps, and dissecting needles. A scalpelis not needed unless you wish them to gain experience in using one.

Students need to be reminded that the body wall of Ascaris is paper-thin and that they must be careful to avoiddamaging underlying organs. Pinning Ascaris near an edge of the dissecting pan will enable examination of the speci-men with a dissecting microscope. The addition of water to the pan to cover the worm after it is pinned out will preventdrying of the tissues and facilitate observations.

A slide of living Turbatrix may be set up as a demonstration if time is limited. The use of a microscope video sys-tem is very effective for this demonstration.

Exercise 15. Mollusks, Segmented Worms, and Arthropods

As in other diversity exercises, the emphasis here is on the recognition of distinguishing characteristics.

Mollusks

Soak the clams overnight in water to remove much of the preservative. You may wish to present a dissected clam asa demonstration because it is a difficult dissection for students. Two to four dissected specimens with the major struc-tures labeled with pins are adequate for an entire class. They will keep several days if covered with water.

Frozen squid are better than preserved squid for student study. They are usually available at fish markets. Emphasizethe major adaptations of cephalopods and their predatory lifestyle in this section. Dissection of the squid is optional.

Annelids

Soak the earthworms overnight in water to remove some of the preservative. Students have little difficulty with theearthworm dissection if they pin it to the dissecting pan dorsal side up. They should be reminded to pin it near one edgeof the dissecting pan so that it may be examined under a dissecting microscope. You may wish to again emphasize theobjectives of a dissection.

Arthropods

The emphasis in this section is on crustaceans and insects, although representatives of other classes are to be examined.

Crustaceans

Soak the crayfish overnight in water to remove some of the preservative. It is helpful to remind students to corre-late structure with function as they examine external structures and do the dissection. Caution students to gently removethe carapace to avoid damaging the underlying heart and attached blood vessels. Non-injected crayfish are best for stu-dent dissection, but injected specimens may be helpful for demonstration dissections.

Insects

The prime objective is for students to recognize the distinguishing characteristics of insects, and this can be ac-complished without dissection. If time is limited, the grasshopper dissection may be set up as a demonstration or omit-ted. Soak the grasshoppers overnight if dissection is to be done. Students tend to have difficulty with this dissectionmainly because the coagulated hemolymph makes dissection “messy.” When informed of what to expect and if the con-gealed hemolymph is carefully removed with forceps before proceeding, students perform this dissection quite well.

A good metamorphosis display, including live specimens if possible, helps students distinguish the types of meta-morphosis among insects.


Exercise 16. Echinoderms and Chordates

The exercise focuses on the evolutionary relationships of chordate groups along with their distinguishing characteristics.

Echinoderms

If a marine aquarium is available, observation of tube feet in action in living sea stars or sea urchins is a valuableexperience for students. Soak the preserved sea stars in water overnight to remove much of the preservative prior totheir use in dissection.

Chordates

Observation of living vertebrates is a valuable experience for students when comparing characteristics of the variousgroups. Many students have never observed closely or never handled some of the common representative vertebrates, anddoing so is a valuable experience.

A demonstration dissection of a frog or fetal pig to show the basic vertebrate body plan is a useful addition, if youdo not plan to dissect either a frog or fetal pig in the course. Label the organs with pins.

PART IV ANIMAL BIOLOGY

Exercise 17. Dissection of the Frog

This exercise is provided for those instructors who prefer to use a frog, rather than a fetal pig, as the vertebratedissection animal. This dissection does not cover skeletal and muscle systems. Instead, it focuses on the organs of theventral cavity and the basic organization of the vertebrate body plan. The entire dissection has been placed in a singleexercise to provide optimum flexibility for the instructor. It may be completed in a single lab session or may beworked on over several lab sessions as exercises on the various organ systems studied. Students have no problem withthe dissection if they follow the directions provided.

Exercise 18. Dissection of the Fetal Pig

The fetal pig is the preferred dissection vertebrate for beginners in biology. The entire dissection has been placed ina single exercise to provide optimum flexibility for the instructor. The dissection may be done in a few successive labsessions, or portions may be done in conjunction with exercises on the various organ systems. It is helpful to emphasizethe general dissection guidelines described early in the exercise before students begin.

Students have little difficulty with the dissection if they follow the directions in the manual. However, they usuallyneed help in the identification of the blood vessels anterior to the heart and some of the urogenital ducts and organs.

Exercise 19. Blood and Circulation

Students have little difficulty with this exercise if they understand the procedures to be followed before startingeach part of the exercise.

Human Blood

Students should know the recognition characteristics of the various blood cells from studying the Schilling bloodcharts before they examine the blood slides. It is a good idea to set up demonstration microscopes for students to viewbasophils and eosinophils since they are difficult for students to locate. A microscope video system is excellent fordemonstrating the characteristics of blood cells.

Blood Typing

Some instructors have chosen to not have students type their own blood because of the fear of accidental transmissionof HIV. Others continue this useful experience for students, but only with the direct instructor supervision of each student.If you do not wish to use human blood for blood typing, Wards offers an ABO Blood Simutype Kit and a Rh Blood Simu-type Kit. These may be used in place of human blood and antisera to demonstrate the principles of blood typing.

Blood typing presents no problem for students if they follow the directions and do not contaminate the antisera.You should remind them to (1) not touch the blood with the antisera droppers and (2) use separate toothpicks to mixeach blood–antiserum combination.

Infection Control

When students determine the type of their own blood, be sure to explain the specific procedures that you wish themto follow to prevent the possibility of transmission of blood-borne viruses. The potential presence of HIV and hepatitis


viruses in students’ blood demands special precautions. It is a good idea to have only one student working at a slide-warming box at a time, under the direct supervision of the instructor. Students may need to be reassured that the proce-dures do not cause foreign blood to be introduced into their blood by these tests. They must be reminded to follow yourdirections and laboratory procedures carefully. It is also a good idea to make participation a voluntary activity. The fol-lowing minimal safety precautions are suggested:

1. Students should pierce their own fingers or have them pierced by the instructor to eliminate possible student con-tact with foreign blood.

2. Fingertips should be cleansed with 70% alcohol prior to piercing. Use a good disinfectant or 0.5% Amphyl onpierced fingertips afterward.

3. All materials in contact with blood should be placed in a biohazard bag for autoclaving and disposal. Remind studentsto place used alcohol wipes, toothpicks, paper towels, glass slides, and anything else in contact with blood, immediatelyafter use, into the biohazard bag. Used lancets should be placed immediately in a biohazard sharps container.

4. Use new, clean glass slides for blood typing. They can be placed across the special blood typing plate on the warm-ing box. This allows the use of the divisions on the underlying typing plate as a guide for placing the drops of bloodon the glass slides. Students should place the slides in the biohazard bag immediately after use. If you wish to savethe slides, they should be placed in 0.5% Amphyl or 10% bleach. The slides should not be washed until they havebeen in the disinfectant solution for at least 30 min; and the dishwasher should wear gloves.

5. Tables, warming box, and typing slide should be washed with a disinfectant (0.5% Amphyl or 10% bleach) beforeuse and after the laboratory session.

6. Autoclave the biohazard bag and contents prior to disposal.

Circulation of the Blood

The use of fresh or frozen sheep hearts for the heart dissection is highly desirable. Most students react negativelyto preserved hearts. If you cannot obtain them locally, frozen sheep hearts are available from Wards by special arrange-ment. If preserved hearts must be used, soak them overnight in water to remove some of the preservative. Students haveno difficulty with the dissection if they follow the directions provided. Students should know the structure of the heartfrom a study of the figures and text before they start the dissection.

The frog’s foot used to demonstrate capillary blood flow must be kept wet for good results. Also, keep the wrappingcloth moist as well. The microscope light should be turned off when students are not viewing the capillary circulation. Ifyou prefer, a goldfish’s caudal fin may be used instead of a frog’s foot for this demonstration. Just wrap the goldfish inwet cotton and pin the cotton to a frog board, with the caudal fin projecting over the opening.

Blood Pressure

Students enjoy this activity once they become familiar with the procedures. Caution them about leaving the inflat-ed cuff closing the brachial artery for more than 30 s. They often need guidance in placing the stethoscope over thebrachial artery, located in the medial half of the anterior surface of the elbow joint.

Exercise 20. Gas Exchange

This exercise covers gas exchange in animals, especially humans.

Demonstration of Frog Lungs

Care must be taken to not squeeze the frog too tightly during pithing or the lungs will be deflated. If this occurs, thelungs may be inflated by snugly inserting a small, bore glass tube, or medicine dropper into the glottis and forcing airinto the lungs with a rubber bulb; then, clamp the bulb to prevent escape of the air. The lungs and heart must be keptmoist with frog Ringer’s solution. Keep the desk lamp turned off, except when observations are made, to avoid damageto the tissues by excessive heat. Better yet, use a fluorescent lamp for illumination. Explain to students that the frog isbrain-dead and is not experiencing pain.

Sheep Pluck

Use a fresh or frozen sheep pluck, if at all possible, for students to examine. Most students respond negatively topreserved plucks. It is important that students wear protective vinyl or latex gloves when handling fresh or previously


frozen plucks. And remind them to wash their hands thoroughly afterwards. Inflated and dried sheep lungs also are auseful demonstration.

Lung Capacity

Four to six Propper spirometers are usually adequate per class. Students should be reminded of the importance ofdisinfecting the intake tube of the spirometer with 0.5% Amphyl or 10% household bleach, before adding a sterile dis-posable mouthpiece. The used mouthpieces should be placed in a biohazard bag immediately after use, and the bagshould be autoclaved prior to disposal. Students should also be reminded not to inhale through the spirometer. Thespirometers should be cleaned with 10% bleach after the lab.

Exercise 21. Digestion

This exercise focuses on both anatomy of the digestive tract and enzymatic digestion in humans.

Digestion in Paramecium

Prepare the stained yeast cells for Paramecium as follows:

1. Suspend 5-g dried yeast in 50-ml water and heat to kill all yeast cells.

2. Add sufficient Congo red dye to color the mixture medium red.

3. After 30 min, filter through rapid-filtering filter paper. Discard the filtrate.

4. Re-suspend a portion of the stained yeast cells in the Paramecium culture 1 to 2 h prior to the laboratory session.

Digestion of Starch

This experiment is best performed by groups of 2–4 students who share the responsibilities. The data is shared withthe entire class.

The action of pancreatic amylase is used to illustrate the action of enzymes, in general, and digestive enzymes, inparticular. It is important to emphasize to students that (1) they must understand the procedures to be followed beforestarting the exercise and (2) the droppers of dropping bottles must not be contaminated by touching other solutions.

The rubber bulbs of the droppers should be checked before the lab to be sure that they are in good condition andwill dispense approximately equal-sized “droppers” (about 1 ml) of solution. Remind the students of the difference be-tween drops and droppers of solution.

Preparation of Soluble Starch

Prepare the 0.1% soluble starch solution by mixing the starch into distilled water. Heat the mixture, but not to boil-ing, while stirring until the starch goes into solution. Store the starch solution in a stoppered flask in the refrigeratoruntil needed, but allow time for it to return to room temperature before use. It is important to test a sample of the starchsolution before the laboratory session, to be sure that it gives a positive reaction with iodine solution.

Use light-proof dropping bottles for Benedict’s solution and for iodine solution (I2KI).If it is desirable to divide the experiments among student groups with all groups sharing their data, it is best to do so ac-

cording to assigned temperatures rather than assigned pH. This reduces the movement of students in the lab and minimizesthe need for a large number of water baths. The disadvantage is that most, if not all, of the digestive action occurs at 37º C.

When drops of amylase and starch solutions are added to the test tubes, they should not run down the side of thetube, but should land squarely in the bottom, for best results. The tube contents should be quickly mixed using a vortexmixer, or by shaking the tube from side to side, as soon as the solution has been added.

When testing tubes 1B–9B for the presence of maltose, remind students to add boiling chips to the water-bathbeaker and to heat only 3 tubes at a time. These procedures will help prevent the spatter of boiling water.

Exercise 22. Excretion

Kidney structure and function are emphasized in this exercise.

Kidney Dissection

Fresh or frozen sheep kidneys are superior to preserved ones for dissection. If the kidneys are still encased in fat, itcan best be removed by breaking it off with the hands. The use of a scalpel tends to promote damage to the kidney. Stu-dents should use protective vinyl or latex gloves when handling fresh animal parts, and they should wash their handsthoroughly afterward.


Condition Recipe for Urine Preparation

Normal Add tap water to 30 ml of urea broth concentrate to bring to one liter; add 10% HNO3

to adjust pH to between 6.0 and 7.0 (may require about 10 ml); if necessary, add NaClto adjust specific gravity to about 1.010.

Diabetes insipidus Dilute simulated “normal urine” with tap water: 5 parts water to 1 part “normal urine.”

Diabetes mellitus Add glucose to “normal urine” to yield a 5% solution (0.6 g glucose per 12-ml “normal urine”); elute 1 ketone portion from a CheckStix for each 12 ml of solution; adjust pH with 10% HNO3 to 5.5 to 5.8; if necessary, adjust specific gravity to about 1.035 with sodium chloride or additional glucose.

Glomerulonephritis Add albumin to “normal urine” to make a 0.25% solution (30-mg albumin per 12-ml“normal urine”) and add 1 drop sheep blood per each 24-ml solution. If necessary, use10% HNO3 to adjust pH to 5.0 to 6.0.

Hemolytic anemia For each 12-ml simulated “normal urine,” add 1 drop of sheep blood.

Hepatitis For each 12 ml of “normal urine,” elute 1 bilirubin portion from a CheckStix; adjust pH with 10% HNO3 to 5.8 to 6.0; if necessary, adjust specific gravity to about 1.020 withNaCl.

Strenuous exercise and Add albumin to “normal urine” to make a 0.05% solution (6 mg per 12-ml “normal a high protein diet urine”).

Urinary tract infection For each 12 ml of “normal urine,” add 1 drop of blood and 12-mg sodium nitrite.

Preserved, triple-injected kidneys are excellent for demonstrating the glomeruli and nephron capsules when sec-tioned and viewed with a dissecting microscope.

Urinalysis

This experiment uses simulated urine samples that are analyzed using Multistix 9SG. If you wish, students may an-alyze their own urine in the same fashion. Once students get over their initial squeamishness about providing a urinesample, they have no problem doing their own urinalysis. Students have no problem with the experiment if they care-fully follow the directions.

It is imperative that students understand the test provided by each reagent band on the “dipsticks.” To do this, acolor chart provided with the “dipsticks” must be compared to the reagent bands on a “dipstick.” The sequence ofreagent bands from the tip to the handle of a “dipstick” matches the sequence of specific tests listed from top to bottomon the color chart. When ordering your Multistix 9SG “dipsticks,” request a supply of additional color charts. The colorcharts may be laminated for repeated use by students.

Matching the unknowns with clinical conditions provides an intriguing challenge for students, and it helps them tounderstand the clinical value of a urinalysis as well as kidney functions.

If students analyze their own urine samples, use disposable cups for collecting the urine samples. The cups, Multi-stix strips, and any paper towels in contact with urine should be placed in a biohazard bag for autoclaving and disposal.Urine should be flushed down a toilet in the nearest restroom.

Preparation of Simulated Urine Specimens

The use of glucose, albumin, sheep’s blood, and CheckStix (Ames) makes preparing positive controls and un-knowns a simple matter. A CheckStix contains six zones (squares) impregnated with various substances. From thehandle end of the stick to the tip, the substances present are as follows: Square 1: nitrite, blood, glucose, protein;Square 2: nitrite, blood, glucose, protein; Square 3: urobilinogen; Square 4: bilirubin; Square 5: ketone, protein;Square 6: ketone. Simply cut the squares of substances from the stick and place them in the preferred combinationsfor unknowns, one square for each 12-ml tap water for 30 min. Remove the plastic squares prior to use. A drop ortwo of iodine solution may be used to create the desired color. The unknowns will remain viable for 24 h and aregood for at least 12 tests with Multistix, before the values begin to change. The following recipes give good results,or you may create your own.


Exercise 23. Neural Control

This exercise poses no difficulty, but students must use their time well.

Sheep Brain

Preserved sheep brains should be sectioned and soaked prior to the lab session to remove much of the preservative.Students have no problem locating the major parts of the sheep brain if they have learned the background information.

Reflexes

Students have little difficulty with the patellar and photopupil reflexes. Occasionally, it is difficult to divert a stu-dent’s attention to get a good response of the patellar reflex.

Reaction Time

Students enjoy this activity. It teaches them about the factors that determine reaction time and that practice can im-prove reaction—but only so much. The experiments will show that reaction time varies among persons and that it islonger in dim light. You may wish to discuss the practical implications of a longer time in dim light, that is, when dri-ving. It is best done by pairs of students.

Exercise 24. Sensory Perception in Humans

This exercise enables students to examine their own perception of sensory sensations.

The Eye

Students enjoy the eye dissection more than they think they will at first, and it is quite informative. Fresh or frozeneyes are best. Fresh or frozen beef eyes can be obtained from Wards’ New York Office by special arrangement, and it isworth the effort.

Students have little difficulty with the visual tests, and they enjoy doing them. You may have difficulty obtaining anIshihara Book of Color Plates for the colorblindness test. It is distributed only by the Graham-Field Surgical Company,415 Second Avenue, New Hyde Park, New York, 11040. This company can identify a supplier near you. An alternate se-ries of test plates, the Ichikawa color plates, for detecting colorblindness are available from Wards and other biologicalsuppliers. If you use the Ichikawa plates, the normal and abnormal responses will differ from those noted on the labo-ratory report for this exercise.

The Ear

The most difficult part of the hearing tests is finding a quiet place to perform the tests. Students sometimes need tobe reminded of the correct use of the tuning fork.

The static balance test may be demonstrated using one student for the entire class, if desirable. Students enjoy thistest, and they are usually amazed at the importance of visual input in maintaining balance in sighted persons.

Skin Receptors

Students generally have no difficulty with these experiments. However, it takes a little time for students to deter-mine the smallest change in water temperature that is detectable. If time is limited, you may wish to omit this part ofAssignment 5 under Intensity of Sensations.

Taste

The purpose of these tests is to allow students to discover that (1) substances must be in the solution to be tasted,and (2) taste cells for the four tastes are scattered over the tongue and are not restricted to the taste zones shown in Fig-ure 24.7. Tests for bitter have been omitted because of the likelihood of producing gagging. If you wish to try it, alumpowder will produce a bitter taste. It is important to prevent contamination of the test solutions and to ensure the safetyof introducing the solutions in students’ mouths.

Exercise 25. Chemical Control in Animals

The role of the thyroid gland on metabolic rate and the action of acetylcholine and epinephrine on heart rate are ex-amined in this exercise.


Thyroxine and Metabolic RateThis experiment requires respirometers similar to the one shown in Figure 23.1 and several days of prior prepara-

tion. Students should work in groups of 2–4 to provide a division of labor.Begin this experiment 10 days prior to the laboratory period. Best results are obtained by using thyroxine and

potassium perchlorate to accentuate the normal condition of the mice. Do this by determining the metabolic rate foreach mouse. Use the mouse with the highest metabolic rate as Mouse A, hyperthyroid. Use the mouse with the lowestmetabolic rate as Mouse B, hypothyroid.

The hyperthyroid condition is produced by providing Mouse A with drinking water containing 0.02% syntheticthyroxin (sodium levothyroxin). It may be obtained in 10-ml vials containing 500-mg thyroxin from companies thatsupply veterinary drugs. One such company is the Flint Division of Trovenol, Deerfield, Illinois.

Contact a local veterinarian to determine the source nearest to you. Prepare the drinking water by placing 4-ml (200mg) sodium levothyroxin in a liter flask and by filling with water to 1 liter. If the solution is a bit bitter, add artificialsweetener to make it more palatable.

The thyroxin solution decreases the surface tension of water, so the drinking tube must be constricted over a flameto prevent dripping. Some experimentation is needed to determine the optimum bend and terminal diameter of thedrinking tube.

The hypothyroid condition is produced by providing Mouse B with drinking water containing 0.25% potassiumperchlorate. This solution tends to be bitter, so add sufficient artificial sweetener to yield a palatable taste.

Suspend the drinking bottles outside the cages for easy accessibility. Bottles holding 200 ml or larger will ensure a con-tinuous supply of water overnight. Mark the level of the drinking water in the bottles to enable a check on fluid consumption.

Place the food nuggets in a feeding rack outside the cage to prevent food from being lost in bedding material. Thiswill facilitate the keeping of an accurate record of food consumed since residual food is easily located.

Chemical Control and Heart Rate

This experiment works very well to show the effect of very small quantities of acetylcholine and epinephrine ontarget tissues. It may be done as a demonstration if you prefer, especially if a video camera and monitor can be used toprovide easy viewing by the entire class at once.

Double-pith the frogs before the laboratory session, not in front of the class. Be sure that the students understandthe result of double-pithing a frog. (This usually leads to consideration of indicators of death and the progressive natureof death in various tissues.) Most students handle the dissection well, but they should be reminded to carefully avoiddamage to the blood vessels when cutting through the length of the sternum.

Students must understand how to correctly read the volumetric divisions on the hypodermic syringe to provide an ac-curate dosage of each chemical. Determination of the heart rate without a recorder is best done by one student counting thecontractions and another monitoring the start and end of the counting period (15 s). Careful control of the hypodermic sy-ringes and needles is advised.

Exercise 26. The Skeletal System

This exercise is trouble-free, but it is wise to remind students to use pipe cleaners, rather than pencils or pens, aspointers when examining skeletal material.

Skeletal Adaptations for Locomotion

Observing the locomotion of a live frog, bird, and cat in the lab, in addition to skeletal mounts, is helpful when stu-dents are correlating the adaptations of the skeletons of these animals and their modes of locomotion.

Exercise 27. Muscles and Movement

Students sometimes need help in recognizing the different types of levers, although this usually resolves itself asthey work through the section on levers in the exercise.

Frog Muscles

Freshly-killed frogs are easier to work with, but more costly, than preserved frogs for determining the function ofleg muscles. They also permit students to see the natural appearance of muscles and connective tissue. Generally, stu-dents have no problem with this section once they have correctly located the muscles.


Myofibril Structure and Contraction

Glycerinated rabbit psoas muscle kits, including reagents, may be purchased from biological supply houses. Priorto the lab session, remove the muscle tissue from the tube and cut it into 2-cm strips. Place the strips in a small dish ofglycerol. Tease the fibers apart so that the strands are 0.2-mm thick, or less.

Exercise 28. Reproduction in Vertebrates

An understanding of meiosis is necessary for this exercise. Also, you may wish to provide a brief discussion of theovarian and uterine cycles if they have not been covered in lecture.

Birth Control

Students are very interested in this section. The availability of various birth control devices and chemicals for ex-amination is helpful, but you will need to use your own judgment as to how best to handle consideration of this topic,which is quite sensitive for some students.

Exercise 29. Fertilization and Development

Many students consider this exercise to be the high point of the course. Supply houses providing marine specimenshave developed sea urchin kits that can be shipped overnight by air to almost any part of the country, and they guaran-tee viable urchins in reproductive condition. The kits come with directions for the care of the urchins. The proceduresfor the extraction and preparation of the gametes are not difficult, but must be followed carefully if good results are tobe obtained. One kit will serve 4 to 6 laboratory sections.

It is not necessary to have a marine aquarium in order to do this exercise. The sea urchins can be kept for severaldays in a refrigerator if kept as they are shipped, that is, wrapped in newspapers soaked in seawater.

Allow 5–10 min after injection for the sea urchins to start releasing the gametes. Shedding is usually complete 15min after the start of gamete release. It may be necessary to inject 5–6 sea urchins to obtain gametes from each sex.

Discard the sea urchins after use. Do not return them to a marine aquarium containing other sea urchins. If this isdone, it will initiate the release of gametes from all the ripe sea urchins in the aquarium.

Remind students to place only a few eggs on the depression slide since they can deplete the oxygen rapidly andyield poor results. Usually 6–10 eggs give good results. After activation, the slide must be kept in the refrigerator at20º C between observations. Observations should only be for about 5 min at a time; longer periods expose the embryosto excessive heat from the microscope lamp.

Prepare embryos at different stages by adding 4 to 6 drops of sperm suspension to a fingerbowl containing about200 eggs in 100-ml seawater. Stack the fingerbowls for storage in the refrigerator. Plan to have embryos at approxi-mately the following ages: 6, 12, 24, 48, and 96 h.

You may wish to set up demonstration slides of the embryos of different ages. If students prepare their own slides,transfer about 50 eggs and 100-ml seawater to Petri dishes that are labeled with the ages of the embryos. Students canobtain the embryos from the Petri dishes and should not endanger the embryos in the “stock cultures.” By sharingslides, students can reduce the number of slides to be made.

Chordate Development

This section emphasizes the early similarities with echinoderm development and the basic chordate pattern throughthe neurula development. Color-coding the figures is an important learning tool here and in the next section.

Human Development

Students are very interested in the stages of human development, and they are usually amazed that an 8-week fetusis so humanlike. A series of fetuses, labeled as to age, is very informative.

Exercise 30. Early Embryology of the Chick

Fertilized chicken eggs should be ordered for delivery no more than 3 days before they are to be incubated. Storethem in a cool place until incubation is started, but do not place them in a refrigerator. Lengthy storage or refrigerationwill reduce the viability of the chick embryos.


If fertilized eggs are to be incubated for several laboratory classes in the same incubator, make a schedule of start-ing times for each batch of eggs. The date and hour at which a batch of eggs will reach 48 h of incubation should bewritten on the eggshell. Before placing the eggs in the incubator,

1. put a fingerbowl of water in the incubator to provide necessary humidity,

2. be sure that there is an adequate oxygen supply,

3. stabilize the incubator temperature at 39º C.

Once in the incubator, rotate the eggs 1/4 turn every 8 h, or as close to this schedule as possible.

The 48-Hour Embryo

An egg should be supported while opening the shell by nesting it in crumpled paper towels in a fingerbowl. Keepthe embryo warm with a laboratory lamp while opening the egg and removing it to the warmed slide.

PART V PLANT BIOLOGY

Exercise 31. Structure of Flowering Plants

This exercise poses no difficulty for students.

General External Structure

The first portion of this section focuses on (a) the distinguishing characteristics of monocots and dicots and (2) theexternal structure of flowering plants.

If Coleus plants are difficult to obtain, geraniums or snapdragons are quite satisfactory for the study of externalstructure, but Coleus is best for locating vascular tissue in a herbaceous dicot after they have been stained by uptake ofa water-soluble dye. Corn seedlings, 2 to 3 weeks old, are the best to show the location of vascular bundles in a mono-cot after uptake of a water-soluble dye. Single-edged razor blades are usually better than scalpels for sectioning stemsof both Coleus and corn.

Flowers, fruits, and seeds were covered in Exercise 13 and are not covered here.

Internal Structure

The rest of the exercise focuses on internal structure of roots, stems, and leaves. Students have little difficulty here.

Preparation of Germinated Grass Seeds

1. Place two filter papers in the bottom of a Petri dish and wet them with 0.1% proprionic acid. Proprionic acid is amold inhibitor.

2. Sprinkle the grass seed over the filter paper and add the cover.

3. Incubate at room temperature for 7 to 10 days.

If a shorter germination period is helpful, radish seeds can be germinated by the same technique in 3 days.

Root Types

Tap and fibrous root systems can best be demonstrated by washing away the soil and placing the roots of plants ina beaker or flask of water. This will keep the smallest roots in good condition.

Leaf Types

A good diversity of leaf types helps students to understand the different forms of venation.

Exercise 32. Transport in Plants

The key to the success of this exercise is in preparing the setups for root pressure and transpiration. Students shouldwork in pairs.

Root Pressure

For the root pressure demonstration, use a well-watered geranium plant whose stem is about the same diameter asa 1-ml pipette. The rubber tubing can be placed over the cut plant stem with minimal damage by (1) rolling the rubber


tubing back on itself and (2) then rolling it over the cut end of the stem. A little petroleum jelly on the side of the stemhelps to make a good seal, but do not get it on the surface of the cut stem because it will plug the xylem vessels. If nec-essary, the tubing may be secured with string, but do not tie it tight enough to damage the stem. Add a little water to theopened end of the tubing and insert the pipette far enough to force the water up into the pipette to the 0.1-ml mark. Alittle petroleum jelly on the pipette facilitates its insertion into the tubing. Support the pipette with a ring stand andclamp.

Germinated Grass Seed

Allow 5 to 7 days for the grass seed to germinate. Prepare the germinated grass seeds as described below. If youprefer, radish seeds can be germinated in 3 days by the same protocol.

1. Place two filter papers in the bottom of a Petri dish and wet them with a 0.1% proprionic acid solution. Proprionicacid is a mold inhibitor.

2. Sprinkle the grass seed over the filter paper and add the cover.

3. Incubate at room temperature for 5 days.

Transpiration

For the potometer setup, use a woody stem whose diameter is slightly less than that of a 1-ml pipette. Prepare thesetup as follows:

1. Insert the 1-ml pipette into one hole of the 2-hole rubber stopper before the leafy branch is cut from the plant.

2. Immerse the cut end of the branch or shoot in water as soon as it is cut from the plant.

3. Recut the end under water at an oblique angle using a very sharp knife or scalpel. This cut is to ensure that thexylem cells are not crushed and that air has not entered them.

4. Keeping the cut end of the branch or stem under water, insert it completely through the hole of the stopper takingcare not to damage the stem.

5. Immerse the potometer jar under water and fill it. Keeping the jar and stopper under water, insert the stopper intothe jar while being careful not to trap air bubbles.

After determining the rate of transpiration without and with a breeze from an electric fan, students are encouragedto design their own experiments to evaluate the effect of other climate factors (e.g., temperature or humidity) on tran-spiration. Measuring the effect of temperature is easier since a heat lamp may be placed at varying distances from thepotometer and the temperature easily measured.

Density of Stomata

Students may need assistance in determining the surface area of a leaf and in making the necessary calculations.They are usually amazed at the number of stomata on a leaf.

If leaves of plants adapted to different types of habitats (e.g., sunny and dry vs moist and shady) are used in this partof the exercise, the stomatal density and leaf size can be correlated with habitat adaptation.

Exercise 33. Chemical Control in Plants

Parts of this exercise require several days or a few weeks to obtain results from the experiments. If this is not pos-sible, these experiments may be set up as demonstrations.

IAA and Oat Seedlings

Germinated oat seedlings with intact coleoptiles should be available for student examination so that they will bet-ter understand the described experiments using oat coleoptiles. It is not practical to try to replicate these experiments,except for Experiments 1, 4, or 5. However, a demonstration of one or more of these experiments is very informative tostudents. Suggestions for their preparation are noted below.

1. Husk the oat seeds and soak them for 2 to 3 h.

2. Place a layer of cotton in a fingerbowl and soak it with 0.1% proprionic acid. This provides the necessary moistureand also prevents fungus growth.

3. Place the oat seeds on the wet cotton with the grooved side down.

4. Cover with a second fingerbowl and place in the dark at room temperature for 72 h. All of the seedlings in one fin-gerbowl should be used to demonstrate a single experiment.


5. To remove the coleoptile tips, hold a millimeter ruler alongside a coleoptile and use a razor blade to cut off the top3 mm of the coleoptile. Discard the tips. Be careful not to dislodge the seedlings from the cotton.

6. Measure and record the length of the cut and uncut coleoptiles in millimeters and place the covered finger bowl inthe selected environment for 24 h. Students can make the measurements at the end of the experiment.

7. Seedlings may be grown horizontally by gently manipulating the cotton around the base of the seedlings with for-ceps until they are in the desired position. Alternatively, cover the fingerbowl with Saranwrap and place it on edgewith supports to prevent movement. Mark the side of the fingerbowl that is upward.

8. Add additional 0.1% proprionic acid to the cotton, if necessary, to provide sufficient water for the seedlings.

Tropisms in Dicots

Coleus plants work well, and so do snapdragons and tomatoes, so you can go with whatever is available at yournursery. These experiments allow students to correlate the experiments described for oat seedlings with their own ex-periments.

Biological supply houses sell IAA in lanolin paste (0.5% or 5,000 ppm) and as a powder.

Gibberellins

Gibberellins promote elongation in stems and retard germination of seeds. Biological supply houses sell gibberellicacid (GA3) as powder and in prepared 0.1% solutions. If starting with powder, dissolve 0.1-g gibberellic acid powder in2.5-ml of 95% ethanol. Then add about 90-ml distilled water and gently heat to drive off the alcohol. Add distilled waterto raise volume to 100 ml of a 0.1% gibberellic acid solution. Use this solution to apply to bean stems.

Preparation of Pour Plates with Gibberellic Acid

1. For 10 plates, prepare 100-ml plain agar in a beaker and cool to near 50º C.

2. Add 0.1-g gibberellic acid and mix.

3. Pour agar into 10 Petri plates (about 10 ml/plate), cover and refrigerate until used.

Abscisic Acid

Abscisic acid inhibits seed germination. It is also known as Dormin at nurseries, in case you have difficulty findingit from supply houses.

Preparation of Pour Plates with Abscisic Acid

1. For 10 plates, prepare 100-ml plain agar in a beaker and cool to 50º C.

2. Add 0.01-g abscisic acid and mix.

3. Pour agar into 10 Petri plates (about 10 ml/plate), cover and refrigerate until used.

Assignment 8

If students wish to investigate the inhibitory effect of high IAA concentrations on roots, snapdragons or beanseedlings are good subjects. The plants should be removed from soil, washed, and their roots placed in an IAA solutionin a test tube. Use clay to seal the top of the tube and to support the plant. Prepare the stock IAA solution by dissolving100-mg IAA in 1.5-ml absolute ethanol. Add about 900-ml distilled water and gently heat to drive off the ethanol. Thenadd distilled water to bring to one liter. Use this 100-mg/l solution to make the dilutions. The concentration range test-ed should be 0.01, 0.1, 1, 10, and 100 mg/l.

Alar (B Nine) is a synthetic plant regulator that disrupts formation of IAA, so it has the opposite effect of IAA onplants. It is applied as a 1% solution using an atomizer to wet the entire plant. Adding a wetting agent to the solution (1drop/50 ml) is desirable.

PART VI HEREDITY AND EVOLUTION

Exercise 34. Heredity

As students begin the study of genetic crosses, it is important that they relate (1) gene pairs with chromosome pairs,(2) segregation to gamete formation by meiosis, (3) independent assortment to non-linked genes, and (4) recombinationof genes to the union of egg and sperm nuclei at fertilization. Experiments involving crossing over are not included in thisexercise.


Students should diagram each step in working out the genetic crosses to establish these relationships in their mindsuntil they are proficient in solving genetic problems. The determination of possible gamete genotypes is the key to mostgenetic crosses. Some students may need assistance at this point for dihybrid crosses.

Some of this exercise can be done outside of the laboratory, if time is limited. At any rate, students should completetheir study of the background information before coming to the lab session. If you cover genetics only in lecture, as-signing portions of this exercise as homework will facilitate student understanding.

Dominant-Recessive TraitsIn this section, students are asked to tabulate progeny (corn seedlings) of crosses because this helps to “get them in-

volved” in the lab. Start the trays of corn seedlings 2.5 weeks prior to use so that the phenotypes are easily recognizedand the albino plants are still in good health. Prior to the laboratory session, remove seedlings as necessary so that thenumbers are pretty close to a 3:1 ratio. In this part of the exercise, students will not use Chi-square analysis, so this ar-tificial adjustment is necessary for students to recognize the type of inheritance.

Tabulate the human phenotypes (item 2k on the laboratory report) of the entire class on the board, so the frequen-cies of the phenotypes are readily apparent to students. The objective is for students to recognize that dominant traits arenot always more frequent in a population.

Dihybrid CrossAs noted above, plant the corn kernels 2.5 weeks before the lab session. Prior to the lab session, remove seedlings

as necessary, so the numbers are pretty close to a 9:3:3:1 ratio. A Chi-square analysis is not done at this point in the ex-ercise, so an artificial adjustment is needed so that students will recognize the inheritance pattern.

Chi-Square Analysis

In counting corn kernels on ears of corn, remind students to mark the starting row with a pin, so they won’t losetheir starting point. Some students may need assistance with the Chi-square calculations.

Exercise 35. Molecular and Chromosomal Genetics

If time is a limiting factor, much of this exercise can be done outside of the laboratory.

Molecular Genetics

The use of a DNA-RNA-Protein Synthesis Kit provides “hands-on” activity that helps students learn the interaction ofDNA and RNA in transcription and translation. As far as I can tell, there isn’t a commercial DNA-RNA-Protein Kit availablethat will do exactly what is needed. Wards (14 W 8330) and Carolina (PN-21-1111) have a useful kit consisting of a teacherdemonstration portion and several student hands-on components. The drawback is that the sequence of nucleotides is fixed,so it can’t be used for the nucleotide sequences on the laboratory report. However, you can make an effective and inexpensivekit using pop beads (Wards #36 W 1550) and pop bead connectors (Wards #36 W 1555) together with a couple of sheets ofconstruction paper, cardboard, or plastic. The kit does not include a simulated ribosome. Prepare a kit for each student group.

Kit Composition

1. Use color-coded pop beads to simulate nucleotide bases: red = adenine; white = thymine; black = uracil;blue = guanine; green = cytosine. Pop bead connectors are used to separate the nucleotide bases in triplets for eas-ier recognition.

2. Cut out small circles, squares, rectangles, and so on, from construction paper, cardboard, or plastic and write thenames of the amino acids on them—including methionine/start and stop. You’ll need multiples of methionine/start,stop, and some amino acids according to the coding on the laboratory report.

3. For each polypeptide to be formed, provide each student group with a sheet of paper divided by four equally spacedheadings along the left margin: DNA nucleotides; mRNA codons; tRNA anticodons; and amino acids. This pro-vides a template on which students can lay out their “molecules.”

Directions for Students

1. Using color-coded pop beads, connect them to indicate the sequence of nucleotides in the parent DNA strand. Usepop bead connectors to arrange the “nucleotides” in triplets for easier recognition. Place the strand of pop beadsrepresenting DNA nucleotides on your paper and use clear or masking tape to lightly attach them to your paper.


2. Using color-coded pop beads, connect them in triplets with the connectors to represent the codons of the mRNAstrand. Place the strand of mRNA codons on your paper and lightly tape it in place so that each codon is directlybelow its complementary DNA triplet.

3. Using color-coded pop beads, connect them in triplets with the connectors to represent the anticodons of thetRNAs. Place the pop beads representing tRNA anticodons on your paper and lightly tape them in place so thateach anticodon is directly below its complementary codon.

4. Using the data in Figure 35.1, place a card representing a specific amino acid coded by an mRNA codon directly belowthe anticodon of the tRNA transporting it. Lightly tape the amino acid sequence to your paper to form the polypeptide.

5. Examine your setup. Note how the sequence of DNA nucleotides determines the complementary codons of mRNA.Each codon of mRNA, in turn, codes for a specific amino acid and is complementary to an anticodon of a tRNAmolecule that transports the specific amino acid into its position in the polypeptide strand.

Chromosomal Disorders

For karyotype analysis, obtain sheets of chromosome spreads from either Wards or Carolina. Chromosome spreadsof normal human male and female and a variety of abnormal chromosome numbers are available. Student groups canprepare karyotypes of different chromosome spreads and share the data.

The mechanics of preparing the karyotypes are usually not a problem, but it will take students nearly an hour foreach karyotype. Small dissection scissors are best for cutting out the chromosomes. It is important to emphasize that thechromosomes should not be glued in position until the correct position has been established. Removable gluesticks aregenerally better than rubber cement and they minimize the mess. Once all chromosomes are in the correct positions, useScotch tape to hold them permanently in place.

Exercise 36. DNA Fingerprinting

This simulation of DNA fingerprinting is based on Edvotek Kit #109, which is adaptable to allow completion of theexercise within a three-hour laboratory period. Of course, kits from other suppliers may be used, but they will need tobe modified in a similar way to enable completion in a single lab session. Edvotek kits are reliable, and the booklets re-ceived with the kits are especially helpful, providing additional information about preparation and operation. Edvotek’stechnical assistance personnel are knowledgeable and courteous.

The exercise calls for the instructor or technician to prepare the agarose gel solution and buffer solution prior to thelaboratory session. Preparation of the gels prior to the lab will provide an additional guarantee that students will finishon time. Allow 1.5 h for this preparation. If you want students to prepare their own agarose gel solution and buffer, anadditional lab session is required. Another time saver used is adding methylene blue to the agarose and buffer solutionsrather than staining the gels after electrophoresis. This enables students to see the bands of restriction fragments as theymove down the gel, and provides sufficient definition to enable interpretation without additional staining. However, de-staining may be required. The gels may be saved, if you wish, by placing them in airtight freezer bags and storing themin a refrigerator.

Preparation of 0.8% Agarose Gel Solution

The quantity of gel solution needed depends upon the size of the gel beds and the number of gels to be used. If youare using Edvotek Kit #109, prepare the gel solution for the entire class as follows. This will provide sufficient agarosefor ten 7 � 7 cm gel beds or five 7 � 15 cm gel beds.

1. Add all of the agarose in the kit (2.5 g) to a 500-ml flask.

2. Add 6.25 ml of the kit’s 50x buffer.

3. Add 305.75-ml distilled water.

4. Add 2.1 ml of the kit’s methylene blue.

5. Use a glass-marking pen to mark the fluid level on the flask.

6. Cover the flask opening with plastic wrap and heat to near boiling with frequent swirling of the contents, until allof the agarose is completely dissolved and uniformly blue.

7. Add distilled water to bring the volume to the original level.

8. After cooling to 55º C, divide the agarose solution equally into smaller flasks that are the same number as studentgroups. Place these flasks in a water bath at 55º C. Each group will then have a flask of agarose for filling a gel bed.


If you know the exact amount needed for each gel bed, place only this amount in each flask, so students simply addall of the gel solution to the gel bed.

Size of Gel Bed Total Volume Needed

30 ml60 ml

Preparation of the Buffer

The amount of electrophoresis buffer required varies with the size and number of the electrophoresis chambers.Prepare the electrophoresis buffer required for each electrophoresis setup in separate flasks, so each student group willhave its own flask of buffer. Prepare the buffer by adding 50x buffer from the kit to distilled water to make a 2% solu-tion. The following table may be helpful.

50� Buffer + Distilled Water = Total Volume

4 ml 196 ml 200 ml6 ml 294 ml 300 ml8 ml 392 ml 400 ml

Distribution of DNA Samples

The Edvotek kit #109 contains six tubes of predigested DNA samples labeled A through F. For ease of operation, itis best to divide each sample into labeled micro-centrifuge tubes for each student group. This prevents congestioncaused when each student group must obtain samples from the same stock tubes, and it reduces the possibility of cont-aminating the stock tubes.

Additional Suggestions

It is important for the instructor to demonstrate how to fill a gel bed and how to use a micropipette.The time students spend practicing filling the wells in the practice gel, using a micropipette, is time well spent.

Practice is essential for good results. Practice gels may be purchased (Carolina Biological) or you can make them your-self using Petri dishes and plain agar.

Remind students to wear protective gloves to prevent skin contact with samples and to prevent DNA contamination.Caution students to be sure to use a new sterile micropipette tip for each DNA sample.After electrophoresis, caution students to turn off the power supply and unplug the leads before removing the

safety cover.A transparent plastic sheet (e.g., Saranwrap) placed on the viewing box will keep moisture off the box and help

keep it clean.

Exercise 37. Evolution

This exercise focuses on evidences for evolution and on the comparative examination of hominid skulls.

Evidence from Vertebrate Embryology

The use of models of vertebrate embryos to show the pharyngeal pouch stages is very helpful. You may wish to setup the embryo slides as demonstrations to save time and to facilitate the location of the key structures by students.

Evidence from Biochemistry

In this section, students will obtain data similar to that in Table 33.2 by using a LAB-AIDS Immunology and Evolu-tion Experiment Kit available from Ward’s. The LAB-AIDS Immunology and Evolution Experiment Kit uses syntheticsera and simulates the assessment of the degree of relationship by synthetic antigen–antibody precipitation. Syntheticsera of the human, the chimpanzee, the orangutan, the monkey, and the cow are tested by synthetic rabbit sera that havebeen sensitized by synthetic human sera. The resultant precipitations give clear qualitative, not quantitative, results.Students respond to this kit quite well.

Skull Analysis

The measurements for the skull analysis can be a bit tricky for students since the structures being measured are notflat surfaces, and since the measurements are made along parallel lines and cannot always be made simply between twoanatomical points.

7 * 15 cm7 * 7 cm


It takes 2–4 students working together to determine the skull measurements. One student holds the skull while 1–2students take the measurements with the calipers and meter stick. Once you have made the measurements yourself, youwill see the difficulties encountered and will be able to advise students on preferred procedures. Ultimate precision is notrequired in the measurements to obtain the desired results. Some students may need help in plotting the data on a graph.

Students should be reminded to be careful in handling the specimens. The use of plastic hominoid skulls rather thanplaster casts is highly recommended. Plastic skull replicas may be obtained from biological supply houses. Plaster skullreplicas are excellent for demonstration purposes and may be obtained from the University of Pennsylvania Museum.

If you cover evidences for evolution only in lecture, assigning parts of this exercise as homework enhances studentunderstanding.

Exercise 38. Evolutionary Mechanisms

Initially, some students may need assistance in determining the values of p and q. If the Castle–Hardy–WeinbergLaw is covered in lecture prior to the laboratory session, students have little difficulty. If you cover this topic only in lec-ture, assigning parts of this exercise as homework really helps student understanding.

Once students have learned how to determine the frequency of the recessive allele following complete selectionagainst homozygous recessives for one or more generations by the “long method,” it is helpful for them to use the fol-lowing formula to facilitate the process. This expression will give the frequency of the recessive allele after completeselection against homozygous recessives for any number of generations.

q0 = the initial frequency of the recessive alleleqn = the frequency of the recessive allele after n generations

If a computer is in the laboratory, commercial software is available at reasonable prices that will facilitate and ex-tend this exercise, especially in areas of genetic drift and natural selection. Most of these programs are available in Mac-intosh or IBM formats.

PART VII ECOLOGY AND BEHAVIOR

Exercise 39. Ecological Relationships

Most of the first portion of the exercise should be completed prior to the laboratory session to allow adequate timefor the analysis of the aquatic ecosystems.

Aquatic Microecosystem Preparation

The simulated ecosystems should be set up in 600- or 1000-ml beakers. Each ecosystem contains certain organismsthat represent the natural biota of one of the three types of ecosystems. They are prepared using cultures from Wards asnoted below, and by adding diluted algae culture medium, as necessary, to bring them to 500 ml.

With these cultured organisms, it is not possible to adjust the pH to that expected in such ecosystems, especially tothe pH in acid ecosystems. If this is done, most organisms would be killed. Therefore, students should be told the pHusually encountered in such ecosystems, and the ecosystems should be labeled accordingly.

Keep the ecosystems covered to retard evaporation and at room temperature. Place them near a window for expo-sure to indirect light or expose them to light from a fluorescent lamp for 8 to 12 h per day.

Prepare the ecosystems as noted below. Add diluted algae culture medium to bring each ecosystem to 500 ml, afteradding the organisms.

1. Normal Ecosystem (pH 5.5–7.0)a. Pandorina: 1 culture for 35 studentsb. Euglena: 1 culture for 35 studentsc. Oscillatoria: 1 culture for 35 studentsd. Microspora: 1 culture for 35 studentse. Spirogyra: 1/2 culture for 35 studentsf. Cyclops: 1/2 culture for 35 studentsg. Daphnia (small): 1 culture for 35 studentsh. Gammarus: 1/2 culture for 35 students

qn = q0

1 + nq0 where,


2. Acid Rain Ecosystem (pH < 5.5) a. Microspora: 1 culture for 35 studentsb. Peridinium: 1 MAGNAculture 100

3. Organic Polluted Ecosystem a. Euglena: 2 MAGNAcultures 100b. Oscillatoria: 1 MAGNAculture 100c. Chlorella: 1 MAGNAculture 100d. Cyclops: 1 culture for 35 studentse. Daphnia (small): 1/2 culture for 35 students

Aquatic Microecosystem Analysis

The purpose of this activity is to allow students to “discover” that (1) normal, healthy ecosystems have the greatestdiversity of organisms, (2) few organisms can tolerate strongly acidic ecosystems, and (3) organic polluted ecosystemshave little diversity with a few species in great abundance.

Students should work in small groups and share the data. Then, data from all groups should be pooled to obtain bestresults. If time is short, have students sample from the bottom of the ecosystems only. Most organisms will be found there.

Before the analysis begins, students must be reasonably familiar with the organisms that they will encounter. Thisis best accomplished by setting up slides of the organisms under demonstration microscopes so that students can corre-late name and appearance with Figures 38.5 and 38.6. The small size of most organisms makes identification difficultunless students can refer to the demonstration setups. Advise students to ignore organisms other than those in Figures38.5 and 38.6.

Environmental Pollution

The most difficult part of this experiment is the capture of the Daphnia. Cutting off the tip of plastic droppers,usually sent along by biological supply houses, to provide a larger opening makes the job easier. To prevent air fromgetting under the carapace, ultimately killing the Daphnia, submerge the tip of the dropper in the test solution beforesqueezing the bulb to force Daphnia out of the dropper. The dropper must then be rinsed before collecting moreDaphnia. Flat-bottomed vials are preferred over test tubes because the rounded bottom of test tubes reflect light, caus-ing Daphnia to move toward the bottom of test tubes. Students may need assistance in graphing their results and in de-termining the for each pollutant from the graph.

Exercise 40. Population Growth

Students have no major difficulties with this exercise, but some may have difficulties plotting the curves and in de-termining the data for the growth rate curves. They are vitally interested in the problems associated with an excessivehuman population. A portion of the laboratory session should be spent discussing the problems, and possible solutions,associated with an expanding human population. The difficulties of curtailing the human population in developingcountries will become readily apparent. If this topic is covered in lecture only, this exercise may be assigned as home-work to emphasize your presentation.

Exercise 41. Animal Behavior

This exercise focuses on innate behavior in flatworms, sow bugs, and Drosophila flies. It requires that studentsconstruct simple experiments for testing the hypotheses. Students enjoy the exercise, and it usually presents no prob-lems. Tabulating the results for the entire class provides better data for drawing conclusions.

LC50

1pH 7 7.02

L a b o r a t o r y R e p o r t s

Documents

LABORATORY SET-UP AND OPERATIONAL …testbankcart.com/wp-content/uploads/2017/01/Downloadable-Solution...Laboratory Set-Up and Operational Suggestions 9 ... and the nature of scientific