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Using the Scientific Method to Design a Clinical Research Study

Lab Manual Exercises: None Learning Outcomes:

1) Formulate a research question that can be answered with materials available. 2) Form a hypothesis. 3) Design a clinical research study to test the hypothesis. It should contain both an

experimental and control group. 4) Identify independent and dependent variables. 5) Analyze data and graph appropriately. 6) Form a conclusion based on the data. 7) Discuss importance of sample size and sources of experimental error.

Materials:

Treadmills (1) Computer w/excel spreadsheet

Step-boxes (2) Printer

Automatic Blood Pressure Cuffs w/ batteries (4 AA each) (5)

Blank Data Tables (10)

Stopwatches (10) Metronomes (2)

Calculators (10)

Graph Paper (20)

Suggested Reading: Scientific Method Supplement

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ZOOL 141L: # 2 Scientific Method

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LAB # 2 Scientific Method

Background: In today’s lab you will be designing a scientific study to answer a particular research question. You will use this data to write a lab report following the rubric you received on the first day of class. Objectives:

Step 1. : Formulate a research question: (Introduction) Q: Step 2: What is your Hypothesis? (Introduction) H1 = H0 = Step 3: Design a research study to test the above hypotheses. (Materials & Methods)

What is/are the independent variables?

What are the dependent variables? (What types of data will you collect?)

How do we decide who goes in the treatment and control groups?

What treatment will the experimental group receive?

What will the control group receive?

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When and how will measurements be made?

Design a data table to record data from the experiment. You may want to collect data besides just independent and dependent variables.

What precautions have you taken to minimize bias? Step 4: Conduct the experiment (Methods)

Record data in table

Step 5: Analyze the Data (Results)

Present the data in an easy-to-interpret graph or table Discuss trends

Step 6: Form a Conclusion

State whether or not the results support the hypothesis Discuss sources of error Contrast with previous studies Suggest new hypotheses

Lab Notebook Checklist: Lab 2 Scientific Method (Complete Write-up) � Title � Introduction � Materials & Methods � Results (Including 2 graphs) � Conclusions

Introduction to Microscopy and Cells

Lab Manual Exercises: Exercise 3 (Activities 1, 2, 3 & 5) Exercise 4 (Activities 1-6) Learning Outcomes:

1. Learn how to use a compound microscope to examine animal cells. 2. Calculate total magnification (TM) and Field Diameter (FD). 3. Estimate size of objects in a field. 4. Learn how to make a wet-mount. 5. Identify and describe function of major organelles using a cell model. 6. Identify phases of mitosis. 7. Remove tissues from fetal pig for future sectioning.

Materials:

Compound Microscopes (10) Microscope Accessory Boxes (10)

Letter “e” slide (10) Stage Micrometer (1)

Toothpicks (Box) Biohazzard Box (4)

Disinfectant Solution Lugol’s Stain (Iodine)

Brewers Yeast (1 packet) Immersion Oil (5)

Mitosis Cell Models (8) Sperm Smear Slide (Reproductive) (10)

Blood Slide (Cardiovascular) (10) Smooth Muscle Slide, Teased (10)

Ciliated Columnar Epithelium (10) Animal or Onion Mitosis Slides (10)

Swimming Pool Noodles (4- 2 of each color)

Pond Water with living organisms (500 ml)

Digital Projector Fetal Pig or Cat (1- can be cut up)

Cell Model w/organelles (1) Waterproof paper

Dissecting Implements (4 sets) 5 ml Sample Tubes w/ 70% EtOH (10)

Suggested Reading: Exercises 1, 3, & 4

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ZOOL 141L: # 3 Microscopy and Cells

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Assigned Tasks: (Lab Manual) Exercise 3: The Microscope I. Activity 1: Identifying the parts of a microscope II. Activity 2: Viewing objects through the microscope III. Activity 3: Estimating the diameter of the microscope field IV. Activity 5: Preparing a wet mount (Complete above activities as directed in lab manual, but record your data in lab notebook. Omit activities 4, 6, & 7) Exercise 4: The Cell- Anatomy and Cell Division V. Activities 1-4: Identify cell organelles using cell model, sketch and label in lab

notebook, list major functions for each. VI. Activity 5: Observing various cell structures. Observe 4 cell types as directed in lab

manual. Make sketches of each in your lab notebook listing TM and estimated cell size. Record answers to questions (p 36 ) in your lab notebook.

VII. Activity 6: Identifying the mitotic stages. Obtain a prepared slide of mitotic stages. Locate and sketch cells in each of the 6 cell phases (Interphase, Prophase, Metaphase, Anaphase, Telophase, & Cytokinesis). Include TM and label pertinent structures of each stage (e.g., nuclear membrane, chromatin, chromosome, chromatid, centromere, mitotic spindle, cleavage furrow, metaphase plate, kinetechore microtubules). Write a brief description for each stage and be able to recognize on future quizzes and lab practicums.

Supplemental Exercise VIII. Use dissecting tools to obtain a tissue biopsy from the fetal pig. (Your instructor will

tell you which organs(s) to sample). Note the orientation (e.g., transverse or longitudinal) of the tissue as you remove it. Place the tissue in a vial of ethanol and label appropriately. This sample will be examined in lab # 5.

Homework: Using your text, find out which types of tissues may be present in your sample. Write a brief description of the anatomy and function of each tissue type and bring this to lab # 5 (Tissues). Your summary will be counted towards your quiz grade for lab 5.

Instructors Note: This week’s lab focuses on the proper use of the microscope as well as the identification of cellular organelles and mitotic stages. A complete write-up is not necessary.

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TM = 400x

Measuring objects with your compound microscope

Some of the compound microscopes used in A&P lab are outfitted with ocular micrometers (a small ruler found in the eyepiece). The ocular micrometer can be used to measure objects on a slide (e.g., cells, blood vessels, etc). The diameter of a cell can be expressed in ocular micrometer units (OMU’s for short). For example, the cell pictured above is 30 OMUs in diameter. The value of an omu varies depending on magnification. The absolute value can be determined by multiplying the OMU value by the appropriate value for each magnification (see table below). For example, assuming the picture above was taken at 400X, the diameter of the cell would be 30 x 1.25= 37.5 microns. A micron is 1/1000th of a millimeter and is often abbreviated as µm.

TM 1 omu = ? µm

40X 12.5 100X 5.0 400X 1.25

1000X 0.5

Cell Transport Mechanisms: Osmosis and Diffusion

Lab Manual Exercises: Exercise 5A (1,3,5) Learning Outcomes:

1) Define differential permeability and explain differences between active and passive transport 2) Define diffusion, osmosis, solute, solution, isotonic, hypotonic, & hypertonic. 3) Understand the factors that influence the movement of molecules across a semi-permeable membrane 4) Calculate the formula weight of a compound 5) Predict which way substance will move across a membrane given information on concentration gradients. 6) Perform indicated exercises covering osmosis and diffusion.

Materials:

Petri Dishes w/ Agar (no nutrients) (10) 250 ml Beakers (28)

Graph Paper (20) 25 ml graduated cylinder (7)

Wax Marking Pencils (10) Dialysis Tubing cut into 4” (30)

Stopwatches (10) Dailysis Tubing Clamps (all)

Calculators (10) 40% Glucose Solution (2 L)

Rulers (10) Distilled Water (4L)

3.5% Methylene Blue Solution in Dropper Bottle (5)

10 % NaCl Solution (250 ml)

1.6% Potassium Permanganate in Dropper Bottle (5)

Silver Nitrate Solution (5)

Disposable Pipettes (Box) Benedicts Solution (5)

Small Funnels (7) Small Test Tubes (28) and racks (7)

Hot Plates and Balances (4-5 of each) Test tube holders (7)

Hard-boiled eggs (15) or fresh potatoes (3) 40% Sucrose Solution w/ Congo Red (250 ml)

400 ml Beakers (14) 40% Sucrose Solution (2 L)

Gloves & Safety Goggles (22 pairs) Lab Tape (2 rolls)

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ZOOL 141L: # 4 Osmosis and Diffusion

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Suggested Reading: Exercise 5A

Vocabulary & Reading Comprehension Define the following: Solute Solution Concentration Gradient Hypotonic/Isotonic/Hypertonic Diffusion

Osmosis Active Transport Passive Transport Selective Permeability

List 3 factors that influence diffusion rate: 1) 2) 3) Chemical Tests: What compounds do the following chemical tests detect? List what a positive and negative test would look like (e.g., color change) Purpose? Positive Negative 1) Benedict’s test 2) Silver Nitrate (AgNO3)


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Activity 1: Observing diffusion of dye through agar In this part of the lab, you will be monitoring the rate of diffusion of potassium permanganate (KMnO4 a purple-brown dye) and methylene blue (C16H18ClN3S a blue dye) though agar gel. 1) Calculate the molecular mass of 1 molecule of each dye particle. The molecular mass

can be determined by adding up the mass of all atoms making up a compound. For example, the molecular mass of water (H20) is approximately 18 amu. Hydrogen has a atomic mass of approximately 1 amu whereas oxygen has an atomic mass of approximately 16 amu, so molecular mass of water is 2(1) + 1(16) = 18 amu.

2) Given the molecular mass of each dye particle, form a hypothesis as to which of the two dye particles will diffuse most quickly through the agar gel.

3) Perform an experiment to test your hypothesis following the directions for activity 1 (p. 55). It is not necessary to create a well in the agar, as we will be using dry dye for this lab.

4) Record the distance traveled by the dye field in 15 minute intervals. Measure distance traveled from a reference point on the remaining pile of dry dye (see below)

5) Graph the total distance traveled by each dye after 60 minutes 6) Calculate the average rate of diffusion for each dye. Include this information below

your graph. 7) Conclusion: Did the results of your experiment support your hypothesis? Why or why

not? Can you think of any sources of experimental error?


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Activity 3: Observing diffusion through non-living membranes In this part of the lab, we will be determining which molecules can cross a semi-permeable membrane (in this case, dialysis tubing).

1) Read carefully through the directions for activity 3 (p 56). Set up 4 beakers with the required solutions.

2) Fill 4 dialysis tubes with the required solutions, rinse, blot dry and weigh on a laboratory balance. Record this data in your lab notebook.

3) Place the dialysis tubes in the appropriate beakers and leave for 1 hour. While you are waiting, answer the following questions and include in the introduction/ hypothesis for this activity in your lab notebook: i. Include a sketch of the pictured setup in your lab notebook. Which beaker(s)

contain solutions that are isotonic relative to the solution in the bag? Hypertonic? Hypotonic?

ii. Which bags do you predict will change in weight? Why? iii. Which beakers will test positive for glucose? Salt?

4) Remove each sac, blot dry and re-weigh. Record these results in your lab notebook. (You may cut out table from p 56 of the lab manual)

5) Test assigned beakers/ sacs for the presence of glucose using benedicts test or NaCl using silver nitrate. Record these in the table.

6) Conclusion: Did the results of your experiment support your hypothesis? Why or why not? Can you think of any sources of experimental error? As part of your conclusion, you should answer all questions found on p 57 in your lab manual.


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Activity 5: Investigating diffusion through living membranes (Experiment 1) 1) Obtain 2 deshelled hard-boiled eggs 2) Blot dry and weigh. Record the initial weights in your lab notebook (you may cut or

photocopy the table from p. 58 of the lab manual) 3) Carefully place the eggs in the appropriate solutions (see figure below). 4) While you are waiting, form a hypothesis as to how the weight of each egg will change

during the experiment. In constructing your hypothesis, you should state whether the egg/solution pairs are isotonic, hypertonic, or hypotonic.

5) Re-weigh at intervals of 20, 40, and 60 minutes. Record these data in your lab notebook. 6) Conclusion: Did the results of your experiment support your hypothesis? Why or why

not? Can you think of any sources of experimental error?

Lab Notebook Checklist: Lab 4 Transport Mechanisms and Cell Permeability (Lab Report Required) � Title � General Introduction (What was the overall purpose of today’s lab?) � Vocabulary and Reading Comprehension Activity 1: Dye diffusion through agar � Molecular Mass estimates � Results: Graph and Diffusion Rates � Conclusion Activity 3: Diffusion through non‐living membranes � Introduction/ Hypothesis � Sketch of experimental setup � Data table from p 56 � Conclusions Activity 5: Diffusion through living membranes � Hypothesis � Data table from p 58 � C l i

Histology: Classification of Tissues

Lab Manual Exercises: Exercise 6A (Activities 1-4) Learning Outcomes:

1. Demonstrate proficiency using the microscope. 2. Identify and classify epithelial tissue based on cell shape and degree of layering. 3. Identify and classify connective tissues based on ground substance, fibers, and cells. 4. Identify nervous and muscle tissues. 5. Section and stain mammalian tissue using standard microtechnique. 6. Discuss how sectioning plane affects tissue histology.

Materials:

Compound Microscopes (10) & Kits Fibrous CT & Adipose Slides (Tray)

Digital Projector Jejunum or Ileum Slide (Digestive- G.I. Tract) (10)

Epithelium Slides (All trays) Muscle “3 types” Slide (Muscular) (10)

Skin Slides (Tray) Bladder Slide (Urinary) (5)

Cartilage Slides (Tray) Microscope Camera (2)

Blood Slide (Circulatory) (10) Digital Camera w/batteries (2 AA)

Spinal Cord Giant Multi-polar Neuron slide (Nervous) (10)

Toluidine Blue (2.5% Solution) (20 ml)

Spongy Bone Slide (Skeletal) (10) Tissues from Lab # 3

Compact Bone Slide (Skeletal) (10) JB4 Resin Kit

Microtome w/ light source Glass knives (10)

Jewelers Forceps (2) Dessicator

Hotplate (1) Insulin Syringes (5)

Functional Histology Book (2) DipQuick Stains 2 & 3

Suggested Reading: Exercise 1 & 6A

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ZOOL 141L: #5 Tissues

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Exercise 6A: Classification of Tissues I. Activity 1: Examining Epithelial Tissue Under the Microscope Examine slides of the epithelial tissues listed on the board. Make a drawing of each in your lab notebook. For each, list total magnification (TM), classify as simple or stratified, and identify apical cells by cell shape (squamous, cuboidal, or columnar). Label basal and apical surfaces and basement membrane. For each tissue, list the function (absorption, secretion, or protection) and indicate the organ or area of the body in which the tissue is found. Epithelial tissues: 1. Simple Cuboidal (Kidney) 2. Simple Columnar (Intestine) 3. Simple Squamous (Lung)

4. Stratified Squamous (Skin) 5. Pseudostratified (Trachea)

II. Activities 2- 4: Examining connective, muscle, and nervous tissues under the

microscope Examine slides of the connective, muscle, and nervous tissues listed on the board. Make a drawing of each in your lab notebook. For each, list total magnification (TM). Label visible cells and fibers. For each tissue, list the function and indicate the organ or area of the body in which the tissue is found. 1. Hyaline Cartilage (Trachea) 2. Fibro cartilage (Ear) 3. Areolar Connective Tissue 4. Dense Regular Connective Tissue

(Ligament or Tendon) 5. Dense Irregular Connective Tissue

(Dermis of Skin)

6. Adipose (Skin, Hypodermis) 7. Spongy Bone 8. Compact Bone 9. Blood 10. Motor Neuron (teased or from spinal

cord)

11. Muscle Tissue

III. Supplemental Activity: Sectioning tissue using standard microtechnique In this activity, you will make a permanent mount of tissues you collected from the fetal pig two weeks ago. The samples have been embedded in a water-based resin. If the embedding process worked successfully, the tissue and surrounding resin should be of equal hardness. This will allow you to cut the tissue into very fine sections so that they can be examined under the microscope. The cutting machine (called a microtome) is outfitted with a glass or metal cutting blade. The blade is very sharp and easily damaged. Make sure that you do not touch the blade. SAFETY GOGGLES AND GLOVES ARE MANDATORY!

ZOOL 141L: # 5 Tissues

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1) Remove your tissue “Block” from the embedding capsule by slitting the capsule length-wise with a razorblade. If the block is moist, blot it dry with a paper towel.

2) Trim the conical portion of the block with small cuts from your razorblade until the tissue is exposed.

3) Place your block into the holder (chuck) on the microtome and tighten by turning the bezel clockwise.

4) Insert a new glass knife into the knife holder.

5) Adjust the position of the knife using coarse and fine adjustment so that the block nearly makes contact with the knife when the microtome wheel is rotated.

6) Make sequential sections through the block by turning the microtome wheel. In between each turn, tighten the fine adjustment screw approximately 1/16th turn. Gently pluck sections off the blade using a pair of jeweler’s forceps (BE SURE NOT TO TOUCH THE BLADE WITH THE FORCEPS). Discard inferior sections.

7) Once you obtain a satisfactory section, lift it off the knife and suspend it on a drop of distilled water placed on a microscope slide. This takes some finesse, as the section will stick to your forceps once it is wet.

8) Dry the slide on a hotplate (heat setting 2-3) until the water droplet evaporates.

9) Label the slide with a archival pen.

10) Stain your sample as directed by your instructor.

11) Photograph or draw your section, including TM or scale bar. Include this in your lab notebook.

12) Identify and label all tissues present and list the function(s) of each.

Histology Tutorial (Activity 6B p-PEx-21in the back of your manual) Use the histology tutorial on your PhysioEx CD (located in back of lab manual) to review tissues examined in this weeks lab. Instructors Note: This week’s lab focuses on the identification and classification of tissues. We will not be conducting any experiments per-se, so it is not necessary to include a research question or hypothesis in your lab write-up.

Integumentary System and Body Fat Estimation

Lab Manual Exercises: Exercise 7 (Activities 1,3,4,5,6) Learning Outcomes:

1) Identify the major and minor layers of skin from slides and models. 2) Know which tissues are found in each of the above layers. 3) Compare and contrast the presence of nervous tissue and blood vessels in each layer. 4) Describe the origin and function of skin derivatives (sebaceous glands, etc) 5) Compare distribution of skin glands on different parts of the body. 6) Classify fingerprint patterns and discuss functions of fingerprints. 7) Measure body fat using various methods.

Materials:

Skin Models (all) Treadmills (1)

Skin Poster Stopwatches (10)

Skin Slides (all) Compound Microscopes and kits (10)

Fingerprint Kits (4) Lab Tape (4 rolls)

Print Elimination Cards (30) 0.5 cm squares of # 20 bond paper (30)

Fingerprint Ink Pad (4) Lugol’s Solution (4)

Alcohol pads (box) Skin Calipers (all)

Dissecting Scopes w/overhead illum (2) Body Fat Scales (2) (each takes 4 AA batteries)

Goniometers (all) Surface Anatomy Chart

Hand Tally-Counters (4) 3 x 5 note cards (20)

Omron Fat Loss Monitor (+ 2 AAA) (1)

Suggested Reading: Exercise 7 & Body Fat Supplement

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ZOOL141L: #6 Skin and Body Fat

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Activity 1: Locating structures on a skin model

1) Obtain a 3D skin model from your instructor 2) Photograph or make a drawing of the model in your lab notebook 3) Label the following structures AND list tissues present and their functions

Epidermis Dermis Hypodermis Hair Follicle

Sebaceous Gland Eccrine Sweat Gland Arrector Pili Muscle Dermal Papillae

Pacinian Corpuscle Blood Vessels Free Nerve Ending

Activity 3: Comparison of hairy & hairless skin

1) Obtain a prepared slide of hairy (scalp) and hairless (sole of foot) skin. 2) Make a drawing of each at an appropriate magnification. You should be able to see all

layers of the epidermis and dermis. 3) Label the following layers:

Stratum Lucidum Stratum Granulosum Stratum Spinosum

Stratum Spinale Stratum Corneum Dermis

4) Question: In which way (besides the presence or absence of hair) does “hairy” skin differ

from “hairless” skin? Activity 5: Plotting the distribution of sweat glands. In this activity, we will compare the density of sweat glands on thick (palms) and thin skin (forearm) by doing a short in-class experiment.

1) Form a hypothesis as to which area will have more sweat glands. 2) Select a minimum of 10 subjects for the experiment. MAKE SURE THEY ARE NOT

ALLERGIC TO IODINE! 3) Paint both skin regions with iodine and dab dry repeatedly with fresh paper towels until

no iodine appears on the towel. 4) Securely tape 0.5 cm2 of parchment paper over each skin area. 5) Have the participant engage in strenuous exercise (e.g. treadmill or bicycle) for 2-5

minutes. 6) Afterwards, have the student sit quietly for an additional 5 minutes. 7) Remove the papers and tape, blue-side up, to a 3x5 note card. Label with student name

and region of skin. 8) Count the number of blue dots per each square. Multiply each by 2 to obtain # of sweat

glands per cm2. (Discard any samples which are over-stained or un-readable) 9) Return the 3x5 card to the donor student so that it may be included in their lab notebook. 10) Construct a bar graph comparing average density of sweat glands in each area. 11) Conclusion: Write a short paragraph summarizing your results and stating whether or not

your hypothesis was supported. Discus any other factors which may have affected your results.


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Activity 6: Identifying Fingerprints

1) Working in groups of 3-4, obtain reference fingerprints for each person in your group (left hand only) following the directions on p. 100.

2) Each student should classify the pattern of each fingerprint pattern using Figure 7.8 (p. 99). Write the pattern type next to each fingerprint and include in your lab notebook.

3) After washing the ink off your hands, run your left hand through your hair and place your left index finger on a microscope slide to create a latent print

4) Have someone in your group mix the slides around while your back is turned. 5) Dust the slides for fingerprints. 6) Match each slide with the appropriate reference prints.

Supplemental Activity: Using skin-fold thickness to estimate body fat percent. Although not technically part of the skin, the hypodermis (also known as the sub-cutaneous layer) is closely associated with the skin. The hypodermis contains large amounts of adipose tissue which helps insulate the body and store excess calories. As the amount of body fat increases, the thickness of the hypodermis increases. As such, skin-fold measurements can be used to estimate body fat percentage. In this lab, we will compare the body fat percentage of males and females using skin fold measurements, BMI (Body-Mass-Index) and electrical impedance scales. None of these methods is without bias, so it is important not to read too much into any one measurement.

1) Form a Hypothesis as to which sex will have more body fat. Record in your lab notebook.

2) Have each student weigh and measure themselves using the scale at the front of the record in cm and kg (not inches & lbs!). Do NOT record student name.

3) Calculate BMI using the attached chart and record (Note that BMI does not correspond directly to % body fat!!).

4) Use calipers to measure the skin fold thicknesses in the following 3 areas: Females: Triceps, Suprailium & Thigh Males: Chest, Abdominal & Thigh

5) Record the summed data in the attached data sheet. Use appropriate table to estimate % body fat for females (Table 1) and males (Table 2) (p. 11 of handout)

6) Use the impedance scale or fat-loss monitor to estimate your body fat based on electrical conduction. Record in the data sheet.

7) (Optional): Use goniometers to measure diameter of elbow, knee, biceps & calf. 8) Enter all data in classroom computer. Be sure you have a copy of the class data before

leaving lab! 9) Analysis:

a) Construct a bar graph comparing body fat by gender (use skin fold %) b) Construct a scatter-plot comparing body fat by age (use skin fold %)


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c) Construct a bar graph comparing average body fat for the class using skin fold and electrical impedance measurements.

10) Conclusion: Write a conclusion addressing the following. a) Which sex has a greater body fat %? b) Was your hypothesis supported or refuted? c) Why would one sex have more body fat than the other? (Is there any evolutionary

advantage?) d) Was there any relationship between age and body fat percentage? e) Which of the three indices is most accurate? Least accurate? (Explain your answer) f) If you could re-do your experiment, how would you change the methods or

experimental design?

.

Lab Notebook Checklist: Lab #6 Skin and Body Fat Lab Report Required � Title � Labeled drawing of skin model � Labeled drawing of Thick vs. Thin skin + question � Sweat gland hypothesis � Sweat gland graph � Sweat gland conclusion � Reference Fingerprints w/ patterns labled � Body fat hypothesis � Datasheet � Graph: Body Fat vs. Gender � Graph: Body Fat vs. Age � Graph: Impedance vs. Skin fold � Conclusion


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Bone Tissue and Skeletal System

Lab Manual Exercises: Exercise 9 (Activities 1, 4, & Review) Exercise 10 (Activities 1,3,4 & Review) Exercise 11 (Activities 1-6 & Review) Learning Outcomes:

1. List the five functions of the skeletal system. 2. Identify four main groups of bones. 3. Distinguish spongy bone from compact bone. 4. Describe how bone grows and repairs itself. 5. Identify locations and functions of skeletal cartilages. 6. Identify all bones comprising the axial and appendicular skeleton. 7. Identify required landmarks on the above bones. 8. Articulate bones into a skeleton.

Materials:

Fresh long bone, sawed longitudinally (1)

Skeletal System Poster

Digital Projector Human Skeletons, articulated (2-3)

A&P Revealed CD’s (10) Disarticulated half skeletons (4)

Laptops (10) Skull Models (4)

Compound Microscopes and Kits (10) Disarticulated Skulls (pink boxes) (3)

Spongy Bone Slide (Skeletal)(10) Exploded Skull

Ground Bone Slide (Skeletal) (10) Loose Human Bones

Compact Bone Slide (Skeletal) (10) Cat Skeletons (2)

Osteogenesis Endochondrial Ossification (Skeletal) (5)

X-rays of Human Bones

Suggested Reading:

ZOOL 141L

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Exercise 9: Skeleton Overview Activity 1: Gross Anatomy of a Long Bone (pp. 114). Examine a sectioned long bone. Sketch in your lab notebook. Label all boldface terms. Activity 3: Examining Microscopic Structure of Compact Bone (p 115) Obtain a prepared slide of ground bone and examine it under low power microscope. Identify the following structures: a. Central Canal b. Osteon c. Lamellae

d. Canaliculi e. Lacunae

Activity 4: Cartilages of the Skeleton (p. 118) Identify the following cartilages on the human skeleton. a. Hyaline Cartilage (Joint Surfaces and Costal Cartilages) b. Fibrocartilage (Intervertebral Discs) c. Elastic Cartilage

Skeletal Overview Review Sheet: Complete review sheet on pp. 119-122 and include in your lab notebook.

Exercise # 10: The Axial Skeleton

Activities 1-4: Identify the bones of the Axial Skeleton using Anatomy and Physiology Revealed ® CD as well as the articulated and disarticulated human skeletons. You should be able to identify ALL required bones, processes, and depressions listed on the accompanying table. Axial Skeleton Review Sheet: Complete review sheet on pp. 139-144 and include in your lab notebook.

Exercise #11: The Appendicular Skeleton

Activities 1-5: Identify the bones of the Appendicular Skeleton using Anatomy and Physiology Revealed ® CD as well as the articulated and disarticulated human skeletons. You should be able to identify ALL required bones, processes, and depressions listed on the accompanying table. Appendicular Skeleton Review Sheet: Complete review sheet on pp. 157-163 and include in your lab notebook. Instructors Note: This week’s lab focuses on the identification and classification of bones as well as bone foramina and processes. A full write-up is NOT required. However, be aware that bone identification and anatomy will comprise 30-50% of the questions on the first lab practicum.

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Category Bone Markings & Processes Depressions and OpeningsCranial Bones Frontal Frontal Sinus

Parietal Sagittal SutureTemporal Zygomatic Process External Accoustic Meatus

Mastoid Process Carotid Foramen/CanalStyloid ProcessSquamous Part

Occipital Lamboid Suture Foramen MagnumExternal Occipital ProtuberanceOccipital Condyle

Sphenoid Greater Wing Foramen OvaleLesser Wing Sella Turcica

Optic ForamenEthmoid Perpendicular Plate

Facial Bones NasalMaxillae Infraorbital ForamenZygomatic Temporal ProcessLacrimalPalatine Hard PalateVomerMandible Articular Tubercle Mental Foramen

Coranoid Process Mandibular ForamenHyoid HyoidVertebrae Vertebrae (All) Dens Vertebral Foramen

Atlas Intervertebral ForamenAxis Sacral HiatusBody Sacral CanalSpinous ProcessTransverse ProcessLaminaPedicleFacetIntervertebral DiscCervical, Lumbar, Thoracic

Sternum & Ribs Sternum ManubriumBodyXiphoid

Ribs Costal CartilagesVertebrosternal RibVertebrocostal RibFloating Rib

Required Bones and Processes

Axial Skeleton

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Category Bone Markings & Processes Depressions and OpeningsPectoral Girdle Clavicle Medial End & Acronial End Scapula Coracoid Process Glenoid Cavity

Acromion Subscapular FossaSpine Supraspinous FossaSuperior Angle Infraspinous FossaInferior Angle

Humerus Head Olecranon FossaGreater Tubercle Coranoid FossaLesser TubercleDeltoid TuberosityLateral EpicondyleMedial EpicondyleTrochleaCapitulum

Radius HeadRadial TuberosityStyloid Process

Ulna OlecranonHand Bones Trapezoid

TrapeziumScaphoidLunateCapitateHamateTriquetrumPisiformMetacarpalsPhalanges

Pelvic Girdle Ilium Iliac CrestAcetabulumSacroiliac Joint

Ischium Ischial SpineIschial Tuberosity

Pubis Pubic Symphysis Obturator ForamenFemur Greater Trochanter Intercondylar Fossa

Lesser TrochanterLateral CondyleMedial CondyleHeadGluteal TuberosityNeck

Tibia Medial Condyle Fibular NotchLateral CondyleTibial TuberosityMedial MalleolusAnterior Border

Fibula HeadLateral Malleolus

Foot CalcaneusTalusCuboidLateral CuneiformMedial CuneiformIntermediate CuneiformNavicularMetatarsalsPhalanges Proximal, Medial, Distal

Appendicular Skeleton

Required Bones and Processes

Joint Articulations & Body Movements

Lab Manual Exercises: Exercise 13 (Activities 1-3, 5-6) Learning Outcomes:

1. Name and describe functional and structural classifications of joints. 2. Identify types of synovial joints. 3. Review major bone landmarks. 4. Examine the effects of gender and exercise on range-of-motion (ROM).

Materials:

Human Skeletons, articualted (2-3) Skeletal System Poster

Disarticulated half skeletons (4) Joint Models (all)

Loose Human Bones Step Boxes (2)

Cat Skeletons (2) Bicycle (1)

Metronomes (2) Stopwatches (2)

Exercise Mats (4) Joint ROM Goniometers (all)

Skull Models Duct Tape (Roll)

Measuring Tapes (3) Treadmill (1)

Meter stick (2)

Suggested Reading:

ZOOL 141L

9

Joint Articulations & Body Movements (Exercise 13) Activities 1-2: Identifying different types of joints Activity 5: Demonstrating Joint Movements Activity 6: Demonstrating Uniaxial, Biaxial and Multiaxial Movements Research Questions 1& 2:

Q1) Which synovial joint will have greatest Range of Motion (ROM): Neck, elbow, shoulder or hip? H1) Q2) Which sex is more flexible- males or females? H2)

Methods 1) Measure the ROM for each of the indicated joints.

2) Graph the AVERAGE ROM by gender for each of the four synovial joints.

3) Write a brief conclusion stating a) which joint was more flexible and b) which gender was more flexible.

Research Question 3 Q3) What effect will exercise have on flexibility? H3)

Methods

1) Measure toe-distance before exercising

2) Exercise 2 minutes on bicycle

3) Re-measure toe-touch distance

4) Graph AVERAGE toe-touch distance before and after exercise.

5) Conclusion: Write a brief conclusion stating whether or not your hypothesis was supported and describing any sources of error

ID

Gender

HT

Age

Neck

Elbow

Shoulder

Hip

Before Exercise

After Exercise

Ran

ge of Motion (degrees)

Toe Touch (cm

)

Muscle Tissue and Gross Anatomy of Muscular System

Lab Manual Exercises: Exercise 14 (Activities 1-3) Exercise 15 (Activities 1-6) Learning Outcomes:

1. Distinguish three types of muscle based on histological structure and function. 2. Prepare section of fresh muscle tissue using standard microtechnique. 3. Distinguish between slow-twitch and fast-twitch muscle. 4. Identify major muscles of the human body. 5. Name the action, origin, and insertion for the above muscles.

Materials:

Chicken Breast & Thigh (Fresh) (1) (Keep on ice)

Human Skeletons (2)

0.9% Saline Solution Muscle Models (full body, hand, shoulder, leg. foot, arm)

Compound Microscopes and kits (10) Pool Noodles (2)

Microtome & JB4 Resin (optional) Dissecting Instruments (10)

Muscular System Slides (all) Microscope w/ digital camera

Laptops (10) & A& P Revealed CDs Skeletal Muscle Fiber Model

Dip-Quick Stains # 2 & 3 Dissectible Human Torso (2)

Razor blades Slides- Neuromuscular Junction (10)

Suggested Reading: Exercises 14 & 15

ZOOL 141L

11

EXERCISE 14: Microscopic Organization of Muscle Tissue Activity 1: Examining the anatomy of fresh skeletal muscle

1) Prepare fresh sections of chicken breast and chicken thigh muscle using the microtome or razor blade.

2) Mount sections on separate microscope slides. 3) Stain as directed by your instructor. 4) Draw both muscle types and label. 5) Q: how does the appearance and structure of the two muscle types differ? (why is one

“dark” and the other “light”? Which muscle sample would be best for sustained aerobic exercise? Why?

Activity 2: Histological structure of skeletal muscle 1) Obtain a slide showing cross section of skeletal muscle tissue. 2) Draw or photograph at 400X. 3) Label nuclei, endomysium, perimysium & muscle fibers

Activity 3: Structure of the Neuromuscular junction

1) Obtain a specially –stained slide of a motor unit. 2) Sketch at 100x, labeling the following: muscle fibers, axon, terminal branches 3) Q: What is a motor unit? What type of neurons control skeletal muscle?

EXERCISE 15: Gross Anatomy of the Muscular System (Activities 1-6)

1) Use anatomical models, A&P revealed CDs and text illustrations to identify required muscles (see attached list).

2) Fill-in origins & insertions & MAJOR actions. Although you are not required to include any drawings in your lab manual, please be aware that muscle identification will constitute approximately 30%-40% of second practicum.

3) Complete review sheet # 15 and include in your lab manual.

Muscle Origin Insertion Actions

Temporalis Closes jaw (elevates mandible)

Masseter Closes jaw (elevates mandible)

Platysma Pulls lower lip down

Sternocleidomastoid Flexes neck& rotates head

Pectoralis Major adducts, medially rotates arm

Pectoralis Minor pulls scapula forward

Serratus Anterior

Intercostals

Trapezius

Latissimus Dorsi Forceful Extension of Arm

Infraspinatus

Teres Major

Teres Minor

Errector Spinae Extension & lateral flexion of spine

Levator Scapulae

Triceps brachii Extension of forearm

Biceps brachii

Brachialis Flexes forearm

Pronator Teres

Brachoradialis

Flexor digitorum superficialis

Extensor digitorum

Supinator Supinates forearm

Rectus Abdominis

External Oblique

Internal Oblique

Transverse Abdominis

Tensor Fascia Latae Abducts Thigh

Sartorius

Quardiceps Femoris (4 muscles) Extends Knee/Flexes Thigh

Adductor muscles

Psoas Major/ Iliopsoas

Gracilis Adducts Thigh

Gluteus Maximus

Semimembranosus

Semitendonosus

Biceps femoris

Tibialis Anterior Dorsifelxion of toes

Gastrocnemius

Soleus Plantar flexion of toes L

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Ab

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Ant

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Ant

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T

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h

Pos

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Ant

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Lab # 11: Required Muscles

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Ant

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Muscle Physiology and Gross Anatomy of Muscular System

Lab Manual Exercises: Exercise 15 (Activities 1-6) Exercises 3-5 Learning Outcomes:

Materials:

Laptops (10) & A& P Revealed CDs Lab Tape (2 rolls)

Dissected Cat (optional- 1) Gel Electrodes (20)

Human Skeletons (2) Free Weights

Muscle Models (all from previous lab) Electrode Cream and Paste (2)

Computers (PC’s) (2) Metronomes (2)

PowerLab Unit (Summation Setup; Fig 1) Hand Dynamometer (1)

PowerLab Unit (Fatigue Steup; Fig 2) Printer

Rubber Bands (Bag) Packing Tape (Roll)

Alcohol Pads (Box) Skin Abrasive

Goniometers (all)

Suggested Reading:

ZOOL 141L

12

ZOOL 141L: # 12 Muscle Physiology

2

Objectives: In this experiment, you will explore how muscles work. You will also examine some of the properties of muscle fatigue. In this experiment, you will electrically stimulate the nerves in the forearm to demonstrate recruitment, summation, and tetanus.

Assigned Tasks: (Powerlab Experiments) I. Vocabulary & Reading Comprehension II. Exercise 3: Summation and tetanus III. Exercise 4 & 5: Muscle Fatigue Vocabulary & Reading Comprehension Define the following:

Action Potential Twitch Wave Summation

Tetanus Recruitment Fatigue

Background The skeleton provides support and articulation for the body. Bones act as support structures and joints function as pivot points. Skeletal, or striated, muscles are connected to the bones either directly or by tendons, strong bundles of collagen fibers. Two or more muscles usually work antagonistically. In this arrangement, a contraction of one muscle stretches, or elongates, the other. Skeletal muscle is composed of long, multinucleate cells called fibers. These fibers are innervated by motor nerves. An action potential in a motor axon produces an action potential in the muscle fibers it innervates. This muscle action potential allows for a brief increase in the intracellular concentration of calcium ions ([Ca2+]), and activates the contractile molecular machinery inside the fiber. The result is a brief contraction called a twitch. A whole muscle is controlled by the firing of up to hundreds of motor axons. These motor nerves control movement in a variety of ways. One way the nervous system controls a muscle is by adjusting the number of motor axons firing, thus controlling the number of twitching muscle fibers. This process is called recruitment. A second way the nervous system controls a muscle contraction is to vary the frequency of action potentials in the motor axons. At stimulation frequencies of less than 5 Hz, intracellular [Ca2+] returns to normal levels between action potentials: the contraction consists of separate twitches. At stimulation frequencies between 5 and 15 Hz, [Ca2+] in the muscle has only partly recovered when the next action potential arrives. The muscle fiber produces a pulsing tension called a summation response with a force greater than that of a single twitch and that does not decay completely to zero between pulses. At even higher stimulation frequencies, the pulsing component becomes hard to discern and the muscle enters tetanus, a smooth contraction many times stronger than that in a single twitch.


SPB12c Page 3 of 12 18 August 2010

In Exercise 1, you will observe muscular responses without recording them. In Exercises 2 and 3, you will use a force transducer to measure small forces generated by the adductor pollicis muscle. In the later exercises, the grip force exerted by the hand is recorded with a different transducer as you investigate the phenomenon of muscle fatigue.

Required Equipment A computer system Chart software, version 5.0 or later PowerLab 4/20T Finger pulse transducer Bar stimulus electrode Electrode cream Hand dynamometer Adhesive tape



Procedures

Warning Some of these exercises involve application of electrical stimulation to muscle through electrodes placed on the skin. People who have cardiac pacemakers or who suffer from

neurological or cardiac disorders should not volunteer for those exercises. If the volunteer feels major discomfort during the exercises, discontinue the exercise and consult your instructor.

A. Set up and calibration of equipment 1. Make sure the PowerLab is connected and your computer is turned on.

2. Connect the finger pulse transducer to the BNC socket on Input 1 of the PowerLab.

3. Place the finger pulse transducer diaphragm-side up on the top of the lab bench; tape the transducer in place along the Velcro strap (Figure 1).

4. Connect the bar stimulus electrode to the isolated stimulator output of the PowerLab. The leads are color-coded; plug the red lead into the red socket and the black lead into the black socket (Figure 1).

5. Place a small amount of electrode cream on the two silver contacts of the stimulating bar.

6. Turn on the PowerLab.

7. Locate Chart on your computer and launch the program. If the Experiments Gallery dialog box does not appear in front of the Chart window, choose the Experiments Gallery… command from the File menu.

8. In the Experiments Gallery dialog box, select Muscle from the left-hand list. Select the settings file “Nerve Effect Settings” in the right-hand list and click the Open button to apply those settings.

9. After a short time, the Chart window on the computer screen should be set up for the first exercise. Only one channel should be showing, Channel 1, and there should be a red cross through the Record/Monitor button next to the Start button at bottom right of the Chart window.

Figure 1. Equipment set up for the muscle experiments.



Exercise 3: Summation and tetanus

Objectives

In this exercise, you will demonstrate the effects of changing the interval between paired stimulus pulses and observe a short tetanic contraction.

Procedure

1. Close the open document, and choose “Experiments Gallery…” from the File menu. Open the settings file “Summation Settings”. After a short time, the Chart window on the computer screen should be set up, with two channels appearing. Channel 1 should be named ‘Force’ and Channel 2 should be named ‘Stimulus’, as they were in the last exercise.

2. Windows users: From the Setup menu, choose Isolated Stimulator. This will open the Isolated Stimulator window (Mac OS users choose the Stimulator command to open the Stimulator window). Ensure that the settings are the same as in Figure 5.

3. Windows users: Proceed to step 4. Mac users: Close the Isolated Stimulator window, choose the Stimulator Panel command from the Setup menu, open the Isolated Stimulator Panel and then proceed to step 5.

4. Ensure that the volunteer’s hand and stimulus electrodes are placed as shown in Figure 2, and turn the stimulator switch ON.

5. In the Isolated Stimulator Panel, set the pulse current to about 5 mA greater than the maximal stimulus value you determined in Exercise 2.

6. Windows users: Click Start and then immediately click the “Stimulate” button in the Isolated Stimulator window. Chart will record for a fixed duration of 3 seconds, and then stop automatically. Mac OS users: The stimulus will automatically occur 0.5 seconds after the recording begins and Chart will record for 3 seconds. When the recording has stopped, add a comment called “1 Hz” in the new block of data to note the stimulus frequency used.

Figure 5. The Isolated Stimulator window, showing the initial stimulation settings for Exercise 3. A - Windows, B - Mac OS X.

A. B.



7. Increase the stimulus frequency to 2 Hz in the Stimulator Panel, and then click Start. Note the stimulus frequency (2 Hz) in a comment, as in the previous step.

8. Repeat the stimulation for the frequencies 5, 10 and 20 Hz, noting the values in comments as above.

9. Open the Isolated Stimulator window once more, change the number of pulses from 2 to 3, then close the dialog box. Be careful with this setting: a prolonged tetanus with a large number of pulses may be painful for the subject.

10. Click Start. The volunteer should receive a burst of three stimuli at 20 Hz. Add the comment “Tetanic stimuli (3)” to the new block of data. If it is not causing too much discomfort, you could try again with four pulses.

11. Set the stimulus pulse current to 0 mA in the Isolated Stimulator Panel, then close it.

12. From the front of the PowerLab, turn the stimulator switch OFF. Disconnect the finger pulse transducer and stimulus electrodes from the PowerLab.

13. If you are saving your files, choose Save from the File menu to save the recording. Your data should resemble Figure 6.

Figure 6. Typical results from paired stimuli, viewed with 2:1 horizontal compression (the stimulus delay is about a second here).

Exercise 4: Grip force measurement

Objective

In this part of the laboratory you will calibrate the hand dynamometer with respect to a volunteer’s maximal grip strength.



Procedure

1. Remove any transducers and electrodes from the PowerLab, and then connect the plug of the grip force transducer to the Pod Input 1 (Figure 7).

2. Close the open Chart data file. If a warning dialog box appears, click the No button. From the Experiments Gallery, open the settings file “Grip Settings”. After a short time, the Chart View on the computer screen should be set up, with one channel appearing. Channel 1 should be named “Grip”.

3. The volunteer should loosely grip the hand dynamometer in the fist, as shown in Figure 7.

Figure 7. Connections for measuring grip strength and muscle fatigue.

4. Click Start to begin recording. The volunteer should squeeze the dynamometer as hard as possible for a second or two, and then relax their grip. After recording for a few seconds, have the volunteer repeat the maximum grip and then relax. Click the Stop button.

5. Drag over the largest response to select a range of data that includes both the relaxed and maximum-force signals (Figure 8). Choose the Units Conversion… item from the Channel Function pop-up menu for Channel 1.



Figure 8. Selection of trace for calibrating to relative grip strength.

6. In the Units Conversion dialog box (Figure 9) a roughly correct conversion has already been set. Now you need to calibrate correctly for the strength of the volunteer. Select part of the trace where the force was zero, and click the top Point 1 arrow button. Then select part of the trace at the peak, and click the bottom Point 2 arrow button. Click the OK button to return to the Chart View.

Figure 9. The Units Conversion dialog box, with relaxed signal selected.

Exercise 5: Muscle fatigue

Objectives

In this part of the experiment, you will observe the decline in maximal force during a sustained contraction, and to examine some properties of muscular fatigue.

Procedure

The grip force transducer should already be calibrated for the volunteer, as described in Exercise 4. 1. Adjust the scale for Channel 1 (Grip) to show –20 to 120%.



2. Allow the volunteer to view the computer screen. Click the Start button, and ask the volunteer to maintain 20% maximal grip strength while watching the recorded trace (the Range/Amplitude display for Channel 1 shows the percentage force applied).

3. After 20 seconds, tell the volunteer to relax.

4. Click Stop.

5. Wait for 30 seconds to allow recovery of muscle function.

6. Repeat steps 2–4 above for contractions of 40%, 60%, 80% and 100% of maximal grip strength.

7. Allow the volunteer to rest for two minutes.

8. Turn the volunteer away so that they cannot see the computer screen.

9. Click Start, and ask the volunteer to produce a sustained maximal contraction. After 8 to 10 seconds, or when the force has obviously declined, instruct them to try harder. After another 8 to 10 seconds, repeat the encouragement. After a few seconds, ask the volunteer to relax, and click Stop.

Note: Nearly all subjects can produce temporary increases in muscle force during a fatiguing contraction, when sufficiently motivated by verbal encouragement. Click Start and ask the volunteer to produce a sustained maximal contraction. Every 8 to 10 seconds, allow the volunteer to relax very briefly (half a second), and then return to maximal contraction. Click the Stop button to terminate recording after 30 to 40 seconds. Note: Even brief periods of relaxation allow substantial recovery from fatigue, but the recovery is only temporary (Figure 10).

Figure 10. Fatiguing contraction, with brief periods of relaxation.

10. Allow the volunteer to use their other hand if gripping the transducer has become painful. Turn the volunteer so that they can see the computer screen. Click the Start button, and ask the volunteer to produce a 40% contraction while watching the trace. After 10 seconds, press the Enter key to enter a comment (to mark the time).



11. Have the volunteer close their eyes, and attempt to maintain exactly the same contraction force for the next 30 seconds.

12. After the elapsed time, the volunteer should open their eyes, and adjust the contraction force back to 40%.

13. Click Stop and examine the trace.

Note: Almost all subjects will show a declining force (pseudo-fatigue) while their eyes are shut, that is very similar to fatigue. This is, however, not true fatigue, because the full 40% force can be exerted easily, as can be seen when the subject’s eyes are opened again. Analysis

Exercise 3: Summation and tetanus 1. Calculate the stimulus interval in seconds for each stimulation frequency. Enter your data in Table 2

of the Data Notebook.

2. Using the Marker and Waveform Cursor, measure the amplitude of the first two responses at each stimulus interval. Enter your results in Table 2 of the Data Notebook.

3. Examine the tetanic response. Calculate the stimulation interval in seconds and enter your value in Table 3 of the Data Notebook.

4. Drag across the tetanic response to select it, and examine the selected data in the Zoom window. Determine the maximum force amplitude using the Marker and Waveform Cursor and enter your results in Table 3 of the Data Notebook.

5. If you repeated the tetanus experiment with four pulses, repeat your analysis and add it to Table 3. Otherwise, leave the second line blank in Table 3.

Table 2. Results for summation experiment in Exercise 3.

Stimulus frequency (Hz)

Stimulus interval (sec)

Amplitude of first response (mV)

Amplitude of second response (mV)

1

2

5

10

15

20

Table 3. Results from the tetanus experiment in Exercise 3.

Stimulus frequency

Stimulus interval (sec)

Number of pulses

Amplitude of response (mV)



(Hz)

20 3

20 4

20 5

20 10

20 20

Table 4. Results from fatigue experiment

Time to 70% Fatigue

Constant Contraction

Intermittent Contraction

Lab Notebook Checklist: Lab 12 Skeletal Muscle Physiology Lab Report Required! � Title � Vocabulary and Reading Comprehension Exercise 3: Summation and Tetanus (Complete Write‐up) � Introduction � Question � Hypothesis � Results: Table 2 & Graph � Table 3 � Conclusion Exercise 4 & 5: Fatigue (Complete Write‐up) � Introduction � Question � Hypothesis � Results: Table 4 & Graph � Conclusion � Questions for activities 3 & 4



Study Questions

Exercise 3 1) What are the two ways by which the nervous system can control the force generated by a

muscle?

2) Electromyography, with needle electrodes inserted through the skin into a muscle, has been used to study the frequency of muscle fiber activation during voluntary contraction in humans. During weak contractions, the firing frequency is low, so that each fiber produces distinct twitches. The force produced by the whole muscle, however, is relatively smooth. How do you think this occurs?

Exercise 5 Fatigue is not well understood. Some factors that have been proposed to explain the fall in force during fatigue include: changes in the “sense of effort”, loss of “central drive”, failure of neuromuscular propagation, reduction in calcium release in excitation–contraction coupling, metabolic changes in the muscle, and reduction in muscle blood flow owing to compression of blood vessels.

3) Do your experiments help to decide which factors are important?

4) What explanations can you think of for pseudo-fatigue?

Histology of Nervous Tissue

Lab Manual Exercises: Exercise 17: Histology of Nervous Tissue Learning Outcomes:

1) Identify the anatomy of a neuron and Schwann cells 2) Compare and contrast the distribution and function of gray mater and white mater in the

brain and spinal cord. 3) Be able to distinguish between histological sections from different parts of the brain. 4) Identify the Macro Anatomy of the Brain

Materials:

Compound Microscopes and Kits (10) Slide- “Artery, Vein & Nerve” (Cardiovascular- Vessels) (5)

Digital Microscopes and PC’s (2) Dissecting Microscopes and light sources (5)

Slide- Motor Nerve Smear/ Giant Multi-polar Neurons (10)

Digital Projector

Slide- Nerve Fibers, Osm Tet (10)

Slide- Peripheral Nerve (10)

Slide- “Nerve Tissue combination Mammal” (10)

Suggested Reading:

ZOOL 141L

13

ZOOL 141L: # 13 Nervous Tissue

Exercise 17: Histology of Nervous Tissue Activity 1: Parts of a neuron Examine the following microscope slides and sketch each. As always, indicate total magnification (TM). Also, label indicated structures and list function(s).

Slide TM Structures to identify 1) Motor Nerve Smear 400x Cell Body, Nucleus, Axon 2) Nerve Fibers (Osm Tet) 100x Schwann Cell, Nodes of Ranvier

Activity 2: Microscopic Structure of Selected Neurons Make a detailed sketch of each of the following CNS sections (note all are available on one slide labeled “Nerve Tissue Combination Mammal”) at scanning or low-power (40x-100x TM) and label indicated structures.

Section TM Structures to identify 1) Spinal Cord 40x Gray & white matter, cell body, central canal 2) Cerebellum 40x Gray & white matter, Purkinje cell 3) Cerebrum 40x Gray & white matter, cell bodies, Sulcus, Gyrys Question: 1) What are the structural and functional differences of white and gray matter (what

types or “parts” of neurons do they contain?) 2) How does the distribution of gray and white matter differ between the brain and

spinal cord? Activity 3: Microscopic Structure of a Nerve Make a detailed sketch of a peripheral nerve in transverse/ cross-section and label the indicated parts. Refer to figure 17.7a (p 264) for a schematic of nerve anatomy.

Section TM Structures to identify 1) Peripheral Nerve (cs) 40-100x Epineurium, Perineurium, Fascicle,

Myelinated Axon & Blood Vessels Instructors Note: This week’s lab focuses on the identification and classification of nervous tissue. A full write-up is not required.

Gross Anatomy of the Brain and Spinal Cord

Lab Manual Exercises: Exercise 19: Gross Anatomy of the Brain Exercise 21: Spinal Cord Anatomy and ANS Learning Outcomes:

1) Identify the Macro anatomy of the brain and spinal cord. 2) List major functions of brain and spinal cord regions. 3) Describe the functional differences between gray and white mater. 4) Identify the meninges.

Materials:

Dissecting Microscopes and light sources (5)

A&P Revealed CDs (Nervous System ) (10)

Dissecting Pans (10) Preserved Spinal Cord (cow or sheep) (10)

Dissecting Instruments (10 sets) Spinal Cord Models (2)

Sheep Brains (10) Nervous System Posters (all)

Skeleton with spinal nerves (1) Gloves and Goggles (20)

Laptops (10) Brain Bucket

Disposal Bags (not biohazard) Sharps Container

Suggested Reading:

ZOOL 141L

14

ZOOL 141L: # 14 Brain and Spinal Cord

Exercise 19: Gross Anatomy of the Brain

1) Use Brain Models, A&P Revealed CD’s, and dissected sheep brains to identify the structures on the accompanying list.

2) Make a detailed drawing of the brain in your lab notebook. Label all structures named on the accompanying list.

3) List the function(s) of boldface anatomy. 4) Complete the Review Sheet # 19 and include in your lab notebook.

Exercise 21: Spinal Cord, Spinal Nerves and the ANS

1) Use spinal cord model, A&P revealed CD and dissection of sheep/cow spinal cord to identify the structures on the accompanying list.

2) Make a detailed drawing of the spinal cord in your lab notebook. Label all structures named on the accompanying list.

3) List the functions of boldface anatomy. 4) Complete Review Sheet # 21 and include in your lab notebook.

Instructors Note: This week’s lab focuses on the identification and classification of nervous tissue. A full write-up is not required.

Lab Notebook Checklist: Lab 14: Brain and Spinal Cord � Detailed brain drawing, with labels � Functions of boldface brain anatomy � Review Sheet # 19 � Detailed spinal cord drawing with labels � Functions of boldface spinal cord anatomy � Review Sheet # 21

ZOOL 141L: # 14 Brain and Spinal Cord

Division Anatomy NervesCerebrum longitudinal fissure I. Olfactory

central sulcuslateral sulcusFrontal LobeTemporal LobeOccipital LobeInsula

Diencephalon Olfactory Bulbs II. OpticOlfactory TractsThalamusHypothalamusEpithalamus (Pineal Gland)Optic ChiasmaPituitary Gland

Brainstem Midbrain III. OculomotorIV. Trochlear

Pons V. TrigeminalVI. AbducensVII. FacialVIII. Vestibulocochlear

Medulla Oblongata IX. GlossopharyngealX. VagusXI. AccessoryXII. Hypoglossal

Decussation of PyramidsSuperior ColliculiInferior Colliculi

Cerebellum Arbor VitaeVermisFolia

Meninges Dura MaterArachnoid MaterFalx CerebriFalx Cerebelli

Gross Anatomy of Brain and Cranial NervesRequired Anatomy

Human Reflex Physiology and General Sensation

Lab Manual Exercises: Exercise 22: Human Reflex Physiology Exercise 23: General Sensation Learning Outcomes:

1) Define reflex and reflex arc. 2) Name and identify elements of common reflex arcs. 3) Conduct common spinal and cranial reflexes and discuss their clinical importance. 4) Define exteroceptor, interoceptor, and proprioceptor. 5) Determine the two-point threshold for different areas of the body . 6) Define and demonstrate receptor adaptation. 7) Define and demonstrate referred pain and discuss its physiological basis.

Materials:

Reflex Hammers (10) Black Sharpie (10)

Tongue Depressors (10) Sharpie (any color except black- 10)

Rulers (10) Cooler with Ice water

Pen Flashlights (10) Alcohol Pads (Box)

Reaction Times (6) w 4/ AA batteries each

Insulated Mall Probes (10)

“General Sensation” kit (1) Towel

2-point discrimination calipers (10) Reflex Arc Model (1)

Suggested Reading:

ZOOL 141L

15

ZOOL 141L: # 15 Reflexes & General Sensation

PART I. Reflexes and Reflex Arcs (Exercise 22)

Activity 1: Patellar knee-jerk reflex

1) Test the knee-jerk reflex for each member of the class using a reflex-hammer. Have the

subject sit on the lab bench with the lower leg unsupported (see figure 22.4, p 343).

2) Record positive (+) and negative (-) responses on the attached data sheet. If the patellar

reflex is abnormal (ABN), record this on the data sheet as well.

3) Diagram the patellar reflex in your lab notebook, indicating sensory neurons,

interneurons, and motor neurons.

4) Answer the following questions in your lab notebook:

a. What muscle(s) contract during a normal patellar reflex? Which are relaxed?

b. What nerve is carrying afferent and efferent nerve impulses?

c. Is the reflex ipsilateral or contralateral? Monosynaptic or polysynaptic?

d. What would an abnormal patellar reflex look like? (You may need to do an

internet search to answer this question).

e. What type(s) of conditions may be indicated for an abnormal patellar reflex?

Activity 3: Plantar Reflex

1) Test the plantar reflex for each member of the class using the handle of the reflex

hammer. Record positive (+) and negative (-) results in the attached data table.


a. What muscle(s) contract during a normal plantar reflex? Which are relaxed?

(note, we have not previously discussed some of these muscles in lab)

b. What nerve is carrying afferent and efferent nerve impulses?

c. Is the reflex ipsilateral or contralateral?

d. What would an abnormal plantar reflex look like? (You may need to do an

internet search to answer this question)

e. What type(s) of conditions may be indicated for an abnormal plantar reflex?


Activity 6: Pupillary Reflex

1) Follow the directions outlined in activity 6 of your lab manual (p 346). Record the

“normal” diameter of the pupil by holding a ruler close to the eye. (Do this for both R

and L pupils.) Record these baseline measurements in the attached data table.

2) Have the subject shield the right eye with his or her hand. Then shine a flashlight into the

left eye for five seconds. Measure the pupil diameter in both eyes immediately and

record in the data table.

3) Graph the average pupil diameter for both the left and right eye before (control) and after

the light was shined in the left eye.

4) Answer the following questions:

a. What is the function of the pupillary reflex (why is it “beneficial” for the pupils to

change diameter when exposed to a bright light?)

b. Which pupil showed the greatest change in diameter?

c. Did the right pupil (not exposed to light) change size after the experiment? If so,

explain why.

d. What part of the brain is responsible for initiating the pupillary reflex?

e. What could an abnormal pupillary reflex indicate?

Activity 7: Ciliospinal reflex

1) Test the ciliospinal reflex for each member of the class. Follow the directions on p. 346

of your lab manual. Record papillary dilation (+) or constriction (-) in the attached data

table. If the subject shows no visible change in pupil diameter, record “no response”

(NR) in the data table.

Activity 9: Reaction Time

1) Use the supplied reaction timers to test the reaction time of each student using auditory

(sound) stimulus. Test each student with five trails of three stimuli each given at random

intervals. Separate each trial by 10 seconds. Record the reaction times in the attached

data table and calculate the average for each of the four trials.

2) Repeat using visual (light) stimuli. Record the data and calculate class averages.


3) Graph the average reaction time of each of the four trials for both sound and visual

stimuli. Use the same scale for each graph.


a. Which trial had the shortest reaction time for sound stimulus? For light?

b. How did the reaction time of the fourth light trial compare to that of the first

sound trial? What does this say about the nature of learned reflexes?

c. Is there any evidence that the light trials enabled a greater reaction time for sound

trials?

d. How would you modify the experiment to test for differences in learning to react

to sound vs. light stimulus?

Other Activities (time permitting):

p. 345 ACTIVITY # 4 Initiate corneal reflex. Answer questions.

p. 345 ACTIVITY # 5 Initiate Gag reflex.

p. 346 ACTIVITY # 8 Initiate salivary reflex. Answer questions #1-3.

PART II. General Sensation (Exercise #23)

Activity 2: Determining two point discrimination In this exercise you will test different body areas (face, back of hand, palm of hand, fingertips, lips, back of neck, and ventral forearm) for 2-point discrimination threshold.

1) Which body area do you think will have the finest 2-point discrimination threshold? (Record your hypothesis)

2) Measure the two-point threshold for each body area using the plastic calipers supplied in lab. Record in the attached data table.

3) Graph the average two point threshold for each body area. 4) Was your hypothesis supported or refuted by the data? Include a brief conclusion

statement in your lab notebook. Activity #3 Testing tactile localization As demonstrated in the last activity, different areas of the body contain different numbers of touch receptors. In this activity, you will test the ability of the body to localize (determine exact placement) of touch stimuli.

1) Which area of the body will have the best degree of tactile localization? (Record your hypothesis).

2) Have the subject sitting down, eyes closed, with his or her palm outstretched. 3) Touch the subject’s hand with a felt-tip marker. 4) With their eyes still closed, have the subject try to hit the same spot with their own

marker (must be of a different color).


5) Use calipers to measure the distance between the two dots for each body area. Record this in the attached data table.

6) Graph the average error of localization for each area of the body. 7) Did the results support your hypothesis? Write a brief conclusion statement.

Activity # 7 Demonstrating referred pain

1) Immerse the subjects elbow in a cooler of ice water 2) Record the quality (severity) and locality of pain at 1 minute intervals (see table on p. 360

of your lab manual) 3) Question: how did the progression and quality of pain change during the experiment?

Other Activities (Time permitting)

p. 358 ACTIVITY #4 Plotting relative density of touch & temperature receptors;

359 ACTIVITY #5 Demonstration of adaptation of touch receptors;

p. 359 ACTIVITY #6 Demonstration of temperature receptors, p. 260;


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Special Senses: Vision, Hearing, Equilibrium, Olfaction & Taste

Lab Manual Exercises: Exercise 24: Vision (Activities 3, 5, 7 &10) Exercise 25: Hearing & Equilibrium (Activities 1, 3, 5, & 7) Exercise 26: Olfaction and Taste (Activities 1, 2, & 3) Learning Outcomes:

1) Define reflex and reflex arc. 2) Name and identify elements of common reflex arcs. 3) Conduct common spinal and cranial reflexes and discuss their clinical importance. 4) Define exteroceptor, interoceptor, and proprioceptor. 5) Determine the two-point threshold for different areas of the body . 6) Define and demonstrate receptor adaptation. 7) Define and demonstrate referred pain and discuss its physiological basis.

Materials:

Dissectible Eye Models (4) Compound Microscopes & Kits (10)

Dissecting Pans (10) Snellen Eye Charts (2)

Dissecting Tools (10) Large Test Tubes (10)

Digital Projector Sharpies (10)

Sheep or Cow Eyes (10) Ophthalmoscope/ Otoscope (1)

Gloves and Safety Goggles Astigmatism Chart (2)

Sensory Slides (Cochlea, Taste bud & Retina) (10)

Color-Blindness Tests (2)

Freakishly Giant Ear Model (1) Tuning Forks (2 sets)

Audiometer* (optional) Sterile Cotton Swabs (20)

Lemon Juice Bowl of Sugar

Carolina Biological Smell Kits (2) Vials of Odorants

Rulers Nose clips

Autoclave Bag Sugar

ZOOL 141L

16

ZOOL 141L: # 16 Special Senses

Suggested Reading: Exercises 24-26

PART I. Vision (Exercise 24)

Activity 3: Microscopic anatomy of the retina & sheep eye dissection (p367)

1) Obtain a longitudinal section of a human retina.

2) Draw or photograph at 100X. Label photoreceptors, bipolar cells, ganglion cells, pigmented retinal

epithelium, and choroid.

3) Obtain a whole sheep eye and dissection instruments. Dissect following the instructions on p 368.

Photograph or draw in your lab notebook. Label the following: ciliary body, lens, cornea, retina,

choroid, optic disc, sclera, optic nerve, vitreous humor, iris, and pupil.

Activity 5: Demonstrating the Blind Spot (p370)

1) Demonstrate the presence of your blind spot following the directions in your lab notebook. It is not

necessary to include the results in your lab notebook.

Activity 7: Testing Visual Acuity and Accommodation (p 372)

Question: Does visual acuity change with age?

Hypothesis:

1) Choose a minimum of 10 people from each class to participate in the experiment. The group should

contain representatives from all age groups (preferably spanning 20 years or more).

2) Have the subjects remove eyeglasses or contacts.

3) Evaluate the unaided visual acuity using a Snellen eye chart read at a distance of 20 ft. Record the data

in Table 1.

4) Determine the near-point accommodation distance for each of the subjects by holding a pen (preferably

with writing on the side) approximately 3 ft in front of the subject.

5) Move the pen slowly towards the subject’s nose, stopping when the writing becomes blurry or

indistinct. Measure this distance (in cm) and record in Table 1.

6) Make a scattergraph for both visual acuity and near-point accommodation data. Place “age” on the x-

axis and “Visual Acuity” or “Near Point Accomodation Distance” on the y-axis. Graph all individual

values rather than averages.

7) If possible, draw a “best fit” line though the data points.

8) Conclusion: Do the data support your hypotheses?


Activities 8 & 9: Testing for astigmatism and colorblindness (Optional- p 372)

Activity 10: Effects of Binocular Vision on Depth Perception (p373)

Q: What effect will covering one eye have on depth-perception?

H:

1) Choose a minimum of 10 people from each class.

2) Have each subject try to put a pen into the opening of a large test-tube (see p 373 for specifics) immediately

after it has been moved. Repeat 10 times. Record the number (%) of trials for which the subject got the

pen into the tube.

3) Repeat # 2 with one eye covered. Record data in Table 1.

4) Calculate the average accuracy for 1-eye and 2-eye trials and graph.

5) Conclusion: Did the data support your hypothesis?

Activity 12: Conducting and Opthalmoscopic Examination (Optional- p 374)

PART II. HEARING & EQUILIBRIUM

Activity 1: Identifying structures of the Ear (p 383)

1) Be able to identify all bold face anatomy using the 3-D ear model. It is not necessary to include anything

in your lab notebook for this activity.

2) Examine a histological slide of the cochlea. Identify major anatomical features and compare to figure 25.3.

Activity 4: Conducting Laboratory tests for Hearing

Q: What effect will age have on hearing acuity?

H:

1) Choose a minimum of 10 people from each class to participate in the experiment. The group should

contain representatives from all age groups (preferably spanning 20 years or more).

2) With the subject seated, rub your thumb and index finger together close to one ear.

3) Slowly move your fingers away (while still rubbing) until the person can no longer hear them. Measure

this distance (in cm) and record in Table 2.

4) Repeat for the opposite ear.

5) Make a scattergraph for auditory acuity data. Place “age” on the x-axis and “Auditory Acuity” on the y-

axis. Graph all individual values rather than averages.

6) If possible, draw a “best fit” line though the data points.

7) Conclusion: Do the data support your hypotheses?


Part III. OLFACTION & TASTE

Activity 2: Microscopic Examination of Olfactory Epithelium

1) Obtain a microscope slide of human tastebuds.

2) Examine at 400X and draw in your lab notebook.

Activity 3: Stimulating Taste buds

1) Sketch a tongue in your lab notebook.

2) Dry the surface of your tongue with a paper towel.

3) Dip a cotton swab in a small quantity of sugar. Sequentially apply the sugar to different parts of the tongue

(tip, middle, sides & back). Indicate area(s) where sensation was most intense by writing “S” at those

locations on your tongue map.

4) Repeat step #3 with lemon juice. Indicate area(s) where sensation was most intense by writing “O” at those

locations on your tongue map

5) Q: Based on the above experiment, do you think “sweet” and “sour” sensing taste buds are distributed

equally across the tongue?

Lab Notebook Checklist: Lab 16: Special Senses Lab Report Required � Retina drawing w/labels � Sheep eye drawing w/ label � Visual Acuity Hypothesis � Visual acuity scattergraph � Visual acuity conclusion � Depth perception hypothesis � Depth perception graph � Depth perception conclusion � Hearing acuity hypothesis � Hearing acuity graph � Hearing acuity conclusion � Taste bud drawing


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