Human Physiology BIO 5 Laboratory Manual - cabrillo.eduytan/Bio5/Bio5LabManF11NEW.pdf · Introduction to the Physiology Laboratory Physiology is commonly described as the study of

Human Physiology

BIO 5

Laboratory Manual

Cabrillo College Spring 2011

Table of Contents

Introduction to the Physiology Laboratory................................................................2 Exercise 1: Acids, Bases, and Buffers .......................................................................3 Exercise 2: Aerobic Capacity...................................................................................13 Exercise 3: Movement Across Cell Membranes......................................................25 Exercise 4: BioPac Tutorial - Physiological Instrumentation..................................33 Exercise 5: Electromyography.................................................................................51 Exercise 6: Contractility of Skeletal Muscle ...........................................................65 Exercise 7: Electrocardiography ..............................................................................77 Exercise 8: Respiratory Physiology.........................................................................93 Exercise 9: Blood .................................................................................................. 113 Appendix A: Graphing Basics .............................................................................. 123

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Introduction to the Physiology Laboratory

Physiology is commonly described as the study of biological function, while anatomy is described as the study of biological structure. In fact, each discipline must consider BOTH structure and function in order for either discipline to be meaningful. A very practical difference between anatomy and physiology, however, is whether the subject is dead or alive. Anatomical structures can be studied using preserved specimens. Physiology, on the other hand, must be studied in a live subject (or in a close approximation).

Studying a live subject poses certain problems for both the researcher and the subject in a course like Human Physiology. The risk of transmission of infectious agents is too high to monitor bodily fluids, typically saliva, urine, or blood, in a classroom setting. We have also made a conscious decision to minimize the use of live animals, although we have not altogether eliminated them. To circumvent these problems, we are using a variety of alternatives including the non-invasive monitoring of bodily functions (pulse, ECG, etc.), computer simulations, in vitro (in glass) demonstrations, and the analysis of case studies.

Scattered throughout and at the end each exercise are study questions that are meant to help guide you to the relevant points in the lab. Weekly quizzes will be based on these questions and on the assigned background reading from the textbook, Human Physiology: An Integrated Approach, 5th ed., by Dee Silverthorn. The pages for the background reading can be found under the title of each exercise.

These lab exercises are meant to teach you something about experimental design and methods, as well as help you understand the physiological concepts we discuss in lecture. In choosing and writing up these exercises, we attempted to always ask “What is the point of doing this?” and “What do we want the student to learn?”. We want these exercises to be fun, but also relevant and useful. We hope that you will find your experience with us this semester all of that.

A NOTE CONCERNING "AFFECT" AND "EFFECT": Students very commonly get the terms "affect" and "effect" confused. "Affect" is a verb that means to have an influence. "Effect" is a noun meaning the result or outcome. A change or stimulus will affect the body's physiological state (example: Arteriosclerosis affects blood pressure). The resulting effect of that change or stimulus is the response (example: High blood pressure is one effect of arteriosclerosis). In other words, "af-" is the imput and "ef-" is the output.

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Exercise 1: Acids, Bases, and Buffers Readings: Silverthorn 4th ed, pg. 35 – 38; Silverthorn 5th ed, pg. 36 – 39.

The acidic or basic properties of solutions are due to their relative concentrations of hydrogen ion (H+) and hydroxyl (OH-) ion. Acidic solutions have high concentrations of hydrogen ion and low concentrations of hydroxyl ion. Basic (also called alkaline) solutions have low hydrogen ion concentrations and high hydroxyl ion concentrations. Acids will bring about an increase in the hydrogen ion concentration of a solution, while bases will bring about an increase in the hydroxyl ion concentration.

Why are acids and bases important to biology? Because life is a series of chemical interactions that occur in water, and the acidity or alkalinity of a water solution greatly influences the chemistry that is possible. Acids and bases most strongly affect the enzymes that serve as catalysts for life's chemistry. If the proper enzymes are not functioning because of acid or alkaline conditions, life processes will cease.

Enzymes are composed of proteins whose biological activity is dependent upon an exact, three-dimensional shape. Substances that alter the shape or charge on proteins are going to have significant effects on life processes. For example, the reason high temperatures kill living things is because the high temperatures change protein structures.

Like high temperatures, the relative concentration of hydrogen ion and hydroxyl ion exert effects on the shape of proteins, primarily by disrupting hydrogen bonding. This results in a loss of enzyme activity. Most multicellular organisms have the ability to regulate the acid-base conditions in their bodies and can live in environments that are quite acidic or basic. Individual cells on the other hand, can generally tolerate only a very narrow acid-base range.

The human body must maintain a very narrow range of pH between 7.38 and 7.42 (note that this pH is not precisely neutral). The human body uses three main mechanisms for maintaining this narrow range of acid/base balance. The first and quickest acting mechanism is the chemical buffering systems, the most important of which is the carbonic acid/bicarbonate ion system. When chemical buffers are overwhelmed, usually by increasing concentrations of H+, the second mechanism, the respiratory system, can regulate acid/base balance by eliminating CO2 (see your textbook for an explanation of this mechanism). These two mechanisms can only neutralize pH by converting H+ or OH- into H2O. The only way to actually remove excess H+ is by the third mechanism, the renal system. This mechanism actively secretes excess H+ into the urine during acidosis, or HCO3

– during alkalosis.

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Today’s Objectives

1. Review the basic principles of acid/base chemistry.

2. Model a carbonic acid/bicarbonate buffering system using molecular models.

3. Test the pH of a variety of common liquids.

4. Observe pH fluctuations in an unbuffered solution.

5. Create a chemical buffering system and examine its effect on pH fluctuations.

The Chemistry of Acids and Bases

Logically, you might only expect to find only water molecules (H2O) in a beaker of pure water. However, if you perform an exact analysis, you find very small but equal concentrations of hydrogen ion, (H+) and hydroxyl ion (OH-) in addition to water molecules. The reason is a small fraction of the water molecules have dissociated to give equal concentrations of hydrogen ion and hydroxyl ion:

H2O → H+ + OH-

The solution is neither acid nor base, and is said to be neutral.

Further observations on pure water would demonstrate that the number of water molecules dissociating is always the same, namely 1X10-7 M (0.0000001 moles per liter). This doesn't sound like much but actually accounts for more than 1X1016 (more than a thousand trillion) dissociated molecules in a liter of water. Since each dissociated water molecule creates one H+ and one OH- both of these ions are at concentrations of 1X10-7M.

When an acid such as hydrochloric (HCl) is added to the water solution, the acid dissociates:

HCl → H+ + Cl-

The acid adds to the [H+] ([H+] is read as "Hydrogen ion concentration") and it increases [H+] above 1X10-7M. An interesting thing happens here. As the [H+] increases, there is a proportional decrease in [OH-]. In fact, you would find that if the [H+] is multiplied by the [OH-], the result will always be 1X10-14. This number is known as the water dissociation constant.

Acids can be defined as H+ (or proton) donors, while bases can be defined as H+ acceptors. In other words, acids increase the [H+], while bases decrease it. Examine the following list of common acids and bases. Notice how the acids all dissociate to produce H+, thus increasing the [H+]. Some bases, like NaOH, can dissociate to produce OH- as the H+ acceptor. Compounds like ammonia (NH3) and bicarbonate ion (HCO3

–) are also H+ acceptors and are considered bases.

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Some Common Acids

Hydrochloric Acid HCl → H+ + Cl-

Nitric Acid HNO3 → H+ + NO3–

Acetic Acid CH3COOH → H+ + CH3COO–

Some Common Bases

Sodium Hydroxide NaOH → Na+ + OH–

Potassium Hydroxide KOH → K+ + OH–

Ammonia H+ + NH3 → NH4

It is awkward and time consuming to always write out the [H+] in Molar notation. Instead, there is a better way of denoting the [H+]. The term for this notation is pH, and is defined as the negative logarithm of the hydrogen ion concentration, or abbreviated:

pH = –log[H+] Even though no units are used with pH notation, remember that pH notation represents a concentration in moles per liter (M). Let's say you want to know the pH of a neutral solution. You know that at neutrality:

[H+] = [OH–] and that [H+] X [OH–] = 1X10–14 M

thus: [H+] = 1X10–7M

and: pH = -log [H+] = -log(1X10–7M) = -(-7) = 7

Notice that the log is simply the exponent of 10 of the number in question. Therefore, the pH of a solution at neutrality is 7.

Now, suppose we add some HCl to increase the [H+] to 1X10-4M. What is the pH at this [H+]?

pH = -log [H+] = -log(1X10–4M) = -(-4) = 4

What happens if a base, such as NaOH is added to a neutral solution? The [H+] decreases because [H+] X [OH–] must equal the dissociation constant. Let's say the [H+] is decreased to 1X10–9M. What is the pH?

pH = -log [H+] = -log(1X10–9M) = -(-9) = 9

You should now see a relationship between [H+] and pH developing. At pH 7, a solution is neutral; as the pH decreases, the [H+] is increasing and the solution is becoming more acidic; as the pH increases above 7, the [H+] is decreasing and the solution is becoming more basic.

Modeling the Carbonic Acid/Bicarbonate Buffer

In order to maintain a narrow pH range, the blood and other body fluids must be buffered. Buffers are composed of conjugate acid-base pairs. Conjugate acid-base pairs are compounds

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that differ by the presence of one proton, or H+. There are several buffering systems in body fluids, but the most common is the carbonic acid/bicarbonate ion system:

H2O + CO2 → H2CO3 → H+ + HCO3–

Water and carbon dioxide combine to form a weak acid, carbonic acid (H2CO3) some of which dissociates to form H+ and bicarbonate ions (HCO3-). Note that the reactions are reversible and thus may go in either direction, depending on relative concentrations.

Materials Bond models:

• Single bonds: fat gray pegs • Multiple bonds: skinny gray flexible pegs. These are used to form two bonds between

the same atoms. Ball models:

• Carbon (C): black balls with 4 holes (each hole represents one potential bond) • Hydrogen (H): white balls with 1 hole • Oxygen (O): red balls with 2 holes • Nitrogen (N): blue balls with 4 holes (we won't be using these)

Procedure:

In the following diagrams, single lines connecting atoms indicate single bonds. Two lines indicate a double bond. Double bonds are two bonds that form between the same two atoms. The two atoms are sharing a total of 4 electrons.

Use the long flexible connectors to form double bonds. Each connector represents one pair of electrons, so you need two connectors to make a double bond.

1. Start out by making a water molecule and a carbon dioxide molecule:

Water: H2O O

H H

Carbon dioxide: CO2 O = C = O

2. Combine these two molecules to form carbonic acid: H2CO3

O || C

/ \ HO OH

3. Remove one of the hydrogen atoms to make bicarbonate ion: HCO3 –

4. Combine your carbonic acid or bicarbonate ion and hydrogen ion with everyone else's to make a buffering system. The instructor will demonstrate how this conjugate acid and base pair makes a buffer and prevents fluctuations in H– concentration.

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The pH of Some Common Solutions

You will now use a hand held Checkmite pH meter to measure the pH of some common solutions. Make sure they are all at room temperature before making any measurements. You will need to calibrate the meter before you begin your measurements.

Calibrating the pH meter 1. The pH meters are stored with buffer solution in the cap to prevent the sensor from drying

out. Carefully remove the cap on the sensor tip of meter. Put the cap some place safe so it doesn’t get lost. Rinse sensor tip with tap water and blot to remove excess water. Press ON/OFF button to turn the meter on.

2. Immerse about 1/2" to 1" of the sensor tip in pH 7.00 buffer.

3. Press CAL button to enter Calibrate (CA) mode. 'CA' flashes on the display. Then, a pH value close to 7 will flash repeatedly.

4. After at least 30 seconds (about 30 flashes) press the HOLD/CON button to confirm calibration. The display will show 'CO' and then switch to the buffer value reading.

5. Rinse the electrode in tap water before pH testing.

Measuring the pH of a Sample 1. Select about six different samples to test.

You do not have to test all available samples. Space is available for samples not listed in Table 1.

2. Pour about 1" of the sample into the small plastic cup provided. Immerse the sensor tip in your sample solution. Let the reading stabilize.

3. Note the pH or press HOLD/CON button to freeze the reading. Press HOLD/CON button again to release the reading.

4. Rinse and blot the sensor tip with tap water. Always remember to rinse the tip between each sample.

5. If you do not press a button on the pH meter for 8.5 minutes, the meter will automatically shut off.

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Table 1. The pH of some common solutions

Solution pH Deionized water Apple juice (canned) Vinegar Coffee (instant, decaf) Tea (black from bag) Lemon juice (bottled) Coke (Classic) Milk (nonfat) Mylanta

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Unbuffered and Buffered Solution

In this exercise, you will compare the effect of adding a strong acid and base to first an unbuffered solution, then a buffered solution.

Measuring pH Fluctuations in an Unbuffered Solution (Negative Control) 1. Start with clean 1000 ml beaker filled with about 300 ml DI H2O.

2. Place the beaker on the magnetic stirrer and drop a clean stirring bar into it.

3. Turn on the stirrer and adjust the speed so that a small whirlpool is just barely established.

4. Add 2 ml of Universal Indicator, a pH indicator that will change color as the pH changes.

5. Record the pH and the color of the water in Table 2.

6. Add 1 drop of the 1.0N HCl to the beaker. When the pH has stabilized, record the pH of this solution.

7. In the same way, add two more drops of the 1.0N HCl to the beaker, recording the pH after each drop is added.

8. This time, add one drop of 1.0N NaOH. Be sure to continue to record the pH after each drop of acid or base is added and the pH has stabilized.

9. In the same way, continue to drops of NaOH and record the pH until a total of 6 drops has been added, or until the pH of the solution has increased to around pH 9 to 10.

Table 2. pH Fluctuations in an unbuffered water

Event pH Color

Deionized water at start

1st drop 1.0N HCl added

2nd drop 1.0N HCl added

3rd drop 1.0N HCl added

1st drop 1.0N NaOH added

2nd drop 1.0N NaOH added

3rd drop 1.0N NaOH added

4th drop 1.0N NaOH added





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Developing a Buffer System Earlier, you made a carbonic acid/bicarbonate ion buffer using molecular models. Now you will make one using the actual chemicals. We will use Alka Seltzer as a source of CO2 and baking soda as a source of HCO3

–.

H2O + CO2 → H2CO3 → H+ + HCO3–

Making the buffer 1. Start with a fresh 1000 ml beaker filled with 300 ml DI H2O. Be sure to rinse the beaker

well between this experiment and the last. Place the beaker on the magnetic stirrer and drop a clean stirring bar into it.

2. Turn on the stirrer and adjust the speed so that a small whirlpool is just barely established. Add 2 ml of Universal Indicator.

3. Record the pH and the color of the water in Table 3.

4. Place 50 ml of DI H2O in the stoppered flask. Drop 2 Alka Seltzer tablets into the stoppered flask and snugly replace the stopper.

5. Place the end of the glass tubing from the flask stopper into the swirling water. You should see CO2 bubbles swirling out of its tip.

6. When the Alka Seltzer tablets are just about dissolved, measure the pH of the solution in the beaker.

7. Add two more Alka Seltzer tablets to the flask and quickly replace the stopper.

8. Keep bubbling CO2 into the beaker until the last two Alka Seltzer tablets are dissolved. Measure the pH of the solution in the beaker.

9. Add about 1 to 2 teaspoon of NaHCO3 (baking soda) to the beaker. Allow all of the NaHCO3 to dissolve and record the pH.

Testing the buffer 1. Add 1 drop of the 1.0N HCl to the beaker. Record the pH in Table 3.

2. After about 30 seconds, add another drop of HCl. Record the pH.

3. After another 30 seconds, add a third drop of HCl. Record the pH.

4. You should observe only momentary fluctuations in the pH of the test solution.

5. Allow the pH stabilize, but now add one drop of 1.0N NaOH and record the pH.

6. In the same way, add a total of 6 drops of the NaOH solution, recording the pH after each drop.

7. When you are finished with the pH meter, turn it off and recap it. There should be a damp piece of sponge in the cap to keep the sensor moist.

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Table 3. pH Fluctuations in a buffered solution

Event pH Color

Deionized water at start

After first 2 Alka Seltzer tabs

After second 2 Alka Seltzer tabs

After adding baking soda

1st drop 1.0N HCl added

2nd drop 1.0N HCl added

3rd drop 1.0N HCl added

1st drop 1.0N NaOH added

2nd drop 1.0N NaOH added

3rd drop 1.0N NaOH added






About the Buffer

The carbonic acid/bicarbonate buffering system made today is the same buffer found in blood. The bubbling of CO2 into water mimics the CO2 produced by cellular respiration that diffuses out of cells and into the plasma. CO2 combines with H2O to form first carbonic acid, then bicarbonate ion:

H2O + CO2 → H2CO3 → H+ + HCO3–

It is the dissociated H+ that caused the initial drop in pH. To make our buffer, we added an excess of HCO3

– by adding the sodium bicarbonate, which dissociated to form Na+ and HCO3–:

NaHCO3 → Na+ + HCO3–

The Na+ ions have no effect on the pH and can be ignored. The HCO3– behaved as a weak base

and binds free H+, raising the pH and forming carbonic acid. This reaction is written below from right to left to emphasize that adding baking soda increased the amount of product, bicarbonate ion, to our initial reaction and shifted the equilibrium of the reaction to the left.

H2CO3 <–– H+ + HCO3−

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It is the combination of both the carbonic acid and the excess bicarbonate ion that forms the buffer. If acid is added, the bicarbonate ions will bind up the added H+ ions and prevent them from decreasing the pH. If base is added, the carbonic acid will release H+ to combine with the added OH– (or other proton acceptor) to form water and prevent them from increasing the pH. Any chemical buffering system requires both a conjugate acid and a conjugate base to donate or accept compensatory H+.

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Questions

1. Was the pH of the different beverages predominantly acidic or basic? Do you think drinking acid beverages such as Coke is damaging to the esophagus, stomach and intestine? Why or why not?

2. When you made the molecular models, how many steps did it take to reshuffle the CO2 model and the H20 model into carbonic acid? How many steps did it take to convert carbonic acid into bicarbonate? How many steps to convert bicarbonate into carbonic acid?

3. What is the formula for pH? What is the meaning of this number?

4. What is the pH of a solution whose H+ concentration is 1 X 10-3M? Be able to calculate the pH of any solution of known H+ concentration.

5. What is the H+ concentration of a solution whose pH is 4? Be able to calculate the concentration of any solution of a known pH.

6. What happens to the pH of a solution when H+ are added? When OH- are added?

7. Suppose you start with a solution at pH 7, and add acid until the pH of the solution is 4. How many times more concentrated in H+ is the new solution compared to the starting one? Be able to compare the concentrations of any two solutions of known pH.

8. Explain the purpose of each component of the buffer system you developed in this experiment. Explain how this buffer system resists changes in pH, using chemical equations to illustrate your explanation. What happens if you add an acid? What happens if you add a base?

9. What are the ingredients of the Alka Seltzer tablets? Explain how Alka Seltzer can relieve acid indigestion. How do you explain our use of the Alka Selter in our experiment versus how it is used as an antacid?

10. What is the normal homeostatic range of pH found in the human body? What effect do pH levels outside this range have at the cellular level?

11. What are the three main mechanisms for regulating plasma pH balance in the body?

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Exercise 2: Aerobic Capacity Reading: Silverthorn 4th ed, pg. 3-6, 191-200, 810-811; Silverthorn 5th ed, pg 4 – 5, 196 – 204, 816 – 817.

The purpose of this first exercise is to illustrate the relationship between the consumption of oxygen, energy production, and physical work.

Metabolism comes from the Greek word for “change” and refers to all the chemical and energy transformations that occur in a cell. Food is transformed into ATP in the mitochondria through aerobic respiration and the electron transport system. The overall chemical reaction for converting glucose into ATP is:

C6H12O6 + 6O2 + ADP 6CO2 + 6H2O + ATP

ATP is then transformed into mechanical energy to move our muscles and maintain our bodily functions. Oxygen is required for making ATP, so oxygen consumption is a reflection of ATP production (and useage). Energy expenditure can be indirectly measured by measuring the rate of oxygen consumption, or VO2. The units of VO2 are ml O2 consumed/kg body weight/min (the units L/min are sometimes used for non-weight bearing exercise). VO2 is the rate that oxygen is actually used by the body, not the amount of oxygen inhaled. Oxygen consumption changes as the body's activity level changes and more ATP is required. VO2 measured at rest will be lower than VO2 measured during physical activity.

VO2 max reflects a person’s overall fitness level, which is one's maximum potential for using oxygen to make ATP. This is also known as aerobic capacity. VO2 max is defined by the limits of ones oxygen transport system, so this value does not vary from moment to moment in the same way that VO2 will change. Individuals who are more physically fit will be able to use oxygen more efficiently (have a higher VO2 max), and can therefore produce more ATP and perform more work with less effort (huffing and puffing). Factors that can affect VO2 max include general respiratory and cardiovascular health, the amount of mitochondria in the muscle cells, and vascularization of the muscle tissue. A person can increase their VO2 max over time with physical training.

We will use a method called the McArdle-Katch Bench Stepping Test to estimate VO2 max. It is a convenient, low budget method of fitness assessment. We can measure heart rate while performing a sub-maximal amount of work, then extrapolate what our oxygen consumption would be if we were working maximally at VO2 max. This extrapolation is based on data compiled from the VO2 max measurements of many thousands of people of many different fitness levels (see Table 5). We will NOT be measuring VO2. Rather, we will be measuring heart rate and will use this measurement to extrapolate what VO2 max would be if we were working at a maximal level.

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Figure 1. Relationship between heart rate and oxygen consumption

This test is based on the assumption that a defined amount of work will require a set rate of oxygen consumption, or VO2, regardless of the fitness of an individual. The amount of work performed in this test is defined by the height of the step stool and the pace and number of steps performed. Everybody will perform the same amount of work and presumably consume the same amount of O2. Differences in fitness will be reflected by the amount of effort the heart must make in order to deliver that defined amount of O2.

A person that is more fit will be able to consume the same amount of oxygen with fewer heartbeats than a less fit person. In other words, if two people of the same age perform the same amount of work, the person who is more fit will have a lower heart rate than the person who is less fit. See Figure 1.

In addition to estimating VO2 max, recovery time after exercise can also be used as an indicator of cardiovascular fitness. The oxygen debt incurred during exercise can be paid back more quickly if the ATP-producing apparatus in the cell is working more efficiently. Therefore, faster recovery time indicates better aerobic fitness. We will be using the 1-minute post-exercise heart rate as a second indicator of aerobic capacity.

As activity level decreases and oxygen debt is repaid, heart rate decreases with the decreased need for oxygen. The heart rate's response to changes in the body’s demands is called homeostasis. Homeostasis is the dynamic fluctuation of a physiological process in response to changes in the body’s internal environment. Homeostasis can also be described as the maintenance of a body function within set boundaries. This chain of cause and effect is part of a monitoring system known as negative feedback: as activity level decreases, less ATP is needed. This decreases oxygen consumption which then causes O2 levels to increase and CO2 levels to decrease in the blood. The change in blood gas is monitored by chemoreceptors and sent to the brain, which then sends a signal to the heart to slow it down. Thus, the heart rate will match the body's need for oxygen delivery.

The maintenance of homeostasis underlies most physiological processes. Homeostasis is one of the physiological themes that we will return to again and again throughout the semester.

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1. Use the McArdle-Katch Bench Stepping Test to predict VO2 max for each member of your group.

2. Correlate VO2 max to each individual's aerobic fitness and the process of cellular respiration.

3. Measure post-exercise heart rate and correlate length of recovery time to aerobic fitness.

4. Apply post-exercise recovery of heart rate to the concept of homeostasis.

5. Determine an appropriate exercise regime for each of the various fitness levels.

Predicting V02 max

The McArdle-Katch bench stepping test is used to predict VO2 max by having the subject perform a defined amount of work and estimating their energy expenditure based on their heart rate immediately after. The subject steps at a rate of 88steps per minute (females) or 96 steps per minute (males) for 3 minutes. The bench height is 16.25 inches. Validity of the test is highly dependent on the accurate measurement of pulse rate (W.D. McArdle et al. (1972) Medicine and Science in Sports, Vol. 4, p 182-186).

Before You Begin 1. Do a quick self-assessment of your fitness to predict what your fitness level might be. If you

have any health concerns that would prohibit you from participating, use data from a lab partner. Table 6 is a prescription for an exercise regimen recommended by the American College of Sports Medicine. Read the description of exercise intensity in Column 2 of Table 6. This is how intensely you should feel comfortable exercising, depending on your fitness level (Column 1). Make a guess as to where you might fit.

2. Practice taking either your radial pulse in your wrist or your carotid pulse in your neck. If you use your radial pulse, place your index and middle fingers over the radial artery on the anterior side of the wrist just lateral to the tendon of your flexor carpi radialis muscle (remember any of your anatomy?) and press lightly.

If you use your carotid pulse, place two fingers just below the angle of your mandible, or lower jaw, just over the carotid artery.

Whichever method you choose, do not press too hard or it will occlude (close) the artery and make it difficult to feel the pulse.

3. Convert the number of beats you counted in 10 seconds to Beats Per Minute (BPM), which are the standard units used to measure heart rate, by multiplying this number by 6 as per the following formula:

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heart beats X 60 seconds = beats per minute

10 seconds 1 minute

4. Record your Heart Rate (HR) results in Table 1.

5. Take your resting pulse two more times and convert to Heart Rate. How close were your three measurements? Calculate your average resting HR by adding the three numbers together and divide by 3.

McArdle-Katch Bench Stepping Test 1. One lab partner will be your recorder, a second will be your timekeeper. The recorder will

write down your data. The timekeeper will signal you at each of the following time points:

• 10 seconds before the 3-minute mark as a warning

• 5 seconds post-exercise when you will begin counting your pulse beats for 10 seconds

• Every 30 seconds for the next 7 minutes when you will continue to count pulse beats for 10 seconds

2. Be sure you are wearing skid-resistance shoes, like athletic shoes or sneakers. Warm up a bit by gently stretching your leg muscles.

3. Set the metronome to 88 beats per minute for females or 96 beats per minute for males. You can turn off the sound of the metronome and use the flashing light to keep time if you prefer.

4. With the front of the steps facing the wall, brace the stepstool against the wall to prevent it from slipping. You will be stepping up to the top step of the stool, using a four-step cadence (right foot up, left foot up, right foot down, left foot down). If you are female, you will take 88 steps each minute. If you are male, you will take 96 steps in a minute. At the signal from the timekeeper, begin stepping in time with the metronome for a period of three minutes.

5. At the end of three minutes of stepping, remain standing and immediately find your pulse, using the same method you used to take your resting pulse rate. Begin counting pulse beats 5 seconds after you stop exercising for 10 seconds. Have your lab partner record this number in Table 1 as your zero time point. Leave your fingers on your pulse. Continue to report and record your pulse beats every 30 seconds by counting for 10 seconds, then waiting for 20 seconds , then counting for 10 seconds again. You will be counting your pulse beats every 30 seconds for 7 more minutes.

6. Convert beats counted in 10 seconds to Beats Per Minute (BPM).

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Table 1. Heart Rate Data (in BPM) Beats/10 sec Heart Rate (BPM) Resting Heart Rate #1

Resting Heart Rate #2

Resting Heart Rate #3

Average Resting HR

0:00 (5 sec after exercise)

0:30 (minutes:seconds)

1:00 = 1 minute Recovery HR

1:30

2:00

2:30

3:00

3:30

4:00

4:30

5:00

5:30

6:00

6:30

7:00

*NOTE: Only the zero time point (0:00) will be used for the McArdle-Katch Test. The rest of the time points will be used to plot a graph illustrating homeostasis of recovery heart rate. The 1 minute time point (1:00) will be used to calculate fitness from Table 6.

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Data Analysis

Determine Your Aerobic Capacity & Develop an Exercise Regimen

1. Determine your estimated Aerobic Capacity, or VO2 max (ml/kg/min). Only the zero time point (0:00) will be used to estimate as part of the McArdle-Katch Test. Use the following equations, where HR = heart rate at the 0:00 time point converted to BPM.

Females:

VO2 max = 65.81 – (0.1847 X HR)

Males:

VO2 max = 111.33 – (0.42 X HR)

Enter aerobic capacity in line 1 of Table 4.

2. Once you determine your aerobic capacity, you can use this information to design a personalized exercise regimen that will help you develop and maintain your aerobic fitness:

a. Use the standards provided in Table 2 for females and Table 3 for males to determine your Fitness Level based on your estimated VO2 max. Enter your Fitness Level in line 2 of Table 4. The standard error of prediction using these equations is +/- 16% of actual VO2 max.

b. Based on your fitness level, use Table 5 to determine the appropriate Exercise Intensity for your personal exercise regimen. Exercise intensity means working out at a pace that causes your heart rate to increase a given percentage of your heart's maximal ability. For example, if you are not very fit, you should begin your exercise regimen working your heart at only 60% of its maximum ability. If you are very fit, you can work your heart at 90% of its maximum ability. Enter your recommended exercise intensity in line 3 of Table 4.

c. Your heart's maximum ability, or your maximum heart rate, is referred to as HRmax. Your HRmax is estimated as 220 – your age. If you are 32 years old, then your HR max would be 188 BPM. Enter your HRmax in line 4 of Table 4.

d. Your personal fitness regimen will be determined by your Target Heart Rate. This is how fast your heart should be beating while you are exercising in order to safely strengthen your heart and improve your aerobic capacity. Calculate your target heart rate by multiplying your HRmax times your recommended exercise intensity. Enter this number in line 5 of Table 4.

e. Columns 3 and 4 of Table 5 suggests how long and how often you should exercise at your target heart rate in order to improve your aerobic fitness. Enter these recommendations in lines 6 and 7 of Table 4.

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Now you have developed a personalized exercise plan without the cost of a personal trainer! The American College of Sports Medicine recommends using these guidelines for about six weeks, then retesting your aerobic capacity for improvement. If your aerobic capacity has improved to the next level, then you can increase your exercise intensity accordingly. Good luck and happy exercising!

Table 2. Maximal Oxygen Consumption Rates Standards (ml/kg/min)

FEMALES

Age (years) Low Fair Average Good High

10–19 <30 30–37 38–46 47–56 >56

20–29 <26 26–32 33–42 42–52 >52

30–39 <24 24–29 30–38 39–48 >48

40–49 <21 21–25 26–35 36–44 >44

50–59 <19 19–23 24–33 34–41 >10

60–69 <18 18–21 22–30 31–38 >38

70–79 <16 16–19 20–27 28–35 >35

Table 3. Maximal Oxygen Consumption Rate Standards (ml/kg/min)

MALES

Age (years) Low Fair Average Good High

10–19 <38 38 – 46 47 – 56 57 – 66 >66

20–29 <33 33 – 42 43 – 52 53 – 62 >62

30–39 <30 30 – 38 39 – 48 49 – 58 >58

40–49 <26 26 – 35 36 – 44 45 – 54 >54

50–59 <24 24 – 33 34 – 41 42 – 50 >50

60–69 <22 22 – 30 31 – 38 39 – 46 >46

70–79 <20 20 – 27 28 – 35 36 – 42 >42

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Table 4. Calculations

1 Aerobic Capacity: VO2 max in ml/kg/min (from formula)

2 Estimated Fitness Level (from Table 2 or 3)

3 Recommended %HRmax (from Table 5)

4 Your HRmax (HRmax = 220 – age)

5 Target Heart Rate for your fitness level (%HRmax X HRmax)

6 Duration of Exercise per session

7 Frequency of Exercise per week

8 Fitness level based on the 1 minute recovery HR

Table 5. Exercise Prescription Guidelines Based on Fitness Level for Healthy Young Adults

Fitness Classification Based on VO2 max

Exercise Intensity Exercise Duration

Exercise Frequency

Low Females: < 29 ml/kg/min

Males: < 34 ml/kg/min

60-70% HRmax

Perceived exertion: fairly light to somewhat hard

Unaware of ventilation rate; Breathing and depth is comfortable; Capable of passing the “talk test”

20-30 min/session

3 days/wk

Average Females: 30-44 ml/kg/min

Males: 35-49 ml/kg/min

70-80% HRmax

Perceived exertion: somewhat hard to hard

Aware of ventilation rate (i.e. increased breathing rate and depth)

30-45 min/session

4 days/wk

High

Females: > 45 ml/kg/min

Males: >50 ml/kg/min

80-90% HRmax

Perceived exertion: hard to very hard

Hyperventilatory response; respiratory distress (i.e. rapid breathing rate with deep or large breaths); Incapable of passing the “talk test”

45-60 min/session

5 days/wk

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Questions

1. What is VO2 and how does it differ from VO2 max? How does VO2 max relate to cardiovascular fitness?

2. What physiological factors contribute to an increased VO2 max and consequently a greater efficiency in energy production (ATP production).

3. What is the relationship between physical activity, O2 consumption, and heart rate?

4. Why can heart rate be used to estimate VO2 max?

5. The McArdle-Katch Bench Stepping Test is only about 85% accurate. What steps could contribute to potential errors? What might be some of the advantages and disadvantages to using this test to assess aerobic fitness?

6. What is the target heart rate and why can it be used to determine how hard an individual should exercise, depending on their fitness level? What was your target heart rate and how did your assessed fitness level compare to your prediction?

7. Why should fitness level be considered when deciding on a fitness regimen?

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Homeostasis of your Recovery Heart Rate 1. To see how your heart recovers from exertion, make a graph of your recovery heart rate by

using the 7 minutes of data from Table 1. This graph will be turned in. Put "Recovery Time" on the X-axis and "Heart Rate" on the Y-axis. Label your graph with a title, and each axis with the proper units.

Plot your Resting HR across the bottom of your graph as baseline. On the same graph, plot each time point for your post-exercise heart rate. Do not use your resting HR as a zero time point or included your resting HR in the plot of your post-exercise heart rate.

Label on your graph the 1-minute post-exercise heart rate.

2. Use Table 6 to determine your fitness rating based on your 1-minute recovery heart rate. Enter this number in line 8 of Table 4. How does this rating compare to the rating you determined using the McArdle-Katch method?

NOTE: For a review of how to construct a graph, consult Appendix A: Graphing Basics, at the end of this manual. Remember, you can graph your data by hand or with a computer spreadsheet program. If you choose to graph by hand, you must use graph paper. Do not use binder paper or unlined paper.

Table 6. Ratings for 1-minute Recovery Rates (Heart rate one minute after exercise)

Fitness Rating Gender Heart Rate (BPM)

High Male <71

Female <97

Good Male 71 – 102

Female 97 – 127

Fair Male 103 – 117

Female 128 – 142

Low Male 118 – >147

Female 143 – >171

For more information on VO2 max, go to http://home.hia.no/~stephens/vo2max.htm

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http://home.hia.no/~stephens/vo2max.htm

Questions:

8. Why should time be plotted on the X-axis and heart rate on the Y-axis?

9. Draw out the feedback loop that governs changes in heart rate at rest and during physical activity. Describe with words how this diagram explains why your heart rate decreases with time after you stop exercising.

10. What makes this an example of negative feedback rather than positive feedback?

11. Explain why the 1-minute recovery rate is an indicator of cardiovascular fitness. Include the concept of homeostasis in your discussion.

12. How does the fitness level you estimated using the McArdle-Katch method compare to the fitness rating estimated using the 1-minute post-exercise heart rate? What, if anything, does this tell you about using different methods to obtain similar information?

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Exercise 3: Movement Across Cell Membranes Reading: Silverthorn 4th ed, pg. 132 - 136, 153 – 159; Silverthorn 5th ed, pg. 136 – 140.

A selectively permeable barrier is one of the defining features of a living cell. The cell membrane and the associated transport proteins found in the membrane are responsible for regulating the movement of hundreds, if not thousands, of different types of molecules into and out of the cell. All molecular motion is influenced by diffusion, which is the tendency for particles to spread from higher concentrations to lower concentrations until they are evenly distributed, or reach equilibrium. This movement towards equilibrium is the driving force behind a majority of physiological processes, from neuronal impulses to renal function.

Today we will investigate the movement of several different types of molecules across a cell membrane, including water, and we will examine the physical properties of these different molecules to see how they influence this movement.


1. Observe the movement of water across a membrane in model cells (decalcified eggs) and examine the environmental conditions that determine the direction of osmosis.

2. Compare the rate of osmosis when the concentration gradient varies.

3. Observe the effect of molecular size on the movement of solutes across a membrane.

4. Observe the effect of polarity on the movement of solutes across a membrane.

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Osmosis

When a selectively permeable membrane can inhibit the movement of some types of solutes and a concentration gradient exists, water will diffuse towards the higher solute concentration to equalize the concentration on both sides of the membrane. If you have a hard time remembering which way water moves in the presence of an osmotic imbalance (concentration gradient), just remember that SOLUTES SUCK! Water will always be drawn towards more concentrated solutes.

A solution can be described by its tonicity. Tonicity describes how a solution affects cell volume. A hypotonic solution will cause a cell to stretch and swell as water enters because it has a lower solute concentration (hypo = below) than a cell. A hypertonic solution will draw water out of a cell and make it shrink because it has a higher relative solute concentration(hyper = above). An isotonic solution produces no change cell volume because there is no difference in concentration (iso = same); an isotonic solution is said to be in osmotic equilibrium with the cell.

You have observed this phenomenon when your fingers get wrinkled after soaking in bath water, a hypotonic solution. Your skin wrinkles because the skin cells swell with water and your skin becomes too large to fit smoothly on your finger tips. Conversely, your skin may feel dry and tight after a day swimming in the ocean, a hypertonic solution, as the salt from the sea water draws water out of your skin cells.

In the following experiment, you will be using decalcified eggs as model cells. The eggs have been treated with vinegar to remove the calcium from the shell, leaving behind a membrane that is permeable to water (solvent), but not to other molecules (solutes).

Materials: • 5 decalcified eggs

• 5 weigh boats, one for each egg

• 3 Beakers or plastic containers with solutions A, B, & C

• 2 Beakers or plastic containers with solutions 1 and 2

• Paper towel

• Gram scale

Procedure IA: Determining the Tonicity of Extracellular Fluid: 1. Fill three beakers with enough of solution A, B, or C to cover an egg, about 300 ml.

2. Obtain three decalcified eggs. Gently dry and weigh each egg before immersing it in Solution A, B, or C. Dry the egg by gently rolling it on a paper towel. Do not dry the egg for too long because the paper towel will begin to draw out water from inside the egg and will change the weight of the egg. Record the weight of each egg in Table 1.

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3. Let the three eggs soak in solutions A, B, and C for 20 minutes. Go on to Procedure IB while you are waiting. The soak time should be at least 20 minutes, but can be longer if it is more convenient.

4. After at least 20 minutes, dry and weigh each egg and record your results. Use a "+" sign to indicate an increase in weight and a "–" sign to indicate a decrease in weight.

5. The change in weight reflects the movement of water into or out of the egg. Based on the movement of water, determine if the Solutions A, B, and C are hypotonic, isotonic, or hypertonic.

Table 1. Weight of Eggs and Tonicity of Solutions A, B, and C

Weight Before Soaking (g)

Weight After Soaking (g)

Difference in Weight (g)

Tonicity of Solution (hyper-, hypo-, or iso-)

Egg in Solution A

Egg in Solution B

Egg in Solution C

Questions:

1. Compare the presoak weight for eggs A, B, and C with their weights after the 20 minute soak. What is the tonicity of each solution?

2. Explain the physiological cause of the change in the weight of each egg.

3. What physical conditions are required to cause the water to move in a particular direction, into or out of the egg?

4. Explain why osmotic homeostasis must be closely regulated for the all the body fluid compartments.

Procedure IB: The Rate of Osmosis

The rate of osmosis is dependent on the difference in solute concentration across the membrane. Water will diffuse more quickly if the concentration gradient is steeper (the difference in concentration is greater). In this experiment, you will be soaking your eggs in two different hypotonic solutions and measuring the rates of osmosis.

1. Gently dry and weigh the last two eggs. Record the results in Table 2 as your zero time points.

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2. Immerse one egg in solution 1 and the other in solution 2. Dry and reweigh each egg after two minutes. Return the eggs to their solutions after weighing. Reweigh the eggs every 2 minutes for about a half hour.

3. Plot your results on graph paper (not binder paper!). Plot the data for both solutions on the same graph using different symbols or colors. You do not need to begin your Y-axis at zero.

4. Compare the rate of water movement into each egg by calculating the slope of each line using the formula below, where ∆Y is the change in the egg weight and ∆X is the change in the time.

slope = ∆Y

∆X

Table 2. Weight of Egg every 2 minutes in Solutions 1 and 2

Solution 1

Solution 2

Time (minutes)

Egg Weight (grams)

Change in Wt (Relative to 0 Time

Point)

Egg Weight (grams)

Change in Wt (Relative to 0 Time

Point) 0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30

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Questions

5. How does the rate of osmosis differ for the two solutions?

6. Which solution is more hypotonic, solution 1 or 2?

Membrane Permeability

We have just observed how osmosis can change the volume of a "cell" depending on the tonicity of the external environment. Water moves until either osmotic equilibrium is achieved or osmotic pressure prevents further osmosis from occurring.

The diffusion of solutes into or out of a cell also influences osmosis. Diffusion of a solute will occur if 1) there is a concentration gradient and 2) if a solute is penetrating, or able to cross the membrane. As a penetrating solute enters a cell, water follows the solute to maintain osmotic balance and the volume of the cell increases. If a solute is nonpenetrating, or unable to cross the membrane, then osmolarity does not change and osmosis does not occur. Figure 1 illustrates this principle.

Penetration, however, is dependent on the permeability of the cell membrane to a particular solute, which is in turn dependent on the physical characteristics of that solute. In this part of today's exercise, we will examine how solute size and solute polarity influence membrane permeability.

Figure 1. A cell is placed in an isotonic solution (a). Cell volume does not change initially (b), but the penetrating solute diffuses into the cell and changes the intracellular osmolarity (c). Water follows the penetrating solute to maintain osmotic balance, increasing cell volume (d).

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The Effect of Molecular Size

In order to study the effect of molecular size on membrane permeability, you will place red blood cells into isosmotic solutions of alcohols of increasing molecular size. If an alcohol molecule is able to penetrate the membrane, diffusion will occur due to the concentration gradient and water will follow the alcohol molecule. The volume of the cell will increase until hemolysis occurs (the cell bursts open). The amount of time required for hemolysis to occur depends on how easily the solute can penetrate the cell membrane. Since these solutions are isosmotic, no lysis will occur if the alcohol molecules are unable to cross the membrane.

We will use three different alcohols: ethylene glycol, glycerol, and ribose (Figure 2). They differ in size, having 2,3 and 5 carbon atoms respectively. Any difference in observed hemolysis time must be due to differences in size.

Ethylene Glycol Glycerol Ribose

Figure 2. Molecules of about the same polarity (one OH per carbon), but increasing size (number of carbon atoms).

A Note to Clarify the Difference Between Isosmotic and Isotonic: The suffix

"osmotic" is used to describe the relative concentration of a solution compared to a cell when the solute is penetrating, or able to enter the cell. The suffix "tonic" is used when a solute is nonpenetrating. More about this later.

Materials and equipment • Isosmotic Alcohol solutions (0.3M): Glycerol; Ethylene Glycol; and Ribose.

• Horse blood diluted with saline. Mix well before taking a sample!

• Disposable latex or nitrile gloves

Procedure IIA 1. Obtain one tube each containing 3 ml of Glycerol, Ethylene Glycol and Ribose solutions.

2. To each tube add two drops of horse blood. Mix by gently finger vortexing (instructor will demonstrate). Record the time the blood is added in the second column of Table 3.

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3. The tubes should be cloudy immediately after the addition of blood. As hemolysis occurs, the solutions will become transparent.

4. Record the time when the tubes become transparent in the third column of Table 3.

5. Calculate the "Hemolysis Time" by subtracting the time the blood was added from the time the tube became transparent and record the number of minutes and seconds

Table 3. Effect of Molecular Size Data

Molecule Time blood was added

Time solution became transparent

Hemolysis Time Carbon Atoms (size)

Ethylene Glycol 2 Glycerol 3 Ribose 5

The Effect of Molecular Polarity

Polarity is a chemical property that affects a molecule's solubility. Remember from our review of chemistry that polar molecules are more water soluble because water is itself a polar molecule. The uneven distribution of electrons create some areas on a molecule that are more negative which are balanced by other areas on a molecule that are more positive. The more negatively charged areas tend to associate with the –H side of H2O, while the more positively charged areas tend to associate with the –OH side of H2O.

What does water solubility have to do with membrane permeability? Remember that cell membranes are composed of a lipid bilayer. A lipid bilayer is more permeable to hydrophobic, or fat soluble molecules. Conversely, a lipid bilayer is less permeable to hydrophilic, or water soluble molecules.

Molecules with –OH groups, like alcohols, tend to be polar because oxygen is such an electronegative atom. The number of –OH groups on a molecule affects the degree of polarity a molecule will exhibit. Examine the structures of Propyl Alcohol and Glycerol in Figure 3. Note that both have a backbone of three carbon atoms and about the same overall size. The main difference between them is the number of polar –OH groups: Propyl Alcohol has only one (C3H7OH), while Glycerol has three (C3H5OH3), one for each carbon atom. This makes Glycerol a much more polar molecule than Propyl Alcohol. One would predict that cell membranes would be less permeable to more polar or hydrophilic molecules. These molecules will have a harder time crossing the membrane and have a longer hemolysis time.

Propyl Alcohol Glycerol

Figure 3. Molecules of about the same size (3 carbons), but increasing polarity (more OH groups)

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Procedure IIB 1. Obtain two test tubes. Add 3 ml of Propyl alcohol to one and 3ml of Glycerol to the other.

2. To each tube add two drops of horse blood and mix by finger vortexing. Record the time the blood is added in Table 4.

3. The tubes should be cloudy immediately after the addition of blood. As hemolysis occurs, the solutions will become transparent.

4. Record the time when the tubes become transparent.

5. Calculate the "Hemolysis Time" by subtracting the time the blood was added from the time the tube became transparent and record the number of minutes.

Table 4. Effect of Molecular Polarity Data

Molecule Time blood was added

Time solution became transparent

Hemolysis Time Polar OH Groups

Propyl Alcohol 1 Glycerol 3

Questions:

6. How do ethylene glycol, glycerol, and ribose molecules differ from each other?

7. How do propyl alcohol and glycerol differ from each other?

8. What is the purpose of having these solutions isosmotic?

9. What causes a blood solution to go from cloudy to clear?

10. Based on your data, how does a molecule’s size affect its ability to cross a cell membrane?

11. Based on your data, how does a molecule’s polarity affect its ability to cross a cell membrane?

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Exercise 4: BioPac Tutorial - Physiological Instrumentation

Understanding the methodology for obtaining physiological data is as important as understanding the physiological concepts themselves. This exercise is designed to familiarize you with the computer interfaced data acquisition unit called Biopac, equipment we will be using for many of our coming labs. Today, we will be using the physiological concepts we learned previously as an application for this technology. BE SURE TO BRING THIS TUTORIAL FOR FUTURE REFERENCE TO ALL LABS THAT USE THE BIOPAC WORK STATIONS.

Various physiological events can be measured by connecting a human subject to the computer using a device called a transducer. A transducer converts mechanical events into electrical signals. Each type of physiological event uses a different transducer. Today, we will be using a transducer called a plethysmograph, which detects pulse. Later on, we will be using other transducers, depending on the type of physiological activity we want to study. A transducer sends its electrical signal to the Biopac MP35 Acquisition Unit, otherwise known as the Biopac unit or the blue box, which amplifies these electrical signals and displays them on the computer screen. The Biopac Student Lab software allows you to control how the electrical signals are displayed and to analyze the data you collect. Lab partners should take turns using the equipment so that everyone becomes familiar with using both the hardware and the software.

A plethysmograph responds to the surge of arterial blood that occurs in your fingertip due to the contraction of the heart. The photoelectric plethysmograph we will be using emits a beam of light that is then detected by a light sensor. The amount of light reflected back from your finger increases when ventricles contract and blood flow to the finger increases. This change in light is converted into an electrical signal that is plotted as a function of time on the computer display.


1. Learn how to use the Biopac workstation.

2. Record pulse using a plethysmograph pulse transducer.

3. Compare the results of the aerobic step test, using manual versus computerized data collection.

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Equipment List

√ Computer

√ Biopac MP35 Acquisition unit

√ Plethysmograph pulse transducer plugged into Ch. 2

√ Step stool

√ Metronome

√ Biopac Student Lesson L07-ECG&P-1 (or L07 ECG & Pulse for PC's)

Getting Started

The equipment for the day's experiment will usually be ready to go when you come into lab, but you should always double-check your equipment yourself before starting. Here is a checklist of the basic set-up when using the Biopac data acquisition units:

Figure 1. "On" button for MacIntosh computers

1. Turn the computer on. For Mac laptops, press the on silver "ON" button (see Fig. 1) located above the computer keyboard. For PCs, the "ON" button is found on the front of the tower. Login using the password obtained from the instructor.

2. Check that the Biopac unit (blue box) is connected to the computer via a black USB cable.

3. Check that the plethysmograph (SS4LA) is plugged into the front of the Biopac unit in Channel 2 (see Fig. 2).

Figure 3. BSL application icon

Figure 2. Front panel of Biopac MP35 Acquisition Unit

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4. Turn the Biopac unit ON. The Power switch is on the back panel on the right. The down position is OFF. The up position is ON. Check that the AC adapter for the unit is plugged into the back. When the Biopac unit is initially turned on, two green lights on the right side of the front panel of the Biopac unit will light up. Do not launch the Biopac Student Lab application until the left "busy" light goes off. If you don't wait, the computer will not connect to the Biopac unit. If the busy light does not come on, proceed to the next step. If the busy light is flashing rather than steady, turn the Biopac unit off and on again. If the busy light does not go off, try turning the unit off and on again. If all else fails, ask for help from the instructor.

5. Launch the Biopac Student Lab software application. For Mac laptops, click on the Biopac Student Lab application icon in the dock (see Fig.3). The dock is the row of application icons found at the bottom of the screen (usually). For those of you unfamiliar with the MacIntosh operating system, ask the instructor or a Mac-proficient lab partner for help.

For PC users, click on the green "Start" button in the bottom left corner, go to "Programs", then "Science", then choose BSL Lessons 3.7.

6. If the computer connects to the Biopac unit without problem, go on to the next step. If you get a message that the computer cannot find the hardware, check the USB cable connecting the Biopac unit to the computer, all other power cables, the ON/OFF switch, and make sure the power light is on and the busy light is off. For Macs, the error message will look like Fig. 4. For PCs, the error message will look like Fig. 5.

Figure 4. "Can't Find the Hardware" warning for Macs

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Figure. 5. "Cannot Find the Hardware" warning for PCs

For Macs, if the Biopac unit is turned on properly, click on the pull-down menu in the warning box and choose "usbmp353B1" (see Fig. 6).

Figure 6: Pulldown menu to connect the Biopac unit to the computer

For PC users, click "Retry". If the computer recognizes the Biopac unit, continue on to the next step.

7. When everything is connected properly, a window will open asking you to choose a lesson. Select L07-ECG&P-1 (or L07 ECG & Pulse, for PC's) and click OK. Type in a file name for your experiment. Use a name that distinguishes both your lab group and the exercise you are performing, like "Barbara's Pulse", not just "Pulse". Write down the name of your file here:

_________________________________________. Get into the habit of writing this information down every time you open a new file.

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Recording A Pulse

1. When you enter a file name and click OK, the screen that opens will look like Figure 7:

Figure 7: Opening screen of Biopac Student Lesson L07-ECG&P-1

The upper Data window displays two channels, one for recording an electrocardiogram, which we will not be doing, and one for recording the pulse. These channels are labeled on the left side of the window. The bottom Journal window displays instructions we will not be using either. Ignore both the top channel and bottom window in today's exercise. We will be able to hide both of these once you have finished collecting your data.

2. To prepare your subject (lab partner), use the Velcro strap to attach the plethysmograph to the ventral surface (fingerprint side) of the index finger. It should be just snug enough to stay on the finger, but not so tight that it disrupts circulation (no purple finger tips!). The subject should rest her or his arm palm up on a table while sitting quietly. The hand should be relaxed.

WARNING: The plethysmograph is a fragile piece of equipment. All care should be taken to protect the detector window and the cord connections.

3. Click on the Calibrate button in the upper left corner of the screen. An error message will appear. Click Ignore.

Any transducer must be calibrated before it can be used. The calibration process determines the minimum (baseline) and maximum reference points. If the peaks of your calibration recording seem small, software will correct the recording to fit the screen nicely. Do not move while the computer calibrates the plethysmograph.

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4. If your subject moved during the calibration, you can redo it by clicking on Redo Calibration. If you got a recording of a steady pulse that looks like Figure 8, continue.

Figure 8. Calibration trace

5. Click the Record button to begin recording. You will get another error message; click Ignore. The recording of the pulse should appear, moving from left to right in the recording window. The recording is also referred to as a trace. The pulse trace is blue and the ECG trace (that you are ignoring) is a flat red line.

6. Experiment to see how moving your finger affects the recording. Click Suspend to stop recording and change the tightness of the strap. Click Resume to see how this affects the recording. Note that if the strap is too tight and blood can no longer flow into the finger, the pulse trace flattens. Click Suspend and readjust the plethysmograph to take a normal reading.

7. Continue recording for at least 20 – 30 seconds without moving until there is a good, steady pulse recording. Click Suspend to stop recording. If you want to record more, click Resume. Do NOT record more than a minute of data to avoid wasting memory space. If you want to erase your data and try again, click Redo. Your goal is to get at least six steady, uniform pulse beats in a row for analysis. When you are satisfied with your recording and are ready to analyze it, click Done.

8. When you click Done, you will be asked what you would like to do next. If another member of your group would like to record their pulse, select Record from another Subject. Otherwise, select Analyze Current Data File and click OK. If you want to analyze a previously recorded data file, choose Analyze a Different Data File. A finder window will open to the main Biopac folder. Select the Data Files folder, then your folder, then the file you want to analyze.

NOTE: If at any time you want to open a new file or go back to an old file, go to the Lessons menu. Choose the lesson to open a new file (for example, L07 ECG & Pulse) or go to the bottom of the menu and choose Review Saved Data.

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Navigating The Biopac Student Lab Software

The primary purpose of this exercise is to give you practice in collecting and analyzing data using the Biopac unit and Biopac Student Lab software. Take the time to familiarize yourself with the display tools and analysis functions. Becoming comfortable with the software is more important than calculating your heartbeat and getting to the end of the exercise. You will be needing these skills for future Biopac labs. Becoming proficient today will help you perform future exercises more smoothly and efficiently. The following steps are designed to help you manipulate the screen to optimize the viewing of your trace.

1. Editing and Selection Tools: There are three editing and selection tools, the Arrow, I-Beam, and Zoom. The icons to choose these tools are found in the bottom right corner of the data screen (see Figure 9).

Figure 9. Editing and Selection Tools. From left to right: the Arrow, the I-Beam, and Zoom.

The arrow is used to point and click, the I-Beam is used to highlight areas of the trace for analysis, and the zoom enlarges the trace.

2. To activate or hide a channel: Since we are only interested in the Pulse data we collected, we can hide the ECG channel and enlarge the Pulse channel. The Channel boxes are used to activate or hide the different channels and are found in the second row below the "Overlap" button in the upper left corner of your screen (for Macs, see Figure 10). In today's exercise, the Channel 1 box controls the ECG channel and the Channel 40 box controls the Pulse channel. Clicking on a channel box activates that channel so that editing and selection tools can be used on that trace. Values measured in an active channel will appear in the display windows found in the first row directly below the "Overlap" button. We will talk more about the display windows later.

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Figure 10. Upper left corner of analysis screen for Macs (left) and PCs (right). The square Channel boxes two rows below the "Overlap" button indicate the active channel. In the Mac example on the left, Channel 1 is inactive; the Channel 40 box has been selected (gray) and the Pulse Channel is the active channel. In the PC example on the right, Ch 1 has been hidden (X'ed out).

• Alternately click on the two channel boxes to see how the ECG and Pulse screens are activated. Note how the screen title running vertically on the left edge of the screen is highlighted when that channel is activated.

• Since we did not collect any ECG data, we do not need to keep Channel 1 on the screen. To hide this channel, Mac users hold down the option key and click on the Channel 1 icon. PC users hold down the Ctrl key and click on the Channel 1 icon. The channel box will have a slash through it when it is hidden (see the example on the right in Fig. 10). Hiding a channel does not prevent it from being an active channel. Displaying a channel does not make it an active channel unless the channel box is clicked. Be sure to leave the Pulse channel active once you hide the ECG channel.

3. Optimize the displayed channel: One of the most important things you need to learn how to do today is to optimize the way the trace is displayed so that your measurements can be as accurate as possible. The trace will be easier to analyze if the Pulse channel is enlarged to fill the entire screen. To do this, close the journal window at the bottom of the screen:

• Mac users click on the red button in the upper left hand corner of the bottom window (see Figure 11).

Figure 11. Window control buttons. The red button (far left) will close the window. The yellow button (middle) will hide the window in the dock. The green button will maximize the window so that it fills the entire screen.

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Be careful not to click on the red button of the data window at the top of the screen, as this will close your file. If this happens, click on the Lessons pull-down menu and choose Review Saved Data at the bottom of the list. Select the Data Files folder, then the folder you named, then the file you were working on (this is why you always write down your file names). Make the pulse channel fill the entire screen by clicking on the green button of the data window. Alternatively, click and drag the bottom right corner of the data window downward.

• PC users simply click on the upper border of the journal window and drag it down to the bottom of the screen.

4. Control the display of the X and Y axes: All of the data you collected will initially be displayed on a single screen, no matter how many minutes you recorded. The longer you record, the more squished together your pulses will appear and it can be difficult to place the cursor accurately when measuring your pulse rate. You will want to enlarge specific areas of your data for accurate analysis.

• Choose the zoom tool icon in the lower right hand corner of the screen. The zoom tool looks like a magnifying lens (see Fig. 9). Click and drag the zoom tool to draw a box around the specific area of the recording you would like to analyze. To undo the zoom, pull down the Display menu and select Zoom Back. There is also a Zoom Forward function to redo a zoom.

• You can also click the zoom tool anywhere on your trace to enlarge the entire display. The peaks will get higher and wider and some of the trace may be off screen; the scroll bar should now be blue. To adjust your trace so that the peaks and valleys are visible, pull down the Display menu and select Autoscale Waveforms. This automatically adjusts the Y-axis so that the highest and lowest points are on screen. If your pulses are still too squished together, click on the trace again to zoom in further. Readjust the Y-axis by selecting Autoscale Waveforms again.

Note the Autoscale Horizontal function in the Display menu. This readjusts the X-axis so all the data you collected is once again displayed on the screen.

• You can move to different locations along your trace by either clicking and dragging the blue scroll bar or by clicking the left and right arrows (see Fig. 12).

Figure 12. Horizontal scroll bar and arrows from the lower right side of screen.

• There is also a vertical scroll bar along the right edge of the screen. The vertical scroll bar can be used to reposition the trace on the screen if necessary.

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5. Show or hide grids: Grid lines give you a visual reference point when looking at a trace. To add grid lines to your screen, pull down the File menu and select Display Preferences. Choose "Show Grids". To adjust the grid lines of the X-axis (time), click anywhere in the margin below the display. A dialog box will open (see Figure 13).

Figure 13. Horizontal Scale dialog box

Enter the number of seconds you would like between grid lines for Major Division. Increments of 1, 2 or 5 seconds are usually appropriate. Click OK. The minor divisions cannot be adjusted.

• To adjust the grid lines of the Y-axis (mV), click anywhere in the margin along the right side of the screen. A similar dialog box will open. Look at the units of the Y-axis to determine the Major Divisions. If the maximum and minimum Y values are around 1 mV, then choose 0.1 or 0.2 mV increments for your grid divisions.

Analyzing Your Data

Now that you know how to adjust the display settings, practice using what you learned by calculating your subject's heart rate.

1. Display all the data you collected by pulling down the Display menu and choosing first Autoscale Horizontal and then Autoscale Waveforms. Your trace should be all squished

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together again. If the trace is too squished to distinguish the peaks (this means you have collected A LOT of data), use the zoom tool and the Autoscale Waveforms function to adjust your trace so that it can be more easily reviewed.

2. Select five pulse peaks that appear to be steady and regular and use the zoom tool to click and drag a box around those peaks. Be sure to include the lowest point before the first peak and the lowest point after the last peak. All five peaks should be displayed together now. See Fig. 14 for an example.

3. The I-Beam is used to highlight areas of the trace you want to measure. To highlight a region of your trace:

• Click on the I-Beam icon in the bottom right corner of the screen (see Fig. 9). Highlight a pulse peak valley-to-valley by positioning the I-Beam at the lowest point of the trace in front of the pulse peak you want to measure; click and drag the I-Beam until the I-Beam is positioned at the lowest point of the next pulse peak; release the mouse. See Fig. 14 for an example of a highlighted peak.

Figure 14. Screen shot of five pulse peaks with one peak highlighted.

Figure 15. The measurement region is along the top of the data window and directly below the "Overlap" button. In this example, the channel select box is set to Channel 40, the measurement function box is set to Delta T (the highlighted time interval), and the reading in the result box is 0.815 sec.

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4. Values that correspond to the highlighted areas appear in the result box in the measurement region found at the top of the screen (see Fig. 15). The measurement region will allow you to make up to five different measurements or calculations from the data you have highlighted. Use the following settings to calculate heart rate:

• The channel select boxes allows you to choose the channel you want to take your measurements from. Do not confuse the channel select box with the channel boxes directly below. We used the channel boxes previously to hide or activate a channel (see Fig. 10). The channel select boxes are in the row directly above the channel boxes. In today's lab, all your measurements and calculations will be from Channel 40. Change the first two measurements from channel 1 to channel 40 by clicking on both the first and the second channel select boxes and choosing Ch40, Pulse from the pop-up menu. You can tell which channel your box is set to from both the label on the channel select box and from the color of the box. The box will match the color of the trace.

• The measurement function box allows you to choose the variable to be measured or the calculations you would like the computer to perform. Today we want to measure the time interval of each pulse beat, so choose Delta T from the middle of the pop-up menu (see Fig. 16). Note that Delta T is different from Delta at the top of the menu. Delta T measures the highlighted distance along the X-axis, while Delta measures the highlighted distance along the Y-axis.

• The Biopac software can also calculate heart rate based on the Delta T measurement, so set the second measurement function box to BPM (beats per minute). We won't be using the other result boxes today.

5. Record the Delta T and the BPM for this first pulse beat in Table 1 (page 14), using whole numbers for the heart rate. This calculated heart rate is based on a single beat. Compare the calculated heart rate of several different pulse beats by highlighting three other pulse beats on your screen. Record this data in Table 1 as well. Do these values differ at all? How would you decide which value to use if there are differences?

• Since individual pulse beats can vary in length from beat to beat, calculating an average based on several beats is a more reliable way of measuring heart rate. Highlight all five pulse beats using the I-Beam. Delta T now represents the time of five pulses, not one, so the BPM calculation in the second results box is no longer correct. Use the formula below to calculate BPM based on five pulse beats:

Heart Rate (beats/min) = Number of beats X 60 sec.

Delta T 1 min.

Figure 16. Measurement function box pop-up menu.

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How does the BPM averaged from five pulse beats compare to the BPM calculated from a single beat? How does this formula compare to the formula we used to calculate heart rate earlier in the semester?

Pulse bePulse bePulse bePulse beFor 5 b

A

Earlier in the semestecapacity using the McArdle-Kgroup who performed the Stemeasure pulse. This will allowBiopac unit. We will not be m

1. Attach the plethysmograIMPORTANT: The pletshould be loose enough tgive a good pulse reading

2. Open a new file by going& Pulse for PCs). Nameof your subject for aboutrecording. Go back to thegetting started. Slip the pfastener. No recordings wrecording.

3. Have your subject perfor

• Set the metronome to 88can turn off the sound of

August 2011

Table 1. Resting Pulse Data

Delta T (sec.) BPM

(beats/min)

at #1 at #2 at #3 at #4 eats

erobic Capacity Revisited

r, we measured our pulse rates manually to estimate our aerobic atch Bench Stepping Test. If possible have a member of your

p Test previously repeat the Step Test using the plethysmograph to you to compare data collected both manually and using the easuring recovery heart rate over time this week.

ph to the finger of the subject who will do the step test. hysmograph should not be wrapped tightly around the finger. It o slip off the finger without readjusting the velcro fastener, yet still during the calibration.

to the Lessons menu and choosing L07-ECG&P-1 or (L07 ECG your file, calibrate the plethysmograph and record the resting pulse 15 seconds to get a good baseline reading. Click Suspend to stop instructions for "Recording A Pulse" if you need a reminder for lethysmograph off the subject's finger without loosening the velcro ill be made during exercise because the movement affects the

m the Step Test:

beats per minute for females or 96 beats per minute for males. You the metronome and use the flashing light to keep time if you prefer.

page 45

• Brace the step stool backwards against the wall to prevent it from slipping. Facing the back of the step stool, step up to the top step of the stool. Use the same four-step cadence you used last week (up right foot, up left foot, down right foot, down left foot) for a period of three minutes, stepping in time with the metronome. LISTEN CAREFULLY TO THE METRONOME TO MAINTAIN THE CORRECT PACE.

4. Immediately after 3 minutes of stepping, slip the plethysmograph back on the subject's finger and click Resume to begin recording the pulse while the subject stands quietly and rests their arm on the tabletop. It is crucial that the recording begin as soon as the individual stops exercising.

5. Record the pulse for about 30 seconds, the click Done. Select Analyze Current Data File and click OK.

Using Markers To Navigate Your Recording: 1. Markers are used to reference important events or locations in the data. The marker bar is

the white bar located at the top of the screen just below the measurement region (see Fig. 17).

Figure 17. The white marker bar is located below the measurement region. Note the marker

tools located at the far right side of the marker bar (see Fig. 18 for a close up).

2. There are two types of markers:

a. Append markers. These markers are automatically inserted when you begin a new recording. They appear as a black diamond above the text box and appear blue when active. The marker will always be identified by text. Ex: “seated and relaxed”.

b. Event markers. These markers can be manually entered during a recording by pressing either the ESCAPE key (Macs) or F9 (PCs). They appear as inverted triangles below the marker text region and are yellow when active. You can manually add the text you wish in the text box above the inverted triangle by double clicking on a selected marker. Ex: “Robin's pulse post-run”.

You can add an event marker after recording by clicking in the marker region with the selection tool (arrow). The new marker becomes the current active marker and you may label it by typing in the marker text box.

3. The marker tool allows you to navigate from marker to marker without tediously scrolling through your entire recording. The marker tool is located at the far right side of the marker bar. See Fig. 18 to see what it looks like.

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Figure 18: The Marker Tool

Using the marker tool to find a specific marker in your data: a. Click on the right pointing marker tool to reach the next marker to the right of the

data section you are looking at.

b. Click on the left pointing marker tool to reach the previous marker to the left of the data section you are looking at.

c. Click on the downward pointer marker tool at the far right end of the marker region. This will generate a pop-up menu. Select the Find option. Enter the name of the marker (Ex. “Robin's pulse post-run”). Click Find next and the selected marker will light up on the screen. If you wish to see all markers and their labels listed, select Marker Manager. This will list all markers with their labels and time stamp.

Data Analysis 1. If necessary, adjust the trace so that pulse peaks can be easily measured. Scroll along the

trace until the 15 second point is reached. This should be approximately where recording resumed after the Step Test was completed. The exact point when recording was resumed will have been marked by the diamond shaped marker found just below the measurement region at the top of the data window (see Fig. 19).

Figure 19. Diamond-shaped marker below the measurement region.

2. Use the zoom tool (see Fig. 8) to select five pulse peaks beginning at 5 seconds after the Step Test was completed. These pulse peaks may be irregular and jagged since the subject was recovering from exercise. If the peaks are difficult to distinguish, choose the closest peaks that are readable for analysis.

3. Use the I-Beam to highlight five pulse peaks and measure Delta T. If the valleys between pulse peaks are difficult to distinguish, the peaks can be used instead. Be careful to count to correct number of pulses that are highlighted. Calculate heart rate using the formula above.

4. Estimate maximal oxygen consumption rate (VO2 max) using the following equations.

Females:

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VO2 max = 65.81 – (0.1847 X HR)

Males:

VO2 max = 111.33 – (0.42 X HR)

Use the standards from Tables 2 or 3 from Exercise 2: Aerobic Capacity to determine fitness level based on estimated VO2 max.

Table 2. Aerobic Capacity Delta T (sec) Heart Rate (BPM) At rest

After exercise

VO2 max in ml/kg/min (from formula)

VO2 max from previous Step Test

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Questions:

1. Define and describe the function of the following equipment: transducer, amplifier, plethysmograph.

2. How does the plethysmograph that you used detect pulse?

3. What should you do if you get a dialog box that says "I can't find the hardware."?

4. What is the purpose of calibrating a transducer?

5. What are the functions of the arrow, the I-Beam, and the zoom tools?

6. How do you adjust the X and Y axes in the data window (how do you adjust the way the trace is displayed)?

7. How do you choose the data you wish to view in the data window?

8. Where is the measurement region? Where is the channel select box, the measurement function box, and the result box and what are they used for?

9. How do you insert a marker into your trace, both while you are recording and if your recording has already been made? How do you use the marker tool?

10. Be able to calculate a heart rate from a trace using the formula provided.

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August 2011 page 50

Exercise 5: Electromyography Readings: Silverthorn, 4th ed. pg. 407 – 412; 5th ed. pg. 417 – 42

Your brain communicates with your muscles through action potentials on the motor neurons, which are then transmitted across the neuromuscular junction to the surface of the muscle fiber. The muscle fiber action potential triggers the release of calcium into the cytoplasm, which in turn results in the initiation of the actin-myosin sliding filament mechanism responsible for muscle contraction. The coordination of these electrical and mechanical events is called excitation-contraction coupling (see Fig.1). A single contraction-relaxation cycle in a skeletal muscle fiber that results from a single action potential is known as a twitch.

Figure 1. Excitation-Contraction Coupling (Fig. 12-11, pg. 400 from Silverthorn)

There are two ways that the force of a muscle contraction can be increased. One way is called summation. During normal voluntary contraction, the motor neuron releases a volley of action potentials, rather than one at a time. Increasing the frequency, or rate, of these action potentials from a single motor neuron results in an increase in the force of muscle contraction, or summation.

The second way to increase the force of muscle contraction is called recruitment and involves increasing the number of motor units that contract simultaneously in a whole muscle. Each motor unit is composed of many muscle fibers that are innervated by a common motor neuron. More motor neurons firing simultaneously will cause more motor units to contract together, which produces a greater force, or recruitment. We will be learning more about both summation and recruitment in next week's lab exercise, but this week we will focus only on recruitment.

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Each motor unit is made up of one of three muscle fiber types: slow twitch fibers, intermediate twitch fibers, and fast twitch fibers. Each fiber type has its own advantages and disadvantages. Slow twitch fibers are resistant to fatigue, but develop tension the slowest (hence the name) and are the weakest. Fast twitch fibers fatigue more easily, but develop tension more quickly and produce more force. Slow twitch fibers are able to sustain a lower tension for longer periods of time, while fast twitch fibers produce stronger contractions for shorter periods of time.

If a voluntary muscle contraction is sustained for a long enough period of time, however, even slow twitch muscle fibers will begin to tire. As the tension produced by these motor units decreases with fatigue, stimulus from the brain will increase and result in the recruitment of more and more motor units, until all the motor units in a muscle are being used.

Recruitment of motor units always occurs in the same sequence: slow twitch motor units are recruited first, then intermediate twitch motor units, then fast twitch motor units. The motor neurons of slow twitch motor units have the lowest threshold and so are the first to respond to a weak stimulus. As the stimulus increases (ie. the brain tells the muscle to produce more force), threshold is reached next for the intermediate twitch motor units, then finally the fast twitch motor units.

This increase in motor unit activity and the corresponding electrical activity on the sarcolemma of the muscle fibers can be detected by electrodes placed on the skin. A recording of this electrical activity is called an electromyogram or EMG. In an unfatigued muscle, the electrical activity is proportional to force production. In other words, the recorded EMG signal reflects motor unit recruitment and summation taking place within the muscle tissue; greater force is reflected by an increase in the EMG activity. An EMG is a quantitative measure of muscle activity for a given workload. As fatigue sets in, however, the relationship between force and electrical activity changes. An action potential on the muscle fiber will not produce greater force if the muscle fiber is fatigued.

Electromyography can be used to diagnose the cause of muscle weakness. Electromyography can help to differentiate primary muscle conditions, such as muscular dystrophy, from muscle weakness caused by neurological disorders, such as multiple sclerosis. EMG is also used in studies of kinesiology and exercise physiology.

In today's lab, we will be using the EMG to observe electrical activity of your arm muscles and the resulting motor unit recruitment. Keep in mind that the electrical activity we are detecting is primarily the result of action potentials generated on the sarcolemma of the muscle fibers, not the electrical activity of the motor neurons. Consider the difference in surface area between a motor neuron and the muscle it innervates.


1. Investigate the EMG response in the forearm flexors to increasing workloads.

2. Investigate the EMG response to fatigue when the muscle is fully recruited.

3. Investigate the EMG response to fatigue when the muscle is partially recruited.

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Getting Started

1. The equipment for today’s experiment is listed below. You should be able to explain the purpose of the dynamometer, electrode leads, and electrodes.

• Computer

• Biopac MP35 Acquisition unit

• BIOPAC Hand Dynamometer plugged into Ch. 1

• BIOPAC electrode lead set plugged into Ch. 3

• Disposable paste-on electrodes

• Biopac Student Lesson L02-EMG-2 for Macs or L02 EMG II – Electomyography for PCs

A dynamometer is a device that measures force. In this case, the force generated by your hand when the dynamometer is squeezed is converted to electrical activity by the transducer. The unit of force is kilograms.

2. Use the checklist below to ensure that the basic set-up is working:

• Turn the computer on. For Mac laptops, press the on silver "ON" button (see Fig. 1) located above the computer keyboard. For PC towers, push the large button on the front of the tower.

• Check that the Biopac unit (blue box) is plugged in to a working electrical outlet.

• Check that the Biopac unit is connected to the computer via a black USB cable.

• Turn on the Biopac unit AFTER the computer has finished booting up. The Power switch is on the back panel on the right. Check that the AC adapter for the unit is plugged into the back. Wait until the "Busy" light stops blinking before launching the Biopac Student Lab 3.7 (Mac) or BSL Student Lessons 3.7 (PC) application (see Step 5 below).

3. The subject should remove all jewelry from wrists and ankles. Use the dominant forearm for the first part of the experiment. (“Dominant” refers to the right hand if the individual is right-handed, or the left hand if they are left-handed.)

Clean and scrub three regions on the forearm for electrode attachment using an alcohol pad. The first area is about 2 inches below the anterior cubital fossa of the elbow over the mass of forearm muscles that originate on the medial epicondyle of the humerus (Remember your anatomy?). If you aren't sure where this is, clench your fist to make the muscle bulge. The second and third are distal, over the lateral and medial anterior surface of the wrist. Let the areas dry after scrubbing with the alcohol pad. Refer to Fig. 2 for placement of the elctrodes.

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Figure 2. Electrode and Lead Placement

4. Snap the ends of the three color-coded electrode cables onto the disposable electrodes attached to the subject's arm:

• the white“VIN-” lead is attached to the electrode near the elbow.

• the black “GND” (ground)lead is attached to the electrode on the medial (little finger) side of the wrist.

• the red “VIN+” lead is attached to the electrode on the lateral wrist.

The pinch connectors work like a small clothespin and latch onto the nipple of the electrode. Be sure the metal backing of the connectors is in contact with the metal nipple.

5. Launch the Biopac software application. For Mac laptops, click on the Biopac Student Lab application icon in the dock (see Fig.3). The dock is the row of application icons found at the bottom of the screen (usually). Choose lesson “L02-EMG-2” from the menu. For PCs, click Start (lower left corner of screen) > Programs > Science > BSL Student Lessons 3.7. Choose "L02 EMG II – Electromyography".

6. A prompt will appear that asks you to type your file name in the box. Choose a name that identifies and distinguishes you from others, e.g. janedoe_EMG.

7. Once the BioPac data collection window appears, you will be prompted by directions in the lower part of the screen to calibrate the dynamometer. Note that we will not be using headphones to listen to the EMG. Place the dynanometer on the table and click the “Calibrate” button in the left upper corner of the screen. This sets the zero baseline. Pick up the dynanometer with the short grip bar against the palm of your hand. See Figure 3.

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Figure 3. Grip position for the dynamometer.

Following the prompts, wait 2 seconds then squeeze the dynanometer as hard as you are able for 2 seconds, then release the dynanometer and allow the calibration to finish. This sets the highest possible value. The Calibration recording should look something like Figure 4. If it doesn’t, then repeat the calibration.

Figure 4. Calibration Screen.

The software sets the scale of the display based on your calibration grip force so that the recording fits nicely in the window.

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Muscle Fiber Recruitment With Increasing Force; Muscle Fiber Recruitment and Fatigue

You will collect all the data for both the increasing force demonstration and fatigue experiment before you analyze them. The first two segments record data from the dominant forearm while the second two record on the non-dominant forearm. This means that you don’t have to switch the leads back and forth.

• First segment: Demonstrates motor unit recruitment as the dynamometer is squeezed or clenched with increasing force on the dominant forearm.

• Second segment: Demonstrates fatigue in dominant forearm.

• Third segment: Repeats segment one, but on non-dominant forearm.

• Fourth segment: Repeats fatigue experiment (Second segment) on non-dominant forearm.

1. When you click Record (upper left corner of screen), the recording will begin and an append marker labeled “Forearm 1, Increasing clench force” will be added. The screen will change to display only the hand dynamometer channel, and a grid will appear so that you can visually review the force level.

2. You are going to make a series of four or five clenches of increasing strength. During the calibration process, the software determined your Assigned Increment of Force, or the amount of force by which each clench should increase. If your maximum calibration force was 0 – 25 kg, then each clench should be increased by 5 kg. If your maximum force was 25 – 50 kg, then increase each clench by 10 kg. If your maximum force was more than 50 kg, then increase each clench by 20 kg.

3. If your Assigned Increment of Force was 5 kg (your max was less than 25 kg), wait 2 seconds, then begin with a gentle clench of 5 kg that lasts two seconds, then release for two seconds before squeezing again. Increase the strength of each clench by your Assigned Increment, repeating the 2 second cycle of clench and release; i.e. hold the clench for 2 seconds and release for two seconds before beginning the next cycle. Try to maintain an even, sustained force that produces flat peaks. When you have recorded 5 clenches, click Suspend. If all goes well, your data should look something like Figure 5. The last clench should be at maximum strength. If all did not go well, you can redo your recording. Click Redo, then Yes when the computer asks if you are sure, then Resume. But don't tire out your muscles - save some strength for the next segment!

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Figure 5. EMG recording for clenches of increasing strength.

4. In this next recording (Segment 2) you will hold a single sustained clench for as long as you can stand it. This experiment will demonstrate how the action potentials from the brain respond when a muscle begins to fatigue. The window at the bottom of the screen will tell you to clench the dynamometer with your maximum force. Click Resume to begin recording, then squeeze as hard as you can. Make a note of your maximum force. Continue to squeeze until your muscles fatigue to the point of quivering. Click Suspend to stop recording when you have had enough.

Non-Dominant Arm 1. Attach three new electrodes to the opposite arm.

2. Repeat steps 1 through 3 above, collecting data for increasing clench strength. Make a note of the force you generated in your last and hardest squeeze. This is your maximum force for your nondominant hand.

3. In this last recording (Segment 4), when the window at the bottom tells you to clench with maximum force, DO NOT DO THIS. Instead, you will clench the dynamometer with only HALF of your maximum force. For example, if your maximum clench in the previous recording was 30 Kg, maintain a clench of 15 Kg for this exercise on fatigue.

Once you have determined how much force you should exert for the prolonged clench, click Resume. The recording will continue and a marker labeled “Forearm 2, Continued clench at maximum force” will be automatically inserted. Watch the screen to maintain the clench at half-maximum until your muscles fatigue to the point of quivering. Click Suspend when your arm fatigues.

It is very important that you maintain the same force throughout and that you do not stop until your arm is too fatigued to continue squeezing.

4. Click Done when you finish recording the last segment. We won't be listening to the EMG with headphones. When the dialog box asks if you are sure, click Yes, then choose "Analyze Current Data File".

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Analysis of the Data

Refer to the “Physiological Instrumentation Tutorial" from last week that introduced you to the Biopac equipment to refresh your memory about how to navigate the analysis functions. Set up your display for analysis:

Three channels will be displayed, Force (Ch 1), EMG (Ch 3), and Integrated EMG (Ch 40). PC users note that an additional scroll bar will above the segment labels at the top of Channel 1 display.

• Hide the EMG display (red trace) using the square channel boxes in the upper left hand corner of the display (see page 44 of the Tutorial). Mac users hold down the option key and click the Ch 3 box. PC users, hold down the Ctrl key and click the Ch 3 box.

• Close the journal window at the bottom of the screen by clicking on the red button in the upper left hand corner of the bottom window (Mac users) or by dragging the top of the Journal box to the bottom of the screen (PC users). Your display should look like Fig. 6.

Figure 6. Force and EMG Integral channels displayed.

• Select the channels and functions you want to analyze in the first three measurement regions (see pg. 48 and 49 of the Tutorial):

1st: Ch 1 (blue), function: Mean

2nd: Ch 40 (green), function: Mean

3rd: Ch 1 (blue), function: Delta T

Increasing Clench Force

1. For the first part of the analysis, use the data labeled “Increasing clench force” recorded in Segment 1 for the dominant forearm.

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Click on the zoom tool, located at the right lower corner of the screen. Select the clench data from segment 1 by drawing a box around all five force peaks.

You may need to adjust the trace so that the peaks and valleys are visible. Use the Display menu and select Autoscale Waveforms to enhance the visibility of the data.

2. Use the I-Beam cursor to select a flat area on the top of the plateau phase of the first clench (Figure 7).

Figure 7. Plateau of first clench selected.

3. Record the mean Force and mean EMG Integral values for each clench in Table 1 below. These values will be found in the measurement function boxes above the marker region in the data window. “Mean” displays the average value in the selected area. You will be using the mean of the force exerted on the dynamometer by the clench (kg) and the mean of the integrated EMG (mV-sec).

Table 1: Increasing Force of Separate Clenches

Dominant Arm (Mean) Non-Dominant Arm (Mean)

Clench FORCE (kg)

EMG Integral (mV-sec)

1

2

3

4

5

Clench FORCE (kg)


1

2

3

4

5

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4. Repeat the measurements for segment 3, "Increasing clench force with the non-dominant arm".

5. Using the data recorded in Table 1, make a graph of the relationship between the force production and the electrical activity by plotting the Force on the (x) axis and the EMG Integral on the (y) axis. This arrangement may seem counterintuitive, but in this particular experimental design, your brain is determining the force. This becomes the independent variable. The dependant variable is the response being recorded and measured, which is the EMG Integral. Don't forget to include the units in your labels. Plot one line representing the dominant arm and the other line representing the non-dominant arm. Distinguish the lines by making them different colors or making one dotted and providing a key.

6. Calculate the slope of each line using the formula below, where ∆Y is the change in the EMG Integral and ∆X is the change in the Force.

slope = ∆Y

∆X

Questions

1. Describe the molecular events that occur along the path of electrical stimulation from the motor neuron to the sarcolemma of the muscle fiber. In other words, describe the mechanism of excitation–contraction coupling.

2. What is the source of the electrical signal picked up by electrodes on the subject's skin? What is an EMG? Why is the EMG Integral used to determine the mean, rather than the actual EMG?

3. Based on the graph you made, what conclusions can you draw about the relationship between Force and electrical activity (EMG Integral) for each of the five increasing clenches? What conclusions can you make about the difference between your dominant and non-dominant hand strength?

4. Define recruitment. Discuss the relationship between recruitment, motor units and your data. In other words, how did the data you collected demonstrate the concept of recruitment? You may need to consult your textbook to answer this question.

5. What feature of muscle physiology does the slope allow us to compare between the dominant and non-dominant responses?

6. How does the slope of the response differ between the dominant and non-dominant arms? How might this be explained? Is there any noticeable pattern in the class results?

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EMG Activity in Response to Fatigue For the last set of data analysis, use the recordings from segment 2 and 4 that reflects fatigue when the muscle is fully recruited (segment 2) and only partially recruited, ie. the force of the squeeze is at half maximum (segment 4):

1. Use Delta T displayed in the third measurement results box to select the first 2 seconds of the recording after the clench has reached a plateau. Record the mean Force and mean EMG Integral in Table 2. Then move forward and select the last two seconds before the decline of the half-force clench. Go to segment 4 of the recording, and repeat the measurements on the non-dominant arm data. See Figures 8 and 9 for examples of how these values should be measured.

Note: If your recording has a peak at the very beginning, avoid that region when measuring the first two seconds of data. Instead, use the section that immediately follows the peak. Likewise, the end of the recording will show a sharp drop in EMG and force that corresponds to the time you released the dynamometer. Select a section immediately before the drop to analyze the mean of two seconds of the EMG Integral and Force.

Figure 8. Measuring force and EMG integral in the first 2 seconds of the fatigue experiment.

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Figure 9. Measuring the last 2 seconds of the fatigue experiment.

Table 2: EMG Activity During Continuous Sub-Maximal Force

Full Recruitment

(Dominant Arm)

Partial Recruitment

(Non-Dominant Arm)

First 2 seconds Last 2 seconds First 2 seconds Last 2 seconds

Force (Kg)


Calculations

Unit Activity

(mV-sec/Kg)

Percent Change

2. Calculate the Unit Activity, or the amount of electrical/neurological activity needed to generate 1 Kg of force by dividing the EMG Integral by Force:

EMG Int = Unit Activity in mV-sec

Force Kg

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3. Calculate the change in electrical activity as the muscle fatigues by determining the percent change in EMG activity/Kg force. Use a "+" sign to indicate an increase in activity and a "–" sign to indicate a decrease in activity.

Unit Activity for last 2 seconds – Unit Activity for first 2 seconds X 100 = % change

Unit Activity for first 2 second

QUESTIONS:

7. What was the purpose of holding the clench at half your maximum strength with the non-dominant arm, rather than at full strength like the dominant arm?

8. How did the EMG Integral change as your muscles fatigued? How did this change differ when the muscles were already fully recruited compared to partially recruited? How do you explain the difference in the EMG patterns?

9. What are the physiological differences between neural fatigue and muscle fatigue?

10. The cause of muscle fatigue is no longer as clear as it was once believed. What are some of the possible causes of muscle fatigue?

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Exercise 6: Contractility of Skeletal Muscle Text Reading: Silverthorn, 4th ed. pg. 403 – 409, 413 – 417; 5th ed. 412 – 419, 425 – 427

In this exercise, we will investigate the physiology of contraction in the gastrocnemius muscle of a frog. The Biopac system will produce the electrical stimulus to bring about muscle contraction and measure the force the muscle is able to generate under different conditions.

There are several similarities between the excitability of neural tissue and the excitability of muscle tissue. Like a single neuron, a single muscle fiber will not respond to any stimulus impulse that is below threshold. Any single stimulus impulse above threshold will produce a contraction, or single twitch (see Fig. 1), of the same strength. There are no gradations in the strength of the muscle response for a single twitch. Muscles outside the laboratory, however, do not use single twitches to produce motion.

Muscles rely on two physiological principles to bring about productive motion: recruitment and summation. Recruitment is increasing the number of motor units responding to a single stimulus, which results in the increase in tension in whole muscle. Summation is the increase in tension that results when a muscle fiber is unable to relax between twitches. Individual twitches overlap and produce a continuous, smooth contraction of increasing strength.

Recruitment requires increasing stimulus amplitude. Below threshold stimulus, there is no response from the muscle. As stimulus amplitude increases, only the muscle fibers with the lowest threshold initially respond and contract. As the stimulus amplitude continues to increase, an increasing number of motor units will be recruited so that the strength of the contraction, or tension, increases as well. Once all motor units in a muscle have been recruited, an increase in the stimulus amplitude will not result in any increase in tension. The contraction of a whole muscle can produce strong, weak or intermediate contractions depending upon how many motor units are stimulated.

Figure 1. A Single Twitch: An action potential on the muscle fiber is followed by contraction

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Summation requires increasing stimulus frequency. At low frequency, the muscle fiber will relax before the next stimulus impulse occurs. As the stimulus frequency increases and the time between the stimuli decreases, the muscle fiber cannot fully relax before the next stimulus occurs (see Fig. 2). This loss of relaxation between stimuli is called tetanus. We say a muscle is in incomplete tetanus if the muscle fiber is able to partially relax between stimulus impulses (see Fig. 3). A muscle fiber is

in complete tetanus if there is no relaxation at all between stimulus impulses.

We can explain the phenomena of summation and tetanus by examining the molecular events that bring about muscle contraction. Remember that tension is generated in a muscle fiber when Ca+2 is released from the sarcoplasmic reticulum (SR), triggering formation of actin-myosin cross-bridges. In order for relaxation to occur, Ca+2 must be pumped back into the SR. Summation occurs because each subsequent stimulus releases additional Ca+2 from the SR, which in turn increases the number of actin-myosin cross-bridges that form. As cytoplasmic Ca+2 increases, tension will increase until all possible actin-myosin cross-bridges have formed. At this point, the muscle fiber will reach maximal tension. This is illustrated in Figure 3. The black arrows along the X-axis indicate the occurrence of each stimulus, or action potential. With each stimulus, tension increases as Ca+2 accumulates in the muscle fiber until maximal tension is achieved.

Figure 3. Summation leading to incomplete tetanus, or unfused tetanus

Figure 2. A single twitch (left) and two summated twitches


1. Analyze the phases of a single muscle twitch.

2. Investigate the effect of stimulus strength on whole muscle response.

3. Investigate the effect of stimulus frequency on whole muscle response.

4. Correlate the molecular events within a muscle cell with your observations.

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Materials and Equipment Setup

This lab will be conducted as a demonstration by the instructor, but you should have an understanding of the experimental design. The frog gastrocnemius muscle will be dissected from the frog in such a way as to leave it firmly attached to the femur bone, which will be secured by a clamp to the ring stand. The Achilles tendon will be attached to a force transducer by means of a hook and strong thread. An electric shock will be sent from the Biopac stimulator through the stimulating electrodes to the muscle. If the voltage of the stimulus is high enough (above threshold), it will cause action potentials and subsequent contraction in some or all of the muscle fibers in the muscle.

When the frog muscle contracts, the force exerted will be converted to an electrical signal by the force transducer and Biopac amplifier will then convert this signal into units of mass in grams.

The Single Twitch

A muscle twitch is usually divided into three phases: 1) the latent period; 2) the contraction period; 3) the relaxation period (see Figure 4). The latent period is the time from when the stimulus is delivered to the first indications of contraction in the muscle. The contraction period, or contraction time, is the time it takes the muscle to reach its peak contraction after the latent period. The relaxation period is the time the muscle takes to return to resting tension after reaching its peak contraction.

1. Clicking Start will trigger the stimulator to send a jolt of electricity to the muscle. When this is demonstrated, watch the muscle to observe the actual contraction. When the stimulus is repeated, watch the recorder to observe the response on the computer screen. The top trace in green is the recording of the tension generated by the muscle and is measured in grams. The bottom trace in blue is the recording of the stimulus that is delivered to the muscle and is measured in mV.

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Figure 4. A recording of a single muscle twitch. Force is recorded in the upper channel (Channel 2) and the stimulus is recorded in the lower channel (Channel 3). The highlighted area measures the contraction time along the X axis as Delta T. The force of the contraction is measured along the Y-axis as P-P.

2. We will try to get you a printed copy of the twitch. In your recording of the single twitch, locate the latent period by measuring the time period between the stimulus on the blue line below and the beginning of twitch on the red line above.

3. Note the length of contraction time, relaxation time, and the total time as well. Record these measurements in Table 1 on the next page.

4. The amplitude of the muscle twitch is proportional to the force, or tension, generated by the twitch. This measurement is in grams.

Table 1. Data for Muscle Twitch Latent Period

Contraction Time Relaxation Time

Time of Total Twitch Contraction Tension (g)

Questions:

1. How does direct electrical stimulation produce contractions of the muscle?

2. Describe the molecular events in the muscle fiber that produce a twitch in response to the stimulus voltage. Specifically, what molecular events occur during the latent period? Which events occur in the contraction period? Which

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events occur in the relaxation period? Be sure to include the role of calcium in your description.

Stimulus Strength and Recruitment

In this experiment, we will measure the strength of the contraction produced in a muscle that is stimulated with impulses of progressively higher voltage. Remember that motor units within a whole muscle have variable thresholds. The slow twitch motor units have the lowest threshold and will be the first to respond as the voltage of the stimulus is increased. Intermediate motor units are recruited next. The fast twitch motor units have the highest thresholds and are the last to be recruited. As more motor units are recruited with increased stimulus voltage, the tension produced by each twitch increases.

1. We will start the demonstration by reducing the stimulus voltage to 0.0V. We will apply a series of stimuli to the muscle, increasing the stimulus voltage each time. Watch the frog muscle for a response when it is stimulated.

2. We increase the stimulus voltage in 0.1V increments. The voltage at which the muscle first responds to the stimulus is threshold. We'll continue to increase the stimulus voltage by 0.1 V increments until the muscle response no longer increases in magnitude. Record the data collected in Table 2.

Baseline tension is the amount of tension measured by the force transducer when the muscle is unstimulated. This is due to the tautness of the string and the attachment of the muscle to the apparatus. This baseline tension must be subtracted from the measured tension to obtain the tension produced by the contraction itself. The tension produced by the contraction itself will be called the "adjusted response".

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Table 2. Data for Stimulus and Response Stimulus Voltage

(Volts) Response Tension

(grams)

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Figure 5. The increase in the force of contraction (upper channel) as the stimulus voltage increases (lower channel) indicates recruitment of additional muscle fibers within the whole muscle. All motor units have been recruited when increasing voltage no longer increases the strength of contraction.

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Questions:

3. Why doesn’t the muscle respond to low stimulus voltages?

4. Graph the relationship between stimulus voltage and the muscle response. Which variable is the independent variable (X-axis) and which is the dependent variable (Y-axis)? What is the physiological explanation for this relationship?

5. Is the single twitch response to the same voltage stimulus always the same? How do you explain this?

Stimulus Frequency and Summation

In this experiment, we will measure how whole muscle responds to increases in stimulus frequency (the number of stimuli per second). The stimulus voltage will remain constant throughout this experiment. As frequency increases, summation and tetanus will begin to occur. Summation can be measured as the increase in the muscle response along the Y-axis. Tetanus will be observed as the loss of relaxation between twitches.

1. Values in the Stimulator Panel:

• Stimulus voltage set to produce a maximal response (the voltage that produced full recruitment in the previous experiment)

• Frequency set to 2 Hz (stimuli per second).

• Total number of impulses set to 10

We will begin by applying ten impulses to the muscle at a frequency of two times per second. You should observe ten separate, individual twitches.

2. For each subsequent recording, we increase the frequency of stimulation in 1 Hz increments up to 20 Hz. We will need to increase the number of impulses to 20 or even 30 once the frequency is set above 10 Hz.

3. Measure the tension produced at each frequency by highlighting the response and measuring P-P. Record the data collected in Table 3.

4. Measure the relaxation time of the muscle after fused tetanus has occurred just as we did for a single twitch (delta T).

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Table 3. Data for Summation and Tetanus Stimulus

Frequency (Hz)

Response Tension (g) Presence of Tetanus? (none, incomplete, or

complete)

Summation Tension (Difference between a

single twitch and tetanus)

2 Hz None

3 Hz

4 Hz

5 Hz

6 Hz

7 Hz

8 Hz

9 Hz

10 Hz

15 Hz

20 Hz

Relaxation Time After A Single Twitch

Relaxation Time After Complete Tetanus

So What Happened?

Remember what causes contraction in a muscle fiber. An action potential causes calcium ions to be released from the sarcoplasmic reticulum into the sarcoplasm. This allows myosin to form cross bridges with actin filaments of the sarcomere and generate the movement associated with the contraction. The more cross bridges formed, the stronger the contraction.

Relaxation is brought about by the removal of the Ca++ from the sarcoplasm. If a second stimulus is delivered to a fiber before all of the Ca++ are removed (the fiber has not relaxed), more Ca++ are released and the fiber starts a second contraction without completely relaxing from the first.

If there is no relaxation at all between stimuli, but rather a smooth sustained contraction, it is called “Fused Tetanus”, “Complete Tetanus” or just plain “Tetanus.” If there is some relaxation between the stimuli, the contraction is referred to as “Unfused Tetanus" or Imcomplete Tetanus". If a muscle completely relaxes in between successive stimuli, there is no tetanus at all.

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As a muscle responds to successive stimuli, the contractions increase in strength. This property is called “Summation of Contractions,” or simply "summation." Summation makes sense when you remember that the increased frequency of stimuli causes increased amounts of Ca2+ to be deposited in the sarcomeres. The increased Ca2+ means there will be increased numbers of myosin-actin cross bridges, and thus, an increase in the strength of contraction.

Sample data can be seen in Figure 6. The data has been squished into one window so that the results for each increase in frequency can be compared side by side. Note the difference in the amplitude of the single twitches on the left versus the amplitude of the contractions showing fused tetanus on the right. The I-bar added to the figure indicates summation, or the increase in force that results from fused tetanus.

Figure 6. Increased summation is observed as a result of increased stimulus frequency. The first stimulus was at a low enough frequency to result in single twitches. As frequency increases, individual twitches begin to fuse (tetanus) and the strength of contraction increases (summation). Summation, or the increase in force produced by increasing the frequency of the stimulus impulses, is illustrated by the vertical black I-bar over the first set of twitches.

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Questions:

6. Can the tension produced by an isolated single twitch vary in a muscle cell? What circumstances would allow a twitch to increase in tension?

7. What do the terms unfused and fused tetanus refer to? What do the terms incomplete and complete tetanus refer to?

8. In what section of the recording can you detect summation, and in what section can you detect incomplete tetanus? Where do you see complete tetanus on the recording?

9. Explain the cellular mechanism that produces tetanus and summation.

10. Compare the relaxation time of the single twitch to the time measured after complete tetanus. Why is the relaxation time much longer after tetanus than after a single twitch?

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Exercise 7: Electrocardiography Text Reading: Silverthorn 4th ed. pg. 477 – 489; 5th ed. 487 – 500

Our heart is a four-chambered organ that relies upon electrical conductivity to accomplish its job of pumping blood throughout the body. The normal electrical conductivity of the heart follows a sequence traveling from the non-muscular right atrium to the powerful ventricles. This wave of depolarization, as it is often termed by physiologists, causes the atria to lightly contract in order to fill the ventricles. The wave then travels through the ventricles causing the muscle to depolarize and contract, ejecting blood to both our pulmonary and systemic circuits.

A normal cardiac cycle begins as an action potential that develops in the sino-atrial node (SA node), in the upper right wall of the right atrium. This action potential spreads through the atrium via internodal fibers and causes the muscle of the atrium to contract. The electrical impulse arrives at the atrioventricular node (AV node) and is delayed for approximately .20 seconds, allowing the atria to contract. The impulse then spreads to the ventricles via the bundle of His, and right and left bundle branches located in the ventricular septum, and finally to the Purkinje fibers in the base of the ventricles and upward. See figure 1 for a diagram of the conductive system pathway.

Figure 1. The heart

Action potentials spread very rapidly through the atria and then throughout the ventricles. The effect of the rapid spread is a very large mass of cells depolarizing and then quickly repolarizing. Each massive depolarization and repolarization (one heart beat) causes an electric field to rapidly spread throughout the body fluids. These changes in electrical fields are very weak, but can be detected as echoes on the skin with sensitive electrodes placed on other parts of our body. The recording of the electrical field is called an electrocardiogram or ECG (also

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EKG) and graphically depicts the electrical events of the cardiac cycle. Figure 2 depicts a normal human ECG.

Figure 2. Recording of a typical human ECG

The Purkinje fibers relay the electrical impulse directly to the ventricular muscles, causing them to contract (systole). The depolarization of the ventricular muscle from inferior to superior forces the ventricles to contract in a manner that pushes blood superiorly through the aorta and pulmonary arteries located in the roof of the ventricles. Repolarization of the heart spreads throughout the atria to the ventricles via the same pathway, allowing the ventricles to relax (diastole). We are able to infer the mechanical activity of the heart from the ECG. The Wigger's Diagram (see figure 3) correlates the electrical events (depolarization and repolarization) with the mechanical events (systole, diastole) of the cardiac cycle.

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Figure 3. Wigger’s diagram portraying correlation between electrical, mechanical, and

auditory physiology of the heart.

An ECG can be recorded by placing electrodes on the skin. The arrangement of the electrodes on the body, one positive and one negative, with respect to a third electrode, the ground, is called a lead. Each lead records the heart's electrical activity from a different perspective on the body. For this experiment, the lead you will be using has one positive electrode on the LEFT ANKLE, one negative electrode on the RIGHT WRIST, and the ground electrode on the RIGHT ANKLE (Figure 4).

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Figure 4. Electrode and lead placement

Factors that affect the cardiac cycle: Temporary minor increases and decreases in heart rate associated with the resting respiratory cycle reflect reflexes of our heart in response to the cycling of intrathoracic pressure. The slight changes in arterial and venous pressure are detected by baroreceptors that relay the signal to our brain, which then alters our heart rate accordingly.

When inspiratory muscles contract the intrathoracic pressure decreases. This decrease in pressure allows veins to expand, causing a brief drop in venous pressure, venous return, cardiac output, and systemic arterial pressure. The brief drop in systemic arterial pressure briefly reduces the carotid baroreceptor firing, causing a brief increase in heart rate.

When inspiratory muscles relax the intrathoracic pressure increases. This increase in pressure compresses the thoracic veins, causing a brief increase in venous pressure and return. This increased pressure and return causes a momentary increase in heart rate from the firing of the venous baroreceptors. This slight increase is short lived as it increases cardiac output and arterial blood pressure. The increase in arterial pressure increases carotid baroreceptor firing causing heart rate to decrease (Figure 5).

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Figure 5. Effects of the resting respiratory cycle on heart rate

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Today's Objectives

1. Become familiar with the electrocardiograph as a tool for evaluating electrical events in the heart.

2. Correlate electrical events as displayed on the ECG with the mechanical events that occur during the cardiac cycle. Figure 3 will aid in this objective.

3. Observe the effects of intrathoracic pressure changes and the baroreceptor reflex on heart rate.

4. Measure the effects of increased heart rate on the systolic and diastolic segments of the cardiac cycle.

Getting Started 1. The equipment for today’s experiment is listed below.

√ Computer

√ Biopac MP35 Acquisition unit

√ BIOPAC electrode lead set plugged into Ch. 2

√ Disposable paste-on electrodes

√ Exercise mat

2. Use the checklist below to ensure that the basic set-up is working:

• Check to make sure the electrode lead set is plugged into Channel 2.

• Turn the computer on. For Mac laptops, press the on silver "ON" button located above the computer keyboard. For PCs, push the large button on the front of the tower. The computer must finish booting up before turning on the Biopac unit (blue box).

• Check that the Biopac unit is plugged into the electrical outlet.

• Check that the Biopac unit is connected to the computer via a black USB cable.

• Make sure the Biopac unit is turned ON. The Power switch is on the back panel on the right. Check that the AC adapter for the unit is plugged into the back. Wait until the "Busy" light stops flashing before launching software.

3. The subject should remove all jewelry from wrists and ankles.

4. Clean and scrub the regions on the right forearm, and medial right and left ankle for electrode attachment using an alcohol pad (see Figure 4 for electrode placement). Let the areas dry after cleaning with the alcohol pad. Place the electrode on the anterior surface of the RIGHT forearm superior to the wrist. Place the second and third electrode on the MEDIAL surface of the lower leg, just superior to the medial malleolus.

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5. Each of the pinch connectors needs to be attached to a specific electrode. Each lead cable is a different color. Attach the WHITE lead (negative) to the electrode on the RIGHT FOREARM or WRIST. Attach the RED lead (positive) to the electrode on the LEFT LEG or ANKLE. Attach the BLACK lead (ground) to the electrode on the RIGHT LEG or ANKLE (see Figure 4). Position the lead cables so that they are not pulling on the electrodes.

CALIBRATION

1. 5. Launch the Biopac Student Lab software application. For Mac laptops, click on the Biopac Student Lab application icon in the dock (see Fig.3). The dock is the row of application icons found at the bottom of the screen (usually). For those of you unfamiliar with the MacIntosh operating system, ask the instructor or a Mac-proficient lab partner for help.

For PC users, click on the green "Start" button in the bottom left corner, go to "Programs", then "Science", then choose BSL Lessons 3.7.

2. Choose lesson L05-ECG-1 from the menu. Type in your file name.

3. Have the subject lie down and make sure he/she is relaxed. Do not laugh, cough, sneeze, or have any other shifts in body position during calibration.

4. Make sure the electrodes are securely adhered to the subject’s skin.

5. Click CALIBRATE.

6. Wait for the calibration procedure to stop. At the end of 8 seconds, there should be a recognizable ECG waveform with no large baseline drifts. If your data resembles Figure 6, proceed to the data recording section. If your data shows any large drifts or does not resemble figure 6 you must redo the calibration. Click on REDO CALIBRATION, then follow the above directions again.

Figure 6. ECG Calibration

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DATA RECORDING

Data will be collected while the subject is lying down, immediately after sitting up, taking deep breaths while seated, and after short vigorous exercise. During all of the recordings do not talk, laugh, or have any other activity that may alter the ECG recording.

LYING DOWN 1. Have the subject lay down on a yoga mat or other pad on the floor.

2. Click RECORD.

3. Record for 20 seconds.

4. Click SUSPEND.

5. Review the data to ensure that it resembles Figure 7 if it does proceed to the next section. If it does not, click REDO and rerecord the data.

Figure 7. ECG lying down

SITTING UP In this experiment the subject is going to sit up quickly and then relax. The goal is to capture the ECG data as close to possible after sitting up. Again, ensure that there is no behavior that may alter the data.

1. Have the subject sit up quickly. Immediately click RESUME. An append marker labeled “After sitting up” will automatically be inserted.

3. Record for 20 seconds

4. Click SUSPEND.

5. Review the data to ensure that it resembles Figure 8. If it does, proceed to the next section. If it does not, click REDO and rerecord the data.

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Figure 8. ECG after sitting up

SEATED, DEEP BREATHING In this section we are going to examine the effects of changing intrathoracic pressures on HR. The subject will be seated and breathing deeply. You will place an event marker at the beginning of each INHALE and EXHALE.

1. Once you begin recording, the subject is going to begin a series of slow prolonged breaths for five cycles. Each time the subject inhales AND exhales place an event marker by hitting the ESCAPE key for Macs or F9 for PCs. Make sure to write down if you started with an inhale or exhale. The event marker marks the recording with an inverted triangle right above the ECG trace.

Click Resume to begin recording. An append marker (blue diamond) labeled “Deep Breathing” will automatically be inserted. Have the subject take 5 deep slow breaths.

2. Click SUSPEND.

3. Review the data to ensure that it resembles Figure 9. If it does, proceed to the next section. If it does not, click REDO and rerecord the data.

Figure 9. ECG deep breathing

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THE EFFECT OF EXERCISE ON THE CARDIAC CYCLE In this section, we want to examine the effects of short intense exercise on the ECG. In order to see a measurable difference, the exercise must be VERY VIGOROUS! The cables will need to be removed from the subject during exercise.

Remember to reattach the leads correctly so that the WHITE lead attaches to the electrode on the WRIST. The RED lead attaches to the electrode on the LEFT LEG. The BLACK lead attaches to the electrode on the RIGHT LEG. After the subject exercises vigorously, have the subject sit and immediately attach the lead cables as fast as possible. This is very important as the goal is to capture the ECG reading as soon as possible after exercise. Once the lead cables are reattached, resume recording,

1. Remove the lead cables from the electrodes.

2. Have the subject perform vigorous exercise to the point that the heart rate is elevated for at least a minute. We recommend running around the building or up and down the stairs several times.

3. Reattach the lead cables as quickly as possible. WHITE to the WRIST. RED to the LEFT LEG. BLACK to the RIGHT LEG.

4. Click RESUME. An append marker labeled “After Exercise” will automatically be inserted.

5. Record for 60 seconds.

6. Click SUSPEND.

7. Review the data to ensure that it resembles Fig. 10. If it does, proceed to the next section. If it does not, click REDO and rerecord the data.

8. When you are satisfied with your data, click DONE, then choose "Yes" and "Analyze Current Data File". Remove the lead cables and the electrodes from the subject and prepare for data analysis.

Figure 10. ECG after exercise

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Analysis of the Data

1. Refer to "Exercise 6: Physiological Instrumentation” that introduced you to the Biopac equipment to refresh your memory about how to use the analysis tools.

2. Adjust the display so that you are viewing four successive beats from your segment 1 recording.

3. Set the first two measurement boxes at the top of the screen so the following are displayed:

Channel Measurement Function

CH2 Delta T (∆T)

CH2 BPM (Beats Per Minute)

• Delta T (∆T) is the time measured along the X-axis by the selected region

• BPM is heart rate calculated based on the Delta T selected and is valid only if the selected region is one complete cardiac cycle

Figure 11: ECG segments for measurements

4. Measure Delta T for a single cardiac cycle: use the I-beam cursor to select the area between two successive R waves (R–R segment; see Fig. 11). Record the ∆T and the BPM from this reading for Cardiac Cycle 1 in Table 1.

5. Measure the Delta T for ventricular systole (Q-T) and diastole (T-Q) of the same cardiac cycle and record the data in Table 2. To measure the Q-T segment, position the I-beam at Q

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and drag to the end of T (See figure 11). To measure the T-Q segment, position the I-beam at the end of T and drag to the next Q.

6. Take measurements for two other cardiac cycles (Cycles 2 and 3) in the "lying down" display and record the data in Tables 1 and 2. Calculate the mean of the these values for all three cardiac cycles by adding them together and dividing by 3.

7. Make similar measurements for the "sitting" data and record in Table 3.

8. To see the effect of deep breathing on heart rate, measure the first heart beat immediately following a deep inspiration or a deep expiration. Record this data in Table 4. Do this for three separate breathing cycles and calculate the mean.

A: SUPINE, RESTING, REGULAR BREATHING DATA Table 1

Measurement Cardiac Cycle

1 2 3

Mean

∆T (seconds)

(R-R Interval)

BPM (Beats Per Minute)

Table 2

Ventricular Readings

Cycle 1 Cycle 2 Cycle 3 Mean

Q – T Interval (seconds)

(corresponds to Ventricular Systole)

T – Q Interval (seconds)

(corresponds to Ventricular Diastole)

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B: SITTING DATA Table 3

Measurement Cardiac Cycle

1 2 3

Mean

∆T (seconds)

(R-R Interval)


C: SEATED DEEP BREATHING DATA

Table 4

Rhythm Cycle 1 Cycle 2 Cycle 3 Mean

INSPIRATION

∆T (seconds)

(R-R Interval)


EXPIRATION

∆T (seconds)

(R-R Interval)


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D: AFTER EXERCISE

Table 5

Ventricular Readings

Cycle 1 Cycle 2 Cycle 3 Mean

Time of Q – T Interval (sec)

(corresponds to Ventricular Systole)

T – Q Interval (seconds)

(corresponds to Ventricular Diastole)

BPM for R – R

(one complete cardiac cycle)

E: COMPILATION OF HEART RATE (BPM) DATA Table 6

Condition BPM Mean

Supine, Regular Breathing (from Table 1)

Seated Regular Breathing (from Table 3)

Seated, Deep Breathing, Inhalation (from Table 4)

Seated, Deep Breathing, Exhalation (from Table 4)

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F: COMPARISON OF RESTING AND POST-EXERCISE CARDIAC CYCLES

Table 7

Q – T (sec)

Systole

T – Q (sec)

Diastole

1 At Rest (from Table 2)

2 After Exercise (from Table 5)

3 Difference between Rest and Exercise

(subtract line 2 from line 1)

4 Percent Difference

(divide line 3 by line 1)

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QUESTIONS:

1. What electrical and mechanical events correlate to the P wave? The QRS complex? The T wave?

2. During which ECG interval does filling of the ventricles occur? During which ECG interval does ejection occur? What is systole and diastole?

3. Describe the path an action potential travels during one cardiac cycle.

4. Describe the flow of blood through the heart and the subsequent flow through the body.

5. What changes occurred in heart rate as position changes from lying down to sitting up? Describe the physiological mechanisms causing these changes.

6. What changes occurred in the cardiac cycle due to the respiratory cycle? Describe the physiological mechanisms causing these changes.

7. What changes occurred in the duration of systole and diastole between resting and post-exercise? Explain.

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Exercise 8: Respiratory Physiology Readings: Silverthorn 4th ed. - pg. 567 - 568, 580 – 582; Silverthorn 5th ed, 578 – 579, 591 – 592.

Cellular respiration is the process in which the body consumes oxygen and produces carbon dioxide. External respiration is the exchange of oxygen and carbon dioxide between the lung tissues and the environment, and relies on a series of tubes called bronchioles that branch and terminate as clusters of small, membranous air sacs called alveoli.

Ventilation of the lungs occurs when chest muscles contract and the thorax expands. Inspiration is achieved by a contraction of the diaphragm and intercostal muscles. Expiration is usually passive in the resting individual as muscle relaxation and gravity act to decrease thoracic volume. Forced expiration, as during exercise, is achieved by contraction of intercostal muscles.

The amount of air that moves in or out of the lungs during breathing is called the lung volume. Lung volume is dynamic, changing according to the requirements of the body. The depth and rate of breathing are controlled by the respiratory control center in the brain, which insures that the exchange of oxygen and carbon dioxide takes place at a level that matches the body’s needs. In addition to variation due to activity, lung volumes are also influenced by a person’s height, physique, age, environmental conditions such as altitude and smoking, and state of health. Therefore, the measurement of lung volume is an important clinical assessment.

Pulmonary Function Tests (PFT) measure lung volumes with an instrument called a spirometer. In this lab you will use a flow-type spirometer to measure standard lung volumes. We measure airflow (ml/sec) by breathing into a sensitive pressure transducer. The mathematics of converting a continuously changing flow to volume is possible because we send information through the transducer to the computer and use software to compute the volume.


1. Measure Tidal Volume (TV).

2. Calculate Respiration Rate and Minute Volume.

3. Measure Vital Capacity (VC) and compare to normative values.

4. Measure Forced Expiratory Volumes FEV1 and FVC. Calculate the ratio of FEV1 to Forced Vital Capacity (FEV1%). Use this value to evaluate respiratory fitness.

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Lung Volume Definitions

1. Tidal Volume (TV) is the volume of air that is inhaled or exhaled with each breathing cycle. It varies with conditions (e.g. rest, exercise, and posture). A more accurate measure is achieved if several breaths are averaged.

2. Vital Capacity (VC) is the volume change that occurs between maximal inspiration and maximal expiration.

3. Forced Vital Capacity (FVC) is the volume of gas that can be expelled during a forced breath from full inspiration to complete expiration. In normal subjects this volume differs little from vital capacity in which expiration is gentle and not forced.

4. Forced Expiratory Volume (FEV) is the volume of gas which can be expired in a short time, during a forced expiration starting from full inspiration. The time recorded is usually for one second, which is designated FEV1.

5. Forced Expiratory Volume Ratio (FEV1%) is the forced expiratory volume expressed a s a percentage of the forced vital capacity, i.e. (FEV1/ FVC) X 100%.

Figure 1. Respiratory Volumes

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Equipment

1. Airflow transducer (SS11A). The transducer for today’s lab is an airflow transducer. You will breathe into this apparatus and the software will convert airflow to volume. In order for this measurement to be accurate you must follow the procedure exactly as described. The airflow transducer should be plugged into Ch. 1.

2. Bacteriological Filter. This is a sanitary device designed to prevent contamination from user to user. This piece is NOT tbe shared.

o

3. Disposable Mouthpiece. Obtain one of these from your instructor. This is the only part that your mouth will come in contact with during the exercise. This piece is NOT to be shared.

4. Nose Clip. The nose clip will prevent you breathing through your nose insuring that the airflow comes only from your mouth.

5. Calibration Syringe. The syringe is for calibrating the Biopac unit, by delivering one full liter of air during calibration procedure. This is will “teach” the software how much acorresponds to 1 liters.

irflow

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STARTING UP

1. Turn your computer ON. When the computer has finished booting up, turn the MP35 unit ON. The "Busy" light on the Biopac unit will flash. Once the "Busy" light goes out, launch the Biopac Student Lab software.

2. Choose Lesson 13 (L13-LUNG-2).

3. Type in your file name. Click OK.

4. Check to see if the airflow transducer (SS11LA) is plugged into Channel 1.

5. Place the calibration filter onto the end of the calibration syringe. NOTE: do not use your personal filter. Use the filter labeled "Calibration". A filter must be included in the calibration procedure because of the way if affects airflow. See Fig. 2.

Figure 2. Calibration syringe with filter attached.

6. Please read ALL of Step 6 before attaching the syringe to the airflow transducer. Insert the calibration syringe with its filter into the Airflow Transducer on the side labeled “Inlet”. CAUTION: The calibration syringes are extremely fragile. NEVER HOLD ONTO THE AIRFLOW TRANSDUCER HANDLE when using the calibration syringe or the tip will break off ( See Fig 3a and 3b for proper handling). BOTH hands must be placed on the syringe itself. One hand should be on the plunger, the other on the body of the syringe (see Fig 3b).

Figure 3a. Incorrect way to hold onto the syringe/transducer assembly.

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Figure 3b. Correct way to hold on to the calibration syringe.

Calibration

You are not holding on to the airflow transducer handle now are you? Good. You should have both hands on the calibration syringe. Practice using the plunger of the calibration syringe until you feel you can get a smooth but firm motion of air moving in and out of the barrel. Continue once you feel confident with the handling of the syringe.

1. Pull the Calibration Syringe Plunger all the way out and hold the Calibration Syringe/Filter Assembly parallel to the ground.

2. Read the Journal window on the lower screen for directions:

The first part of the calibration sets the baseline. It calibrates with no air movement within the barrel. Hold the syringe still without touching the plunger.

Click Calibrate. A prompt will remind you to do nothing during this first part of calibration (Figure 4).

Figure 4. Warning to not touch plunger at this time.

Note the green line traveling across the screen. After two screens, it will stop on its own. Wait for it to end.

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3. The second part of the calibration measures 1 Liter of airflow through the barrel. The method of delivery needs to be very precise and is described in detail in the Journal window on the lower screen of your computer. Read those directions before clicking Yes.

The following directions summarize the directions in the Journal window:

You will push and pull the plunger a total of ten times, 5 in and 5 out, forcefully enough to cause the plunger to produce a whistling sound. Practice this a few times before actually calibrating.

Follow this rhythm as you practice: plunger in, wait 2 seconds; plunger out, wait 2 seconds. Continue this pattern until you have finished the 10 strokes.

Note: The repeated strokes are required because of the complexity of the Airflow to Volume calculation. The accuracy of this conversion is increased when analyzing the airflow variations occurring over five complete cycles of the calibration syringe.

4. When you are ready to begin, click Yes. The second stage of the calibration procedure will begin to record. Wait 2 seconds, then begin by pushing the plunger in. Cycle the syringe plunger in and out completely 5 times (10 strokes) following the rhythm practiced above. Click End Calibration. Your calibration recording should look like Figure 5.

Figure 5. Results from a correct calibration

5. If your data looks like that of Figure 5, you can proceed to the next step. If it does NOT look like the image above, click Redo Calibration. It may take two or three tries to get the calibration right. Check with your instructor to determine what you may be doing wrong and repeat the calibration.

6. When you are satisfied with your recording gently detach the transducer from the calibration syringe and filter. Be gentle, as the torque on the syringe tip can cause it to break off.

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RECORDING TIDAL VOLUME AND VITAL CAPACITY

1. Attach the subject's own filter onto the airflow transducer on the inlet side and then attach the mouthpiece onto the filter (Fig. 6).

Figure 6. Airflow transducer with filter and mouthpiece attached.

2. Seat the subject facing away from the computer monitor. The subject should wear the nose clip, and hold the airflow transducer by the handle (now you are allowed), with its attached filter and mouthpiece parallel to the floor. Keep the airflow transducer upright at all times (Figure 6).

Practice Run

3. Practice the first breathing routine without recording on the computer:

Have the subject breathe in and out through the mouthpiece as normally and relaxed as possible (easier said than done).

It is OK to bite on the mouthpiece as they breathe. A breath is made up of one inhale and one exhale cycle. The subject should try to inhale normally, followed by an unrushed exhaling. There is a tendency to not exhale completely before beginning a new breath because of the awkwardness of the situation. Try to breath as normally as possible without too much conscious thought.

4. Monitor the subject's breathing pattern by the rise and fall of their torso. After five cycles, have the subject inhale as deeply as possible followed by exhaling as completely as possible.

Note: Inhaling as deeply as you can and exhaling as deeply as you can takes EFFORT. The subject should truly force their inhalation and then their exhalation until they are bending over trying to eliminate the last volume of air.

5. Let the subject breathe normally once again for five cycles. Remember a cycle is one inhalation followed by exhalation.

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These steps differ from the prompt in the lower journal window on your laptop. Do NOT follow the computer screen prompt. Follow the directions on this handout instead. From here on, this handout will differ from the prompts on the screen. Ignore the prompts.

Ready to Record (The real thing)

6. When ready and clear on the directions above, a member of the group other than the subject will click Record FEV.

7. Read the following steps aloud to your subject as you record.

a) Breathe normally until we tell you otherwise. (Helper counts 5 cycles)

b) Inhale as deeply as you can, and exhale as deeply as you can.

c) Breathe normally until we tell you to stop. (Helper counts 5 cycles)

8. Click Stop. The recording should look like the Figure 7.

Figure 7. Tidal Volume plus Forced Expiratory Volume tracing.

9. You may redo the recording if you feel there was an error in your methodology and it does not look like the recording above. It is best to fix errors now, early in your data collection than later. Don’t hesitate to re-do. Obtaining a good recording usually takes a couple of attempts.

10. You will collect data from this screen. DO NOT click "Setup FEV".

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Measuring Tidal Volume

1. Set up two measurement boxes on the upper left side of the screen (see Figure 8.). The first should be P-P, the second should be Delta T. P-P will measure volume in liters (Y-axis) and Delta T will measure time in seconds (X-axis).

Figure 8. Measurement boxes set up for P-P and Delta T

Check glossary at the end of this week's exercise for an explanation of the P-P or Delta T calculation.

2. Measure the volume (Y-axis) of one breath (TV) by using the I-beam cursor to highlight the inhalation phase of one breathing cycle. Position the I-beam cursor from the valley to the peak of one breath (see Figure 9). Use three significant digits. Record P-P in liters in Table 1.

Figure 9. I-beam location for measuring Tidal Volume (P-P)

3. Measure the time of the same breath (cycle time) by positioning the I-beam from valley to valley. Record Delta T in seconds.

Repeat these measurements for two more breathing cycles from your recording. (Pick those without any major anomalies).


4. Calculate the average TV and cycle time for all three breaths. Tidal volume values may range from 0.2 L to 0.6 L, depending on the size and respiratory health of the individual. Cycle time may range from 3 to 6 seconds per breath.

Note: Averages are preferable because they account for variability between breathes.

Table 1: Tidal Volume (TV)

Cycle Tidal Volume

(P-P in L)

Cycle Time

(∆ T in sec)

#1

#2

#3

Per Cycle Average

Respiratory Rate (cycles/minute)

Minute Volume (liters/minute)

Calculating Respiratory Rate and Minute Volume

Respiratory rate by itself is of limited clinical value, but may be useful in conjunction with other signs of respiratory distress. The simple act of measuring respiratory rate is often sufficient to alter it. Respiratory rate can vary widely and is often difficult to measure accurately because it is relatively slow. In a clinical setting, the most accurate measurements of respiratory rate should be taken over a full minute and without the subject's knowledge.

Minute Volume takes into account both rate and depth of breathing. It is particularly important in sick or injured patients using ventilators to support their breathing. The volume of air delivered by the ventilator must be closely monitored.

1. Calculate your subject’s Respiratory Rate: Respiratory Rate (breaths/min) is the number of breaths in a minute. Use the average cycle time from Table 1 to calculate respiratory rate using the formula below.

Respiratory Rate = Number of breaths X 60 sec. = (breaths/min)

Cycle Time (sec) 1 min.


Average Respiratory Rate is between 12 and 17 breaths per minute. Record your result in Table 1.

2. Calculate Minute Volume:

Minute Volume (liters/min) is the volume of air inspired in a minute. This is calculated by multiplying Tidal Volume by Respiratory Rate. Use the formula below for your calculation.

Minute Volume = TV X Resp. Rate = L X breaths = (L/minute)

breath minute

3. Record your results on the board or overhead projector with the rest of the class data. We do this to get an idea of variability seen in a random group of individuals.

Measuring Vital Capacity

You will now measure the volume of exchanged air occurring between maximal inspiration and maximal expiration, called vital capacity (VC). Normal values for VC are affected by age, gender, and height. Use the tables of normative values provided to assess your VC. See “Interpretation of Pulmonary Function Tests” below for more information.

1. Use the I-beam cursor to select the area of maximum exhale. You do this by placing the

cursor at the start of exhale (just before the drop) and dragging the I-beam over to the lowest point of the following valley. This highlighted section should be about 3 seconds in length. See figure 10. Record the P-P value in Table 2. Record your results on the board or overhead projector with the rest of the class data.

2. Do not exit your screen yet. We will use the same file to collect one more set of data to measure FEV1 and FVC.


Figure 10. I-beam location for measuring Vital Capacity.

Table 2: Vital Capacity

Vital Capacity (P-P) (Liters)

Room Temperature BTPS Factor (from Table 3)

BTPS Adjusted Volume

The BTPS factor is a conversion factor used to correct for variations in airflow caused by ambient temperature differences. Multiply your vital capacity value by the factor in Table 3 corresponding to your room temperature. Enter the result in Table 2.

Example: Vital Capacity: 3.71 Liters

Room temperature: 21 degrees C. requires a 1.096 factor

BTPS= 3.71 X 1.096= 4.066 Liters


Table 3. Conversion Factors from ATPS to BTPS

Room Temperature (ºC) BTPS Factor*

18º 1.113

19º 1.108

20º 1.102

21º 1.096

22º 1.091

23º 1.085

24º 1.080

25º 1.075

26º 1.069

27º 1.063

28º 1.057

29º 1.051

* Assumes a of 760 mmHg barometric pressure


Table 4. Normative Values for Vital Capacity (in liters)

FEMALES

Age (years)

Height (in.) 16-25 26-35 36-45 46-55 56+

56” < 58” 2.90–2.70 2.70–2.55 2.55–2.35 2.35–2.20 2.20–2.00

58” < 60” 3.15–3.00 3.00–2.80 2.80–2.60 2.60–2.45 2.45–2.25

60” < 62” 3.45–3.25 3.25–3.05 3.05–2.90 2.90–2.70 2.70–2.50

62” < 64” 3.70–3.50 3.50–3.30 3.30–3.15 3.15–2.95 2.95–2.80

64” < 66” 4.00–3.80 3.80–3.60 3.60–3.40 3.40–3.25 3.25–3.05

66” < 68” 4.20–4.05 4.05–3.85 3.85–3.65 3.65–3.50 3.50–3.30

68” < 70” 4.45–4.30 4.30–4.10 4.10–3.90 3.90–3.75 3.75–3.55

MALES

Age (years)

Height (in.) 16-25 26-35 36-45 46-55 56+

60” < 62” 4.10–3.80 3.80–3.50 3.50–3.20 3.20–2.90 2.90–2.60

62” < 64” 4.45–4.15 4.15–3.85 3.85–3.55 3.55–3.25 3.25–2.95

64” < 66” 4.75–4.45 4.45–4.15 4.15–3.85 3.85–3.55 3.55–3.25

66” < 68” 5.10–4.80 4.80–4.50 4.50–4.20 4.20–3.90 3.90–3.60

68” < 70” 5.40–5.10 5.10–4.80 4.80–4.50 4.50–4.20 4.20–3.90

70” < 72” 5.75–5.45 5.45–5.15 5.15–4.85 4.85–4.55 4.55–4.25

72” < 74” 6.05–5.75 5.75–5.45 5.45–5.15 5.15–4.85 4.85–4.55


Forced Expiratory Volume in One Second (FEV1)

Forced Expiratory Volume is a reflection of the mechanical aspects of the functioning of the respiratory system and is measured as FEV1. FEV1 is the volume of gas expired in the first second of forced expiration. The ratio of FEV1 to Forced Vital Capacity is a measure of the speed with which air can be expelled from the lungs and is also referred to as FEV1%. The FEV1/FVC ratio is the most important factor for distinguishing obstructive disease from restrictive disease. In general, healthy individuals are able to expel 75% to 80% of FVC in the first second of expiration.

Reduction in this ratio, coupled with relatively normal lung volumes, indicates obstructive pulmonary disease. Relatively normal FEV1/FVC ratios coupled with reduced lung volumes indicates restrictive disorders. Another factor in distinguishing obstructive from restrictive disorders is the use of bronchodilators. A bronchodilator, such as albuterol, can improve the FEV1/FVC ratio as much as 10% to 15% in obstructive disorders. If there is no effect, the disorder is probably restrictive.

See “Interpretation of Pulmonary Function Tests” below for more information on abnormal values and possible pathologies.

Recording Steps for FEV1

1. We are still using the old screen. Make sure you have recorded all the values in the last exercise because your old screen will now be erased. Click Redo and erase the current FEV data.

2. Click Yes. You are actually performing a new experiment, but tricking the program into thinking it is a repetition of the old one. The next screen will give you the option to Record FEV again. Do not record until you have read the procedure below. If you get the dialog box shown in Fig. 11, click NO.

Figure 11. Dialog box warning before the opening of a new file.

This section requires the subject to take three measurements of FEV1 in a row.. The goal is to exhale the vital capacity as FAST as the subject can manage. The subject will need to do this three times with three normal breaths inserted between each forced, fast exhalation.

3. Read the steps below and PRACTICE the following procedure before actually recording:


a) Breathe normally until we signal you to inhale. (Let them breathe 3 cycles)

b) Inhale as deeply as you can and hold your breath for at least 2 seconds.

c) Exhale as FAST as you can and continue to exhale for at least 5 seconds.

d) Breathe normally until we tell you inhale again. (Let them breathe for 3 cycles)

e) Repeat steps b-d two more times.

Once the subject is clear on the directions, proceed with the recording.

4. Place the nose clip on the subject and click Record FEV.

5. Have the subject perform step 3 so that three forced expirations are recorded.

6. Click Stop. Your recording should look like the one below.

Figure 12. Recording of three Forced Vital Capacities (FEV1).

Measuring FEV1

1. Measure FEV1 by placing the I-Beam on the peak of your inhalation curve and dragging it over to the right as you watch the Delta time value in the measurement box increasing from zero. Remember you are trying to gather data only on the first second of exhalation. When the Delta T value reaches approximately 1.000 second, end the highlighting. You may overshoot the second slightly, or be slightly short of the second, getting values such as 0.980 or 1.010 seconds. These values are acceptable.


Figure 13. Placement of I-Beam for recording of FEV1 data.

2. Make a note of your P-P value for the one second highlighted.

3. Repeat steps 4 and 5 for all three forced exhalations and record your results in Table 5. Since FEV1 is a ratio, you must now record the FVC (Forced Vital Capacity) for each of the three trials.

4. Position the I-Beam on the peak of your first inhalation curve. Drag the I-Beam until it reaches the lowest point in the recording. Record the P-P value. Repeat these steps for the next two forced exhalations. Record your results in Table 5.

5. Calculate the percentage of vital capacity that can be exhaled in 1 second:

FEV1% = (FEV1/FVC) X 100

This is based on the average of the three forced exhalations.

6. Record your results on Table 5 and on the board or overhead projector with the rest of the class data.

7. To exit the screen or record from another individual, click Setup FEV.

8. The dialog box in Figure 14 should appear. Ignore the instructions and highlight any three-second area on your screen. (We are tricking it here, just go with the flow.)

Figure 14. Dialog box to be ignored.

9. Click Done. Click Yes. (More tricking)

10. A dialog box will appear giving you the option to Record from another Subject or to Quit (Fig. 15). At this point, you have finished recording for one individual. If you wish to repeat this for another group member, first check to see whether time will allow for it. If so, request a new filter and mouthpiece from your instructor.


Figure 15. Dialog box giving you the option to record from another individual or quit.

Table 5. Forced Expiratory Volume Data

Trial FEV1 (liters) FVC (liters)

1

2

3

Average

FEV1% = (FEV1/FVC) X 100 =

Interpretation of Pulmonary Function Tests

Accurate clinical interpretation of pulmonary function tests (PFTs) is complicated because of the nature of spirometry technology. Even under the most controlled clinical conditions, it can be difficult to get good, reproducible PFT results for an individual. Reproducibility is dependent on the effort put forth by the subject, the reliability of the equipment, calibration of the equipment and the experience of the operator. Variation is influenced by environmental factors such as temperature or barometric pressure.

If the evaluation is considered reliable, then the results must be compared to normative values depending on age, gender, body size and ethnicity.

• Age: loss of elasticity with age results in smaller lung volumes and capacities.

• Gender: Males usually have larger lung volumes and capacities than females, even when matched for age and height.

• Height and body size: Taller individuals will have larger lung volumes and capacities than shorter individuals, when matched for gender and age. When body size increases due to an increase in the body fat to lean body mass ratio (obesity), abdominal mass prevents the diaphragm from descending as far as it could, resulting in smaller lung volumes and capacities.


• Race: Hispanics, Native Americans, Asians, and African-Americans have different PFT results compared to Caucasians. Race appropriate PFT tables are available for comparison.

In general, PFTs place subjects into three categories: normal lung function, obstructive disease, or restrictive disease.

Obstructive lung disease patients have decreased airflow (decreased FEV1/FVC ratio) and usually have normal or above-normal lung volumes. Chronic obstructive pulmonary disease (COPD) encompasses this category and includes emphysema, chronic bronchitis, asthma, cystic fibrosis, and bronchiectasis, a disease that causes localized, irreversible dilatation of part of the bronchial tree. Involved bronchi are dilated, inflamed, and easily collapsible. Bronchiectasis most commonly results from bacterial infections.

Restrictive lung disease patients have decreased lung volumes or TLC with normal airflow (normal FEV1/FVC ratio, but with reduced values for both FVC and FEV1 individually. Conditions that lead to restrictive lung diseases include pleural disorders, alveolar disorders (pneumonia, cancer, or pulmonary edema), interstitial disorders that decrease the space between, neuromuscular disorders that affect the diaphragm or intercostals muscles (Guillain-Barré syndrome, myasthenia gravis, amyotrophic lateral sclerosis, otherwise known as Lou Gehrig’s disease, poliomyelitis), or skeletal disorders (kyphosis or scoliosis).

Glossary P-P- finds the difference between the maximum and minimum value in the selected area and

subtracts the minimum value found in the selected area. This is used to measure amplitude of the waves recorded or Tidal Volume (L/breath).

Delta T- The Delta Time measurement is the difference in time between the end and the beginning of the selected area. This is used to measure the time taken for a full breath in Liters.


Questions

1. Define tidal volume, respiratory rate, minute volume, inspiratory reserve, expiratory reserve, and vital capacity.

2. Compare your various lung volume measurements to those of others in the class. How do you account for differences? (Consider factors such as age, height, gender, smoking, etc.)

3. How do age, gender, height and body size affect normal lung volumes and capacities?

4. What is the significance of FEV1? Why must it be compared to FVC as FEV1% to be meaningful?

5. What is the physiological difference between pulmonary obstructive disorder and pulmonary restrictive disorder?

6. How can PFTs (Pulmonary Function Tests) help distinguish between these two types of pulmonary disorders?

7. Kim is 62" tall and 18 years old. Her results for her PFT in Physiology class were as follows:

Tidal Volume: 0.385 Liters Delta T for one breath: 4.5 seconds Vital Capacity 2.124 Liters FEV1 1.787 Liters

Calculate Kim’s Respiratory Rate and Minute Volume. Give units of measurement. Does Kim have a respiratory disorder? If so, is it restrictive or obstructive? Explain.

8. The medical records of Kyle and Beth may have gotten mixed up. One of these two individuals has asthma. Given the data below, who has asthma?

Kyle BethAge and Height 25 y.o./63 inches 27 y.o./64 inches Tidal Volume (L): 0.487 0.398 Delta T for one breath (s): 4.1 4.5 Vital Capacity (L) 4.124 3.80 FEV1 (L) 2.685 2.97


Exercise 9: Blood Readings: Silverthorn 4th ed, pg. 536-546, 798 – 799; Silverthorn 5th ed, 547 – 558, 804805.

Blood Typing The membranes of human red blood cells (RBCs) contain a variety of cell surface proteins called blood group antigens. The most important and best known of these are the A and B antigens, also called the ABO blood group, and the Rh antigen. When blood is transfused between a donor and a recipient, blood group antigens must be identified in order to ensure that their blood is compatible.

Blood group antigens are found only on RBCs and their presence is determined by an individual's genetics. An individual who does NOT inherit a particular blood group antigen will produce an antibody that recognizes that antigen as foreign. Most commonly, an antibody is produced after an antigen is introduced into the body, as with the Rh system. The ABO blood group is unusual in that an individual lacking a blood group antigen will automatically produce an antibody against the lacking antigen, even if that antigen has never been introduced into the recipient.

An antibody will specifically bind to the antigen it recognizes in an attempt to eliminate the antigen. This binding reaction is called agglutination (Fig. 1b). Agglutination of blood group antigens causes large clumps of RBCs and antibody to form, which can block and damage the small capillaries, especially in the kidneys. The resulting damage is called a transfusion reaction and can cause permanent kidney damage or even kill the recipient.

So how are these blood group antigens identified before a transfusion is made? A sample of blood is removed from the recipient and mixed in a dish with purified antibodies that are known to recognize a specific blood group antigen. If the purified antibodies cause the blood sample to agglutinate, then the RBCs in the blood sample carry the protein recognized by the antibody. See Fig. 1a for examples of agglutination.


1a.

1b.

Figure 1a. Agglutination reactions: Blood samples in the first column have been mixed with Anti-B antibody. Agglutination reactions with Anti-B antibody have occurred with Type B and Type AB blood, which both contain the B antigen. Blood samples in the second column have been mixed with Anti-A antibody. Agglutination reactions have occurred with Type A and Type AB blood, which both contain the A antigen.

Figure 1b. Antibodies are Y-shaped. Only anti-A antibodies bind to the A antigen on the RBCs.

A person with Type A blood has RBC's that carry the A antigen. Antibodies that recognize the A antigen are called anti-A antibodies. If Type A blood is mixed with anti-A antibodies, the RBCs will agglutinate. If Type A blood is mixed with anti-B antibodies, the RBCs will NOT agglutinate because the B antigen is not present. Conversely, if a blood sample agglutinates with anti-B antibodies but not with anti-A antibodies, then the blood sample contains Type B blood.

Where do these antibodies come from? They are naturally found in the blood of the opposite blood type. Remember that individuals with A antigen on their RBCs will carry anti-B antibodies and individuals with B antigen will carry anti-A antibodies. See Table 1 for clarification of which type of antigen and antibody is found in a given blood type. Type O blood contains neither A nor B antigen, and so can produce both Anti-A and Anti-B antibodies. Type AB blood contains both A and B antigen, and so will produce neither Anti-A nor Anti-B antibodies.

At one time, Type O blood was considered a universal donor since it contained neither A nor B antigens and would therefore not react with any blood group antibodies produced by the recipient. Today, it is recognized that Anti-A and Anti-B antibodies found in the plasma of Type O blood have the potential to produce a transfusion reaction with a Type A or Type B recipient and so is no longer given to recipients with other blood types.


Table 1. ABO Blood Groups

Blood Type Antigen on RBC Antibodies in Plasma

O Neither A nor B Both Anti-A and Anti-B

A A only Anti-B only

B B only Anti-A only

AB Both A and B Neither antibody

The Rh factor is separate from the ABO blood group. An individual carrying the Rh antigen is said to be Rh + while an individual without the Rh antigen is said to be Rh –. The + or – is added to the ABO blood type. For example, a person with Type A+ blood carries the A antigen, the Rh antigen, but not the B antigen.

Individuals who are Rh – will not produce Anti-Rh antibodies unless the Rh antigen has been introduced into their bodies. Rh antigen can be introduced into a woman's body if a fetus she is carrying inherits the Rh antigen from the fetus's father. Fetal RBCs carrying the Rh factor are too big to cross the placenta, so the mother is not exposed to the Rh factor until the baby is delivered. During delivery, or any break in the placenta, fetal blood can mix with the mother's blood and the mother can begin to produce Anti-Rh antibodies. When the mother becomes pregnant a second time, these antibodies proteins are small enough to cross the placenta and attack the Rh+ fetus, causing damage to fetus and possible miscarriage. This is referred to as erythroblastosis fetalis.

Differential Blood Cell Counts In a sample of blood, various formed elements are present. The ratios between red and white blood cells allow us to evaluate the health of individuals. Disruption of these ratios is an indication of disease. Anomalies in the shape of the cells may also be pathological. We will use blood smears and a compound microscope to identify four of these anomalies.

When looking at slide of healthy blood under the microscope one sees mostly red blood cells (erythrocytes), but also white blood cells (leukocytes) and platelets (thrombocytes). Less than 1% of the blood cells are white blood cells. The five types of white blood cells include neutrophils, eosinophils, basophils, monocytes and lymphocytes (Fig. 2). Each type has specific functions and a distinctive appearance. Doing a differential white blood cell count involves determining the relative abundance of the different types of white blood cells. Observing increases or decreases in the numbers of a certain type of white blood cells is helpful in diagnosing certain conditions.


Figure 2. All blood cell types arise from a common stem cell.

Blood Cell Abnormalities Sickle Cell Anemia Inherited abnormalities in hemoglobin may cause anemias. The red blood cells that contain abnormal hemoglobin may lose their shape or lose their ability to deliver an adequate supply of oxygen. In Sickle Cell, the red blood cells contain an abnormal form of hemoglobin, reducing the amount of oxygen in the cell, causing them to become crescent shaped (Fig. 3). These cells block and damage small blood vessels in the spleen, brain, kidneys and bones, reducing their oxygen supply. Sickle-cell cells are fragile and break up easily causing blocked blood flow to these organs.

Eosinophilia Increased numbers of eosinophils (Fig. 4), or eosinophilia, are most often associated with allergic diseases and the presence of parasites (such as worms).

Cancers of the Blood Chronic Myelogenous Leukemia (=chronic myeloid leukemia) is a condition where cell division becomes excessive in myeloid stem cells. This results in an abnormal release of leukocytes from bone marrow, spleen and lymph nodes. observe contain myeloid cells at all stages of maturation.

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Figure 3. Sickle-cell red blood cells

The blood slides that you

igure 4. Normal Eosinophil

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Lymphomas are malignant neoplasms of lymphoid tissue (lymph nodes, spleen, and other organs of the immune system). Lymphocytes are found in great numbers on these blood smears. Keep in mind that a high lymphocyte count in the blood could also be an indicator of infection or even a leukemia.


1. Blood typing with simulated blood.

2. Observe microscopic slides portraying various blood pathologies.

3. Perform a differential blood cell count

Getting Started

Blood Typing Materials:

• 4 blood typing slides

• 12 toothpicks

• 4 unknown blood samples:

Mr. Smith

Mr. Jones

Mr. Green

Ms. Brown

• Simulated Anti-A Serum

• Simulated Anti-B Serum

• Simulated Anti-Rh Serum

Procedure

1. Label each blood typing slide with the name of the sample:

Slide #1 – Mr. Smith

Slide #2 – Mr. Jones

Slide #3 – Mr. Green

Slide #4 – Ms. Brown


2. Place 3 drops of Mr. Smith's blood in each of the A, B, and Rh wells of Slide #1.

3. Repeat with Slides #2 – #4 and their respective samples.

4. Place 3 drops of Anti-A serum in each A well on all four slides.

5. Place 3 drops of Anti-B serum in each B well on all four slides.

6. Place 3 drops of Anti-Rh serum in each Rh well on all four slides.

7. Use a separate clean toothpick to stir each well for 30 seconds. Do not press too hard on the typing tray to avoid splattering.

8. Record the occurrence of agglutination in Table 2. If the sample appears grainy or opaque, assume agglutination has occurred. Determine the blood type based on your agglutination results.

9. When finished, throw away all toothpicks and rinse off and stack the typing trays.

Table 2. Blood Typing Observations

Anti-A Serum Anti-B Serum Anti-Rh Serum

Slide #1: Mr. Smith

Slide #2: Mr. Jones

Slide #3: Mr. Green

Slide #4: Ms. Brown

Table 3. Blood Typing Results

Antigens present Antibodies present Blood Type

Mr. Smith

Mr. Jones

Mr. Green

Ms. Brown

Questions:

1. Explain how you were able to fill out Table 2.

2. For each patient, determine what blood type(s) could be used if an emergency transfusion was needed. It is understood that he/she should receive only his own blood type under optimal conditions, but assume that he/she was in an


emergency situation, out of reach of a hospital, but close to a partially equipped clinic with some blood in its coolers.

Blood Slides Obtain one of each of these microscope slides:

1. Sickle Cell Anemia

2. Eosinophilia

3. Chronic Myelogenous Leukemia

4. Lymphoma

Observe the slides under the compound microscope. Use the descriptions below to help you to

determine what to look for. You should use immersion oil at 1000X magnification. Make a

drawing of each of these slides to hand in.

Sickle Cell Anemia Notice the large numbers of misshapen RBCs. The affected cells have the

characteristic sickle shape or will look like grains of rice.

Eosinophilia Observe the large numbers of eosinophils on the blood smear. The lobed nuclei

are stained purple and the cytoplasm has dark pink granulations. Neutrophils are also present.

Disregard the clumped “worm-like” aspect of the RBCs. This clumping was the result of the

preparation technique. Also disregard the many small purple-stained cell fragments throughout

the slide.

Chronic Myelogenous Leukemia- Notice the high counts of white blood cells throughout the

blood smear slide. Here too, the red blood cells on these slides appear clumped and out of shape.

Disregard this condition. Instead, concentrate on finding granulocytes (basophils, eosinophils

and neutrophils) and agranulocytes (monocytes) in all stages of development. The granulocytes

have visible stained granules within the cytoplasm of the cells.

Lymphoma – Observe the blood smear and notice the large numbers of lymphocytes in all

different stages of maturity. Neutrophils are also present and some of the red blood cells appear

crenated.


Differential Blood Cell Counts

For this activity you will first perform a differential white blood cell count to determine

the relative abundance of the different types of white blood cells in normal human blood. You

will then perform a similar analysis on a slide of blood from a person with a white blood cell

abnormality.

Using your text, and figures available in the lab you will identify the types of white cells.

Some general characteristics to help in differentiating the types are described below.

Neutrophil: Multi-lobed nucleus, clear cytoplasm (looks like a "Mickey Mouse Balloon"). Makes

up 65% of WBC.

Lymphocyte: Large round nucleus that fills most of the cell, a small amount of clear cytoplasm.

Makes up 25% of WBC.

Monocyte: Large cell with heart-shaped/ horseshoe shaped nucleus and clear cytoplasm. Makes

up 2-4% of WBC.

Eosinophil: Bi-lobed blue/purple staining nucleus with red staining granules in cytoplasm.

Makes up 4-5% of WBC.

Basophil: Bi-lobed nucleus with blue-purple staining granules in cytoplasm. Makes up <1% of

WBC.

Normal Blood Focus the normal blood slide at 400X magnification. Starting at one end of the

slide, scan back and forth as shown in Fig. 5, systematically locating white blood cells,

identifying them and tallying the different types you encounter in Table 4. Each group should

count a total of 25 white blood cells. If a particular white blood cell is squashed or ambiguous

looking, skip it and go on to the next one. We will compile the class data to get larger numbers

to use for calculating the relative percentage of each type of cell.


Figure 5. Scanning the blood slide

To determine relative percentage of neutrophils divide the number of those cells by the total number of white cells counted and multiply by 100.

For example:

Relative % = Number of neutrophils counted X 100 Total number of white cells counted


Table 4. Normal Blood

Group data Class data

Cell type # identified Relative % # identified Relative %

Neutrophil

Eosinophil

Basophil

Monocyte

Lymphocytes

Total = (25)

Eosinophilia Blood Repeat the steps above now using the Eosinophilia blood slides. Record your data in Table 5.

Analyze the abnormal blood smear as you did above after counting 25 white blood cells.

Table 5. Eosinophilia Blood

Group data Class data

Cell type # identified Relative % # identified Relative %

Neutrophil

Eosinophil

Basophil

Monocyte

Lymphocytes

Total = (25)

Based on your observations, describe what is abnormal about the blood:


Appendix A: Graphing Basics

Introduction

In most of the laboratory exercises in this course you will be generating data. How you treat and handle these data can make a big difference in your understanding of the material. One of the best ways of expressing data is a graph or plot. The purpose of this appendix is to familiarize you with the basic conventions of scientific graphing and how to use a computer spreadsheet to construct a graph.

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-10

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-901 2 3 4

ME

MB

RA

NE

PO

TEN

TIA

L (m

V)

TIME (ms) Figure 1. Action potential in a neuron

The graph in Figure 1 shows an action potential or "nerve impulse" as measured at the membrane of a nerve cell (neuron). It represents data that were collected over the course of an experiment. To someone experienced at reading graphs, this figure can summarize action potentials very clearly.

A graph is meant to express or demonstrate a relationship between two kinds of things. In Figure 1, the relationship is between time and membrane potential (this is simply the electric charge on the membrane in millivolts). Both time and electric charge can vary within limits and are therefore called variables. Therefore, this graph shows the relationship between two variables; time and membrane potential.

When the relationship is examined, it is clear that one variable changes independently, while changes in the other are dependent on the first. Time is going to pass no matter what the membrane potential and is therefore said to be the independent variable. The variation in membrane potential, on the other hand, depends upon how much time has elapsed since the membrane was stimulated. Membrane potential is therefore identified as the dependent variable.

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Male

Age in Years

Uni

ts o

f Est

roge

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Figure 2. Estrogen levels in human urine


The units of the variables are shown on lines called axes. One axis is a horizontal line, the X-axis; the other axis is a vertical line, the Y-axis. The point where the two axes meet or cross is the Origin. By convention, the independent variable is shown on the X-axis (horizontal) and the dependent variable is shown on the Y-axis (vertical). Note how these variables are plotted in Figure 1.

Figure 2 shows the relationship between the urine levels of Estrogen in the urine of girls and boys of different ages. Notice that again, time is the independent variable and plotted on the X-axis (horizontal). The levels of hormone depend upon the age of the individual, and therefore are considered dependent, and plotted on the Y-axis (vertical). You should also note that this graph clearly summarizes and differentiates relationships that would otherwise require many paragraphs verbal explanation.

Figure 3 is a graph in which time is not the independent variable. The independent variable in this case is the concentration of Carbon Dioxide in the inhaled air. The dependent variable is the volume of air inhaled per minute (Respiratory Minute Volume). Initial inspection of this graph tells us that as the CO2 concentration in the inspired air increases, the volume of air inspired per minute also increases. A closer inspection reveals that about a 20% increase in CO2 concentration results in more than a 400% increase in minute volume... highly significant.

Before going on, you should be able to explain why CO2 concentration is considered the independent variable, and minute volume the dependent variable. You should also be able to explain the relationship between the two just by looking at the graph.

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20

25

30

35

38 40 42 44 46 48 50Carbon Dioxide (mm Hg)

Res

pira

tory

Min

ute

Vol

ume

(l/m

in)

Figure 3. Respiratory minute volume as a function of Carbon Dioxide

Concentration


Now go back to the graph in Figure 1 and see what story it tells. Trace the line with your finger as you look at the values on the X- and Y-axes. There is no change in membrane potential (charge) until about 2 milliseconds into the measurement. For the next half millisecond, the charge changes from -70mV to +30mV. Then in the next millisecond, it returns to nearly what it was before. The point here is not to have you understand the action potential, but that you understand how to read a graph and appreciate how much information it can convey.

Table 1. Oxygen intake at different speeds

Activity Speed (M/sec)

O2 intake (ml/min)

Standing 0.00 350 Walking 0.61 548 Walking 1.18 818 Walking 2.04 1843 Walking 2.38 2732 Running 2.86 3080 Running 3.41 3205 Running 4.05 4175 Running 4.52 4055 Running 4.70 4080

Suppose you figure that in the act of running, a person would consume more Oxygen than when walking; or more generally, that the faster a person is moving the more Oxygen they would consume.

In order to see if your idea is correct, you measure a person's Oxygen consumption while they are walking or running at different speeds. The first job after collecting the data is to present it in a neat easily understood Table, such as the one shown in Table 1.

Inspection of these data reveals that indeed, as a person's speed increases, so does their Oxygen consumption. It is difficult however, to get much more information from the table without a great deal of concentration.

This is where the graph comes in; because finer points of the relationship become obvious when the data are presented in a graph as in Figure 4. Exactly how this graph is put together will be explained later, but for now look at the information that is readily apparent.

First, the graph clearly shows that Oxygen consumption increases as the speed of the individual increases. A closer look reveals even more information. The line connecting the points is relatively flat until 1.2 meters per second, when it begins to sharply curve upwards. It levels off again at 2.4 M/sec, but takes another sharp increase between 3.4 and 4.4 meters per second.

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Walking

Running

Oxy

gen

Con

sum

ptio

n (m

l/min

)

Speed (m/sec)

FIGURE 4. Oxygen Intake at Various Speeds. Data points connected by lines.

What this shows is that as the speed increases, Oxygen consumption does not increase in a regular way. Rather, it increases more rapidly during a fast walk or run.


Think about this relationship for a moment. Also consider why people with severe respiratory problems such as emphysema can't walk as fast as their muscles and coordination might allow.

How to construct a graph from a data set

1. Identify the independent variable. In the example above, Oxygen consumption depends upon speed. Thus, speed is the independent variable. It would be incorrect to assume that a person’s Oxygen consumption sets the speed at which they move (at some point however, Oxygen consumption can limit a person's speed).

2. Once identified, the independent variable will be plotted on the X-axis (horizontal) and the dependent variable on the Y-axis.

3. Decide on the numerical scale for each axis. The origin should be less than or equal to the smallest value of that particular variable, while the largest value on the axis must be equal to or larger than the highest value of the variable.

4. Draw the axes with appropriate tic marks and numbers. Label the axes. 5. Plot each data

point. Locate its value on the X-axis and then move vertically to its appropriate Y-axis value. Mark these points so that they stand out.

6. Draw a line between the points. How you do this depends upon the data and what you want the graph to emphasize. Compare Fline connecting the poWithout any line (Fig.variables, but not the d

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igures 4 and 5. The only difference between them is the ints, but it makes the message slightly different in each. 5), you see a general positive relationship between the etails of the relationship.

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FIGURE 5 Oxygen Intake at Various Speeds.

Points Unonnected by Lines to Show General Relationship.O

xyge

n C

onsu

mpt

ion

(ml/m

in)

Speed (m/sec)

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7. Figure 6 has a straight line drawn in, which best represents the average relationship. Compare Figures 4 and 6, and note the difference in message. Which of these options you choose depends upon the data and the message you want conveyed.

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FIGURE 6 Oxygen Intake at Various Speeds.

Line Drawn In to Represtent General Relationship.O

xyge

n C

onsu

mpt

ion

(ml/m

in)

Speed (m/sec)


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