Laboratory Manual Fall 2014

Rhode Island College Biology 335

Contents

1. Safety in the Laboratory 3

2. Measurements and Computations 5

3. The Cell: Transport Mechanisms and Cell Permeability 13

4. Introduction to iWorx 15

5. Physiology of Skeletal Muscle 27

6. Skeletal Muscle Physiology: Computer Simulation 39

7. Reflexes 41

8. Cardiology with a Vertebrate Heart 51

9. Electrical Properties of the Heart 65

10. Circulatory Physiology 79

11. Mechanisms of Breathing 89

13. Basal Metabolic Rate 111

14. Renal Physiology — The Function of the Nephron 123

12. Restrictive and Obstructive Lung Disease 103

Page 1 Revised Fall 2014


The laboratory is a safe place to work and study. It remains safe when the individuals working there practice the conven-tional rules of laboratory safety. Laboratory work frequently requires the use of reagents, equipment, and organisms that are potentially dangerous for all personnel in the laboratory. On entering the laboratory, you assume responsibility for your own safety and the safety of your neighbors. The ad-vice that follows represents accepted procedure and describes the behaviors assumed to be characteristic of anyone working in a laboratory.

Good laboratory work requires advanced preparation on the part of all that are present. The instructions and directions for a specific activity require careful study beforehand. You need to know: 1) What to do, 2) How to do it, and 3) Why it is to be done in the prescribed manner. Thorough preparation improves the quality, efficiency, and safety of your work. Prepare a set of notes for the materials and procedures organized in the form of a flow chart. Highlight notes that describe safe-ty procedures.

Instructors usually provide a briefing before the start of a laboratory exercise. Unless the instructor tells you otherwise, it is advisable to wait for that commentary before beginning your work. The briefing provides the opportunity for: describing changes in instructions, demonstrating techniques, explaining the procedures for the proper use of instruments, and highlighting safety precautions. Be sure that you understand, and follow exactly, the special procedures for the correct disposal of hazardous materi-als, biological wastes, and body fluids. The briefing is a chance for you to ask questions about equipment and procedures that you don’t under-stand. Making changes in the procedures and using materials other than those described in the instructions can be disas-trous. On the other hand, changes based on new in-sights can be very valuable because they may increase the effectiveness of an exercise. Therefore, be sure to

discuss your ideas with the instructor at the time they occur to you or before leaving the laboratory.

The laboratory bench must be clean and organized, and other extraneous items are a safety hazard when stacked on the surface of the bench. Remove unnecessary items

from the bench top and place them in a secure storage area. The cabinet in the bench pedestal is a good place to store calculators, purses and other valuables.

The laboratory in which you will be working is a general purpose teaching laborato-ry. Instructors and students from different courses make use of the room. As a result, a wide variety of substances and equipment is constantly being moved into and out of the laboratory. Materials remain and form residue that is potentially pathogenic, capable of causing personal injury, and likely to soil or damage your belongings. Check the bench top as a precaution-ary measure before assembling the materials needed for the laboratory activity. Keep your hands away from your mouth. Do not eat or drink in the laboratory, and never pipette by mouth suction.

When gathering materials from central supply areas, label the containers for their transport beforehand. Read the labels on the stock containers TWICE before taking what you need. Immediately replace caps, stop-pers, and covers for all containers. The stock is to re-main in the supply area. Excess amounts of media and reagents are not to be returned to stock supplies because

contamination of the stock supply may result from this process. Materials and equipment are to remain in the laboratory at all times.

A general knowledge of safety procedures is inadequate. You must know what to do in a particu-lar set of circumstances. Quick action is vital when accidents occur that result in materials entering the eyes, fire, broken glass, chemical spills, or injuries to the skin. Therefore, thoroughly study the proce-dures that are to be followed for each of the condi-tions listed below.

Eye Injury. Go immediately to the eye wash station and flush the material from the eyes. When one eye is affected, tilt the head to prevent the stream of water from introducing the material into the other eye. Continue the

Laboratory #1 Safety in the Laboratory

The keys to working safely in

any laboratory are organization,

neatness and being prepared!

Please note that the material in this manual is a compendium of exercises originating from multiple sources. The computer simula-tion laboratories are modifications of exercises contained within the PhysioEx Laboratory Manual. Contributors to this manual in-clude Frank Dolyak, Kenneth Kin-sey, Edythe Anthony, Jerome Montvilo, Eric Hall and others.

Biology 335 Human Physiology: Safety


procedure until directed to stop. Individuals in the immediate area are to clear a path to the eye wash station and take the initiative to lead the injured person to the wash station. NOTE: If you wear contact lenses and also have a framed set of lenses, it is advisable to wear the framed lenses in the laboratory.

Fire. When entering the laboratory for the first time, determine the shortest exit routes from your laboratory station to the adjoining hallways. Locate the exits leading from the hallways to the outside of the building. If the laboratory has fire extinguish-ers and fire blankets, the instructor will explain their proper use and show you their locations.

Broken glass. To pick up broken glass use a dust pan and brush. Gather together small slivers and chips by using a crumbled wad of wet paper toweling or wet cotton and labor-atory tongs. Discard the glass into the special container des-ignated by the instructor.

Chemical spills. Clear the area immediately and tell the instructor. Promptly flush the area with water when spilled material con-tacts your personal belong-ings, clothing or skin.

CAUTION! Sometimes alternative procedures are required, and the instructor will describe them during the laboratory briefing.

Skin injuries. Immediately report accidents that puncture, cut, abrade or burn the skin. In each laboratory, there is a first aid kit for the immediate treatment of minor injuries. Any occurrence is significant, and the injured person should consult the college Health Service.

Long hair, long or baggy sleeves, large bracelets, and long dangling necklaces are safety hazards. When nec-essary, tie back long hair and roll up long baggy sleeves. Store jewelry and other personal items in a secure area. Sandals should not be worn in the laboratory. They do not provide adequate protection against sharp objects or chemical spills.

Before leaving the laboratory, be sure to clean and re-stock the individual bench area. Arrange the materials and equipment as they were initially, and discard wastes into the proper containers for disposal. CAUTION! Disposal of reagents, biological wastes and body fluids in a casual manner endangers yourself and those around you. Unless told to do otherwise, solid waste is not discarded into the sink or solution poured down the drain. You should strive to leave the laboratory station

as you would like to find it. Your colleagues and instructors appreciate your cooperation. Thank you!

Eye Wash Station

First Aid Kit and Fire Blanket


Biology 335 Human Physiology: Safety


1. Review the metric system. 2. Practice converting between different met-

ric units 3. Review simple computations such as using

ratios and proportions. 4. Review the use of significant figures.

At this point in your science education you should be intimately familiar with the standard units of measure-ment used throughout the world. You should also pos-sess a concept of the relationship between these interna-tional standards and the measures commonly used in the USA. This latter is especially important for health care providers who must translate a patient’s ideas of measure into these standard measurements for recording in the patient’s records. In fact, the measurement and admin-istration of medications is always done in metric units. When you pick up a prescription at the drug store it will be measured in international units and if you are admin-istering medications in a clinical setting you will also be using international units. Of course the international standard we are discussing is the metric system which is based on the decimal system where the units are related to each other by powers of 10. The following prefixes have a constant meaning and are in general use through-out the metric system: Kilo (k) = thousand (times) [103] Centi (c) = one-hundredth [10-2] Milli (m)= one-thousandth [10-3] Micro () = one-millionth [10-6] Nano (n) = one-billionth [10-9] A. Length Units Instead of the yards, feet and inches of the English sys-tem, the metric system expresses the length of an object in meters, centimeters and millimeters. Still smaller units are micrometers and nanometers. For comparison, a human erythrocyte is about 7 micrometers in diameter. Table 1 presents some commonly used metric length

Laboratory #2 Measurements and Computations

Objectives

I. The Metric System

Biology 335 Human Physiology: Measurements and Computations

units and their interchangeability. B. Mass Units Instead of weight (pounds and ounces), the metric system expresses mass in grams and multiples thereof. See the conversion table on the next page. It might be more flattering to think of your weight in kg (a 200 lb man or woman has a mass of about 91 kg), but your actual mass is the same regardless of the units you use to express it. C. Volume Units The metric unit of volume is the liter (l; slightly more than a quart) and its multiples. See the table on the next page. D. Time Seconds are divisible into milliseconds (ms) and mi-croseconds (s). There are 1000 ms in 1 s and 1,000,000 s in 1 s. E. Electrical potential difference Volts (V) are divisible into millivolts (mV) and micro-volts (V). F. Temperature Scales In the Fahrenheit scale (still used in the USA), water boils at 212 ºF, and freezes at 32 ºF, and the range comprises 180 units. In the Celsius (centigrade) scale the boiling point of water is 100 ºC and the freezing

Table 1. Metric and English Equivalents

How many English units in a m, l or g?

m 1.093 yd 3.279 39.37 in

l 0.264 gl 1.057 qt 2.114 pt

g 0.0022 lbs 0.035 oz

Continued on page 7. Page 5 Revised Fall 2014

To Convert: To: Mul ply by:

ng or nl or nm g or l or m 0.001 (1 X 10‐3)

ng or nl or nm mg or ml or mm 0.000001 (1 X 10‐6)

ng or nl or nm g or l or m 0.000000001 (1 X 10‐9)

ng or nl or nm kg or kl or km 0.000000000001 (1 X 10‐12)

g or l or m ng or nl or nm 1000 (1 X 103)

g or l or m mg or ml or mm 0.001 (1 X 10‐3)

g or l or m g or l or m 0.000001 (1 X 10‐6)

g or l or m kg or kl or km 0.000000001 (1 X 10‐9)

mg or ml or mm ng or nl or nm 1000000 (1 X 106)

mg or ml or mm g or l or m 1000 (1 X 103)

mg or ml or mm g or l or m 0.001 (1 X 10‐3)

mg or ml or mm kg or kl or km 0.000001 (1 X 10‐6)

g or l or m ng or nl or nm 1000000000 (1 X 109)

g or l or m g or l or m 1000000 (1 X 106)

g or l or m mg or ml or mm 1000 (1 X 103)

g or l or m cg or cl or cm 100 (1 X 102)

g or l or m dg or dl or dm 10 (1 X 10)

g or l or m kg or kg or km 0.001 (1 X 10‐3)

kg or kl or km ng or nl or nm 1000000000000 (1 X 1012)

kg or kl or km g or l or m 1000000000 (1 X 109)

kg or kl or km mg or ml or mm 1000000 (1 X 106)

kg or kl or km g or l or m 1000 (1 X 103)


point is 0 ºC, with a range of 100 units. Thus one Cel-sius unit of temperature is larger than one Fahrenheit unit, specifically 180/100 or 9/5 greater. To convert a Fahrenheit temperature to Celsius, use the following formula: To convert a Celsius temperature to Fahrenheit, use the following formula: H. Problems

Solve the problems found on the accompanying work-sheet.

A. Ratios A comparison of two numbers is called a ratio. For instance, if a tank of frogs contains 150 green frogs and 75 brown frogs, the ratio of green to brown frogs is 150 to 75, which may be written 150/75 or 150:75. This ratio is a fraction which should be reduced to the lowest common values (2/1 or 2:1). [It may be expressed as a single number by dividing it out (in this case “2”). A denominator of 1 is then assumed.] When two numbers are expressed as a ratio their units must be the same. For instance, if a rabbit has a mass of 1.5 kg and a frog has a mass of 100 g, one could express the ratio of their masses, but only after con-verting one of the measurements to the units of the other. Since 1.5 kg = 1500 g, we can express the ratio in grams as 1500: 100 or 15:1. Conversely, we could convert g to kg, getting 0.1 kg for the weight of the frog. The ratio would be the same in kg, i.e. 1.5:0.1 = 15:1.

B. Proportion A proportion is a statement of the equality of two rati-os and can be expressed in this way:

We state that “a is to b as c is to d”. If the numerical value of three of the four terms is known, the fourth can be determined by the following formula (this is called cross multiplying): For instance, assume that for every liter of blood pumped by the heart, 300 mL of this blood goes to the kidneys. When the heart pumps harder and puts out 3 liters of blood in the same amount of time, you wish to know how much blood goes to the kidneys (assuming no independent change in kidney flow). You would solve the problem by using the following proportion:

C. Calculation of an Arithmetic Mean Performing a physiological or any scientific experi-ment involves the collection of a great deal of numeri-cal data. After collecting the data, it is usually neces-sary to determine the significance of the data. Often we want to know if the experimental procedure pro-duced an effect which was different from a control procedure. In order to compare the experimental group of data with the control group it is first neces-sary to condense the data, that is, calculate the arith-metic mean. The arithmetic mean is simply the aver-age value of the group data and is calculated as:

II. Computations and Presentation of Data

When you convert from a larger unit to a smaller unit move the decimal to the right. When you convert from a smaller unit to a larger unit move the decimal to the left:

Large Small


Where:

= mean

= sum x = each individual datum

x = sum of all individual data of one group N = number of data values in the group For instance, a group of six rats was given thiouracil for two weeks in order to repress the functioning of the thyroid gland. A valid question could be raised, “Does administration of thiouracil cause atrophy of the thy-roid gland”? In order to answer this, you could deter-mine the mass of the thyroid gland from each of the six experimental rats. These data would have to be com-pared with similar data from a control group of rats. The first step would be to compute the arithmetic mean of the experimental data and compare it with the arith-metic mean of the control data, as follows: 1. Experimental thyroid mass (mg): 157 174 205 180 181 168 Σx = 1065

= 1065 ÷ 6 = 177.5 mg

177.5 mg is the experimental mean. The next step would be to compute the mean of the control group, in the same way: 2. Control thyroid mass (mg): 160 154 190 142 201 179

Σx =1026 = 1026 ÷ 6 = 171 mg

171 mg is the control mean. By comparing the two arithmetic means, we would conclude that there was no atrophy of the thyroid gland caused by the procedure. Suppose we had found a larger difference between the experimental and control means. Let us assume a dif-ferent set of control values, such that the control mean is 300 mg. In this case, the experimental mean of 177.5 mg is so much smaller than the control mean that we might be justified in concluding that atrophy of the thyroid had been produced by the thiouracil. In many biological experiments the results are not so clear cut. We cannot tell by simply inspecting the means, whether there is a significant difference be-tween the experimental mean and the control mean. In those cases it is necessary to use statistical procedures which will tell us the probability that there is, or is not, a significant difference between the two means. D. Problems Complete the problems included on the following worksheet.

The data gathered in the Human Physiology laboratory is fairly simple and does not require complex graphing techniques or expertise. At the same time there are some things that a student should be aware of when graphing data. The following are general rules that should be used whenever you create a graph (and must be followed when graphing data in this class). 1. You must have a title at the top of the page 2. You must properly label the X and Y axes 3. The X axis (horizontal) is the independent varia-

ble (Time or temperature in the Physiology labor-atories)

4. The Y axis (vertical) is the dependent variable (what you measured and recorded as your data)

5. Labels MUST always include units. Examples: Time (min), Heart rate (Beats/min) 6. Please make every effort to fit the data optimally

to the range of each axis. 7. Use the space provided on the graph paper to opti-

mally display your results

X


Biology 335 Human Physiology: Measurements

Page 8

X

Revised Fall 2014

III. Graphing

xNX =

X

8. Use the appropriate graph type 9. For continuous independent variables (time) use

a line graph 10. For discrete independent variables (temperature)

use a bar graph 11. When a graph contains two discrete data sets on

one graph make sure there is an appropriate label or legend to make it clear.

12. Finally, follow the guidelines provided in the manual.

See the provided examples.

All measurements are approximations—no measur-ing device can give perfect measurements without experimental uncertainty. By convention, a mass measured to 13.2 g is said to have an absolute uncer-tainty of 0.1 g and is said to have been measured to the nearest 0.1 g. In other words, we are somewhat uncertain about that last digit—it could be a “2”; then again, it could be a “1” or a “3”. A mass of 13.20 g indicates an absolute uncertainty of 0.01 g. The number of significant figures in a result is simply the number of figures that are known with some de-gree of reliability. The number 13.2 is said to have 3 significant figures. The number 13.20 is said to have 4 significant figures. A. Rules for deciding the number of significant figures in a measured quantity: (1) All nonzero digits are significant:

1.234 has 4 significant figures, 1.2 has 2 significant figures.

(2) Zeroes between nonzero digits are significant:

1002 has 4 significant figures, 3.07 has 3 significant figures.

(3) Zeroes to the left of the first nonzero digits are not significant; such zeroes merely indicate the position of the decimal point:

0.001 has only 1 significant figure, 0.012 has 2 significant figures.

(4) Zeroes to the right of a decimal point in a number are significant:

0.023 has 2 significant figures, 0.200 has 3 significant figures.

(5) When a number ends in zeroes that are not to the right of a decimal point, the zeroes are not necessarily significant:

190 may be 2 or 3 significant figures, 50,600 may be 3, 4, or 5 significant



Page 9

IV. Significant Figures

Revised Fall 2014

figures.

The potential ambiguity in the last rule can be avoided by the use of standard exponential, or “scientific,” no-tation. For example, depending on whether 3, 4, or 5 significant figures is correct, we could write 50,600 calories as:

5.06 × 104 calories (3 significant figures) 5.060 × 104 calories (4 significant figures), or 5.0600 × 104 calories (5 significant figures). B. Rules for mathematical operations In carrying out calculations, the general rule is that the accuracy of a calculated result is limited by the least accurate measurement involved in the calculation. (1) In addition and subtraction, the result is rounded off to the last common digit occurring furthest to the right in all components. For example,

100 (assume 3 significant figures) + 23.643 (5 significant fig-ures) = 123.643,

which should be rounded to 124 (3 significant figures). (2) In multiplication and division, the result should be rounded off so as to have the same number of signifi-cant figures as in the component with the least number of significant figures. For example,

3.0 (2 significant figures ) × 12.60 (4 significant figures) = 37.8000

which should be rounded off to 38 (2 significant fig-ures).

C. Rules for rounding off numbers (1) If the digit to be dropped is greater than 5, the last retained digit is increased by one. For example,

12.6 is rounded to 13.

(2) If the digit to be dropped is less than 5, the

last remaining digit is left as it is. For example,


(3) If the digit to be dropped is 5, and if any digit fol-lowing it is not zero, the last remaining digit is in-creased by one. For example,


(4) If the digit to be dropped is 5 and is followed only by zeroes, the last remaining digit is increased by one if it is odd, but left as it is if even. For example,

11.5 is rounded to 12, 12.5 is rounded to 12.

This rule means that if the digit to be dropped is 5 followed only by zeroes, the result is always rounded to the even digit. The rationale is to avoid bias in rounding: half of the time we round up, half the time we round down. D. Rules for counting Counted numbers have an infinite number of signifi-cant figures:

10 notebooks + 285 notebooks = 295 notebooks



Page 10

This section on significant figures was taken in part from: http://www.chem.sc.edu/faculty/morgan/resources/sigfigs/index.html For great practice see: http://science.widener.edu/svb/tutorial/sigfigures.html

Revised Fall 2014


Female (g) Male (g) 209 225 222 198 456 356 256 235 185 456 Ratio: 7. Your rats are to be fed a spe-

cial diet in which each 70 g portion of chow will contain 45 g of protein.

a. How much protein would be

present in 500 g of rat chow?

b. What is the ratio of protein

to non-protein in the diet? 8. When you want to quickly

measure a person’s pulse rate you don’t usually count the number of pulses in a full mi-nute. You count the number of pulses in 6, 10, or 30 seconds. You can do this because you know that if you set up a proper ratio you can quickly and easily calculate the pulse rate in pulses per minute.

Measure your pulse rate by

counting the number of pulses

1. Length a) 2.4 cm = _________ m b) 264 cm = _________ mm c) 23 m = _________ mm 2. Mass a) 0.85 g = _________ kg b) 5.3 g = _________ mg c) 280 ng = _________ g 3. Volume a) 53 L = _________ mL b) 7.95 L = _________ L c) 0.058 mL = _________ L 4. Time a) 0.120 sec = _________msec b) 240 msec= _________ sec c) 0.059 msec= _________ nsec 5. Temperature a) 72 ºF = _________ ºC b) 98.6 ºF = _________ ºC c) 2 ºC = _________ ºF

6. You have 5 female rats and 5 male rats, their masses are given below. What is the mean ratio of mass of the male to the female rats? Note: all ratios should be simplified to x:1.

Laboratory #2 Worksheet Name: Date: Section:

I. The Metric System

II. Computations

in 10 seconds (use your left in-dex finger applied to the radial artery of your right wrist).

Pulses per 10 seconds: _________ Calculate the number of pulses in

60 seconds (1 min.) using the following ratio:

(or just multiply by 6)

Pulse rate (pulses/min): ________ 9. On an EKG strip a nurse deter-

mines that a patient’s heart is beating 5 times every 4.5 sec-onds.

Calculate the time for each

heart beat: ______________ Calculate the patients heart rate

in beats/min: ____________ Hint: Divide 60 by the time per beat.

10. On the following page, graph the data set shown below:


Page 11

Pulses

10 =

? Pulses

60 sec

Revised Fall 2014

Heart Rate During Exercise

Time (min) Heart Rate (beats/min)

0 70

3 78

5 88

10 102

Worksheet score: ____________

Laboratory #2 Worksheet Name: (continued)


Laboratory #3 Cell Transport Mech-anisms and Permeability

After completing the following simulation you should firmly understand the concepts of simple diffusion, facilitated diffusion, osmosis and active transport across a cell membrane.

The fluids that bathe all of the cells of the body are water-based (aqueous) solutions. Within any solution the movement of the molecules dissolved within the solution (solutes) is driven by the random collisions of molecules in the solution. These collisions cause mol-ecules which are collected together to be pushed apart. This is diffusion. The movement of molecules from a point of high concentration to a point of low concen-tration is because of random molecular motion. If a semipermeable membrane (like a cell membrane) blocks the movement of solutes (into or out of a cell) but not the movement of the solvent, the solvent will diffuse from a point of high solvent concentration (low solute concentration) to a point of low solvent concen-tration (high solute concentration). This is called os-mosis.

The movement of molecules across a cell membrane can be passive (requiring no direct energy) or active (requiring energy in the form of ATP). Passive transport includes facilitated diffusion and filtration. Facilitated diffusion is the movement of molecules from a point of high concentration on one side of a cell membrane to the other side through protein chan-nels in the membrane. Filtration is the movement of molecules, driven by hydrostatic pressure, through protein channels (pores) across a biological mem-brane.

Active transport processes involve protein pumps lo-cated in cell membranes which utilize energy released by the hydrolysis of ATP. This energy is used to move molecules from one side of a cell membrane to the other from a point of low concentration to a point of higher concentration. In this computer simulation we are going to examine

simple diffusion through a nonliving semipermeable membrane. This is called dialysis. We will also ex-

amine the diffusion of water across such a membrane (osmosis) as well as studying the processes of filtra-tion and active transport. 1. Insert the PhysioEx 9.0 CD-ROM into the CD-

ROM drive of the computer or access the Physio-Ex folder on the desktop.

2. If you started with the CD-ROM a browser win-dow with the PhysioEx opening page should open. If you started with a folder on the desktop click on the StartHere icon .

3. Then click on “Access PhysioEx 9.0” to start the program.

4. Once the PhysioEx 9.0 windows opens click on “Exercise 1: Cell Transport Mechanisms and Per-meability.

5. Beginning with the Overview, complete the Ac-tivities. At the end of each activity you are given the option of saving your work in a .pdf file. Do so, and when complete, submit the files to your instructor (via email or hard copy—whichever the instructor prefers). Save the files with unique file name such as:

Hallsec03pex-01-01 Hallsec03pex-01-02 Hallsec03pex-01-03 Hallsec03pex-01-04 Hallsec03pex-01-05

Make sure the filename includes your name, sec-tion number and the exercise (-01) and activity number (-01, -02, etc.) that you are submitting.

Objectives

Background

Starting the Program

Biology 335 Human Physiology: Transport


http://www.medicinenet.com/dialysis/article.htm#1whatis

Hemodialysis

Artificial kidney machines have been developed that make use of dialysis to purify the blood of persons whose kidneys have ceased to function. Known as hemodialysis, this procedure has saved the lives of many persons suffering from renal failure. In such machines, blood is circulated on one side of a semiper-meable membrane (often cellophane) while a special dialysis fluid is circulated on the other side. The dial-ysis fluid must be a solution that closely matches the chemical composition of the blood. Metabolic waste products such as urea and creatinine diffuse through the membrane into the dialysis fluid and are discard-ed, while loss by diffusion of substances necessary to the body (such as sodium chloride) is prevented by their presence in the dialysis fluid. From: http://www.answers.com/topic/dialysis

http://www.lrh.com.au/home/orginfo/departments/dialysis/1391-image003.jpg



Laboratory #4 Introduction to iWorx

1. Become familiar with the iWorx recording and stimulating apparatus.

2. Examine the effects of exercise on pulse rate and blood flow

3. Examine the effect of increasing stimulus fre-quency on threshold (summation).

A stimulus is a change in the environment to which a cell, organ or organism is sensitive. Irritable (or excit-able) cells, such as neurons, muscle cells, and glandu-lar cells, can respond to such a change if the stimulus is of an appropriate type for a particular cell. (Environmental changes can be thermal, chemical, physical, electrical, etc.). For a cell to respond, the stimulus must also be of sufficient magnitude (i.e., at or above a threshold level of intensity). Some cells, such as skeletal muscle cells, will respond to such stimulation in a manner that can be readily observed. Other cells, such as neurons, are responsive to stimuli, but the response cannot be detected without sophisti-cated instrumentation. Because of the relative ease of observing and record-ing contractions of skeletal muscle, we plan to use this tissue in another laboratory as a means to explore stimulus-response relationships. Specifically, we will examine how the gastrocnemius muscle of a frog re-sponds to various types of electrical stimulation. To be successful in exploring these responses, it is im-portant to learn how to use an electronic stimulator to deliver carefully controlled electrical stimuli. The parameters of electrical stimuli that physiologists often manipulate are voltage (strength or intensity of stimu-lus), frequency of application (number of stimuli de-livered per second) and duration of each individual stimulus. It is also important to record the responses to such stimuli. The iWorx 214 data acquisition system re-ceives electrical inputs via various electrodes and sen-sors which plug into the front panel of the iWorx 214.

This unit also has a built-in stimulator (red) which can be used to mimic electrophysiological events in order to record and analyze electrical responses. The basic system is illustrated in Figure 1. This data collection unit interfaces with the computer via the LabScribe software which permits the display and analysis of a wide variety of physiological responses including frog skeletal muscle contraction, human heart rate, pulse, lung volumes, etc. Recordings of physiological events and their analysis can be saved to the computer desktop and/or printed from the computer.

Turn on the iWorx 214 console using the power button on the rear panel. On the computer, start the Lab-Scribe software by double clicking on the icon located on the desktop. After the software is loaded, select Load Group from the Settings menu. When the dia-log box appears, select 4 Intro to iWorx and then click Load or Enter. Click on the Settings menu again and select the Introduction to iWorx Record-ing settings file. Figure 2 shows the resulting screen. Notice that each channel has its own (white) recording area with a colored bar above it containing a title, Au-toscale and add function select buttons, and the volt-age value. Above the top channel colored bar is a time value, the sampling speed, the display time, the Mark comments, a T2-T1 value and the Record button.

Objectives

Background

Biology 335 Human Physiology: Introduction to iWorx

Figure 1. iWorx 214 Front and Back.

Learning To Use LabScribe-The Basics

Screen Time The default value for the time a signal crosses the screen is 10 seconds. This display time can be changed by clicking the display controls on the Lab-Scribe toolbar. To demonstrate this: 1. Click on Record then

after a few seconds click on Stop. 2. Click the left icon (big mountain) and notice that

the trace spreads out — the display time is five seconds.

3. Click the right icon (double mountains) twice and see that the display time increases to 20 sec-onds.

4. Click the left mountain icon once to return to a 10-second display time.

Marks The recording can be annotated by adding marks in two ways: 1. When not recording, two blue vertical lines or

cursors will overlay the screen (Figure 2). These cursors can be used to make measurements of your data. However, if you type a comment in the Marks area at the top of the screen using the key-board and press the Enter key, the comment will be entered in the lower margin at the left cursor (after you hit the Record button).

2. While recording, you can type short de-

scriptions into the Marks area and after hitting the Enter key the com-ment will appear on the recording. The comments associated with a mark can be moved vertically by clicking on the comment and dragging it using the mouse. Comments in a given view can be reset to the lower margin by select-ing Reset Marks under the View menu. You can use the single cursor to place comments or marks anywhere in your recording. Just move the cursor where you want the mark to appear, type your comment into the marks space and hit Enter.

You can also use marks to move through your record-ed data. Pull down the Windows menu and select Marks, or click the Marks icon on the toolbar. This will bring up a list of all of the marks you have typed into the record. Click on the time or comment for the mark you wish to go to and then press Go To. You can also delete marks from this menu if you inadvert-ently place a mark in the wrong position. Measurements Measurements are made using the cur-sors. These are vertical blue lines that span all channels and can be called up using one of the cursor icons. Using two cursors (left icon button) allows you to de-termine the difference in time or voltage between any two points on the recording. Click and drag the lines to the left or right to display the difference in: Time (horizontal distance) is measured by deter-

mining the time between the positions of the cur-sors. This difference is labeled T2-T1 and is dis-played at the top right next to the Record button.

Voltage (vertical distance) is measured by de-

terming the difference between the two small cross-shaped indicators on the cursor lines. This difference is labeled V2-V1 and is shown in the area on the upper right margin of each channel.


Figure 2. LabScribe Introduction to iWorx screen


Note: If the iWorx 214 unit and computer are not communicating you will receive an error. If this hap-pens, click on the “Tools” menu then “Find Hard-ware”. A small window should appear that says “Hardware Found=iWX114”. Click OK and proceed.

Cursors

Journal Although there is a built in journal feature in the Lab-scribe software, you should use Microsoft Word to rec-ord your data journal for all iWorx laboratories in this class. To start Microsoft Word, click on Start on the Windows task bar then click on the Word icon. Keep Word open while you work and you can copy your Lab-scribe screen using Ctrl-C and paste it into Word using Ctrl-V. This document can be saved to a USB Flash Drive or emailed to yourself or your instructor. Make sure you save your data file and journal frequently. For every exercise which utilizes the LabScribe software you should open a new journal and enter your name, lab partner’s names, exercise, lab section and date in the following format: Your Name Lab Partner’s names Exercise Number (Chapter in the manual) Laboratory Section Date As you complete each exercise you can add headings and insert your results directly into the journal. At the end of the lab you should print it immediately to include with your lab report or email it to your lab partners or instructor. Please note that at the end of each chapter there is a Journal Format. You should insure that your jour-nal contains the exact components listed.

In this experiment we will be using a pulse oximeter attached to the iWorx 214 and the LabScribe software to test the hypothesis that exercise increases pulse rate in a human volunteer. In the process you will learn how to use the LabScribe recording software and the iWorx data acquisition system. Equipment Needed iWorx 214 and computer with LabScribe installed. Pulse Oximeter

Preparation and Using the Software If necessary, turn on the iWorx 214 console using the power button on the rear panel. On the computer, start the LabScribe software by double clicking on the icon located on the desktop. After the software is loaded, select Load Group from the Settings menu. When the dialog box appears, select 4 Intro to iWorx and then click Load or Enter. Click on the Settings menu again and select the Introduction to iWorx Recording set-tings file. Figure 2 shows the resulting screen. In this exercise, the output from a pulse oximeter will be used as a signal source which we will use to determine a volunteer’s pulse rate and the relative rate of blood flow through his or her finger/thumb before and after exer-cise. We are testing the very simple hypothesis that exercise increases pulse rate and blood flow. What do you think will happen? Recorder Procedure: 1. Place your volunteer’s finger

tip into the pulse oximeter according to the diagram on the device.

2. Mark your recording before you start by typing a description in the blank space next to the “mark” button then click on Mark. [NOTE: before you can add any marks to your screen you first have to rec-ord at least one second of data. Quickly hit Record then stop before trying to add marks.] This will insert a mark on the screen wherever the left most cursor is found. It will also be entered into an “index” which will allow you to move to any mark in your data as you proceed.

3. Click Record to record the finger pulse while at complete rest.

4. Click Autoscale in the top channel (labeled Pulse) title area and see the rhythmic signal almost fill the channel recording area. Your data should look like the example shown in Figure 3. If the signal is upside-down you can right click with the mouse on the window and invert the trace. If the base-line in the blood flow window isn’t level you should stop recording and then start it again. Re-peat this process until you obtain a level baseline. The software uses an algorithm to smooth the base-line whenever you hit Record. There are times when this algorithm doesn’t work very well and you need to restart it.

5. After obtaining a good recording, click Stop to halt recording.

6. To determine the heart rate, position the left cursor


Learning To Use LabScribe-Recorder


and post-exercise screens (2 each) should be in the journal. 13. To save your recording, click on the File menu and select Save As. Choose a descriptive file name and save your record-ing to the desktop. Don’t forget to also save your journal. 14. Ask the other lab groups what their results were. How did they compare to your results? Please note: When you proceed to the next exercise you need to use a different file name or you will lose your data for this exercise! You can continue using the same Microsoft Word document for your journal.

Now that you are a little more familiar with the Lab-Scribe software and what it can do, in this next exercise you will be recording some characteristics of stimulus/response relationships. The stimulus will be an artifi-cially applied potential change which will change the membrane potential of receptor cells in your tongue. If the applied depolarization is above threshold for those receptors, you will feel the stimulus as a slight electric shock. The threshold voltage is characteristic of the


on the peak of one of the pulse peaks in the top channel. Position the right cursor on the fifth peak following the first one you chose (see Figure 4).

7. To Copy this screen to your journal switch to your Microsoft Word journal file. Enter your name and other relevant information to the top of the journal page. To insert your data, press the print screen key on your keyboard while in Labscribe then switch to the word document and hit Ctrl-V to paste it into the Word docu-ment.

8. The time for these 5 pulse cycles is listed as T2-T1 in the upper right hand of the computer screen right next to the Record button (see Figure 4). Divide 300 by this number to get the pulse rate (see Figure 5).

9. The height of the peaks (V2-V1) in the blood flow window represents the blood flow through the finger (ml/min). Your results may be a negative number —this is due to the arbitrary positioning of the cursors when you measure the trough to peak height. This isn’t a problem; just take the absolute value of the measured blood flow. We will be using this procedure later in the course (see Figure 6). Copy this screen to your journal as well.

9. Position the left cursor at the trough (bottom) of the first full blood flow (lower window) peak and the right cursor at the peak (top) of the same peak (see Figure 6 on the next page).

10. Copy the screen to the journal. 11. Record the blood flow in the journal. 12. Repeat the above procedure after exercising for 3

minutes. Make sure that you add a mark before you hit record every time! Both the pre-exercise


Figure 3. Pulse and Blood Flow Data

Learning To Use LabScribe-Stimulus/Response

Figure 4. Determining Pulse Rate

the menu. Using the Stimulator To bring up the stimulator panel (Figure 7) click on the View menu then on Stimulator Panel. To Vary Voltage (strength of stimulus) The range of voltages which can be applied are 0 to 5 volts which is adjusted using the Amp (short for amplitude—not amps) win-dow in the stimulator panel (Figure 7) in the upper left hand corner of your computer screen. Note: HP is the holding potential and should always be left at 0 for our purpos-es. W(ms) is the pulses width and should be at 5 or 10 ms.

To vary frequency of stimulation

The frequency of stimulation can be adjusted in a similar manner up to 100 per second using the F(Hz) (frequency

in stimuli per second or hertz) window. To change the number of stimuli The number of stimuli you apply can be adjust-ed by changing the #pulses from 1 up to 9999. A 0 in this box forces the instrument to apply continuous stimuli. Note: Whenever you change any stimulator parameter you need to click on Apply before hitting start. When you click the “Record” button the stimulator will automatically apply the stimulus you have defined. If you click on Apply after you hit record, the stimulator will apply a new set of stimuli every time you click on the button. Setup:

The stimulating electrode should be connected to the red and green terminals of the red stimulator panel on the iWorx 214. On the LabScribe software screen, click on Settings and choose Introduction to iWorx Stimulus. You should now see two screens, the lower recording screen is titled Stimulus while the upper is titled Re-sponse.

cells which are being stimu-lated and how readily the applied potential change penetrates into the tissue. In addition, the frequency of stimulation can also affect the threshold voltage be-cause as you apply more stimuli in a short period of time the stimuli will sum-mate. Thus, the threshold for a single stimulus should be higher than the threshold voltage when using 50 stim-uli per second. This means that many subthreshold stimuli can add together to produce an above threshold potential change in the tar-get cells. This is referred to as summation of subthreshold stimuli. The iWorx 214 apparatus includes a built-in stimulator which we will use in this exercise. We are testing the hypothesis that increasing frequency of stimu-

lation will re-duce the threshold voltage needed to detect a stimulus applied to the human tongue. Set up the Software Click on Settings, then Load Group, and Introduc-tion to iWorx Stimulus. Click on Settings again then on Introduction to iWorx Stimulus at the bottom of


Calculating Pulse Rate

We are using 5 pulses to determine the pulse rate. If we divide the time for 5 pulses by 5 this will give us the time between each pulse. If we then divide 60 by that number we will be calcu-lating the number of pulses in 60 seconds. These calculations can be simplified as shown below:

60

Time for 5 pulses5

60

Time for 5 pulses5

=

60 X5

Time for 5 pulses60 X

5

Time for 5 pulses =

300

Time for 5 pulses

300

Time for 5 pulsesPulses/minute

Figure 5. Calculating Rates

Figure 7. Stimulator Panel

Figure 6. Determining Blood Flow


lus frequency (summation of subthreshold stimuli) on threshold, repeat steps 3 to 7 in the above exer-cise after increasing the frequency to 50.

2. Copy these data into the journal. Skin response. The skin of your forearm has a thick cornified layer of dead cells on its surface and is superfi-cially dry. Do you think that these characteris-tics might alter the ability of an artificially applied volt-age to stimulate the nerve cells found underneath the epithelium?

1. At the maximal voltage (5 V) see if your volunteer can detect the stimulus on his/her forearm. 2. If your volunteer could detect a skin re-sponse place this recording in the journal.

Save your journal.

Find the event marker (Figure 8) and, if necessary, in-sert the DIN cable into the Channel 4 connector of the IWorx 214 unit. Whenever the human volunteer press-es the event button, this will produce a square wave deflection on the top screen (Response) while the stim-ulus will appear in the bottom screen (Stimulus; see figure 9). The stimulus is automatically displayed as a spike indicating every time the stimulator applies cur-rent to the electrodes.

Stimulus/Response Procedure: 1. Locate the stimulating electrode and gen-tly clean it with an alcohol swab. 2. To provide the volunteer with

some idea of the feeling of an above threshold stimulus, set the Amp to 5 volts.

3. Locate the event marker (Figure 8) and give it to your volunteer to hold in his or her left hand.

4. Gently place the stimulating electrodes on the surface of the volunteer’s tongue.

5. Add a mark indicating the stimulus pa-rameters as shown in the small box above. Click on Mark.

6. Click on Record. The volunteer should readily feel the resulting stimulus and press the event marker button accordingly. Click on Stop.

7. To determine the threshold voltage for the same response, reset the voltage to 0.1 V (Amp, make sure you click on Apply) and press Record again. Whenever the volunteer feels the electrical stimu-lus they should press the button firmly and hold it down as long as they feel the stimulus then release it. After the five stimuli are applied click on Stop. If the volunteer cannot feel the stimulus they should not press the button.

8. Increase the voltage in 0.1 or 0.2 V units, click on Apply, enter the stimulus voltage as a Mark then click on Record again. Repeat this process until the volunteer can feel the stimulus.

9. The magnitude of the weakest stimulus the volun-teer can feel represents the threshold for a single stimulus.

10. Copy a recording of the threshold voltage data into the journal. Save your file using a different name than you used for the previous exercise.

Effects of increased stimulus frequency on threshold (summation). 1. To examine the effects of increased stimu-


Figure 8. Event Marker

Amp=5.0 V W(ms)=5 F(Hz)=1 #pulses=5

Figure 9. Stimulus Response

Stimulus

Response

Amp=Varies W(ms)=5 F(Hz)=1 #pulses=0



1) What was your volunteer’s pulse rate (pulses per minute) before and after exercise as determined using the pulse oximeter?

_Before:___________________________________ _After:_________________________________________

2) How does the pulse rate that you recorded relate to that person’s heart rate? ___________________________________________________________________________________________

3) What was blood flow through your volunteer’s finger before and after exercise? _Before:___________________________________ _After:_________________________________________

4) If the blood flow changed, what do you think the physiological significance of that change is? ___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

5) If you change the display time in the main window, will this change the pulse rate that you measured? (Try it) Explain your answer.

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

6) What effect did exercise have on the pulse rate and blood flow of your volunteer? Did your results support the hypothesis mentioned in the exercise?

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________



Learning to use LabScribe - Recorder

Journal: __________ Worksheet Total: ___________


7) Enter the tongue threshold voltages for the class’s volunteers in the table below. Calculate the class average thresholds at 1 and 50 stimuli per second (Hz).

8) You should have seen a difference between the threshold voltage at 1 stimulus per second compared to 50 stimu-

li per second. Why should the threshold voltage change with a change in stimulus frequency? Did your data support the hypothesis stated in the exercise?

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

9) You should see that there was a delay between the administration of a stimulus to the tongue and the volunteer’s response to that stimulus. What caused this delay?

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

10) Did your volunteer feel the stimulus on his or her forearm? Explain these results. ___________________________________________________________________________________________

Laboratory #4 Worksheet (cont.) Name: _______________



Learning to use LabScribe—Stimulus/Response

Tongue Threshold Voltages Class Data

Student: F(Hz) = 1 F(Hz) = 50

#1

#2

#3

#4

#5

#6

Averages:


11) Explain the process whereby an electrical stimulus can induce a receptor cell to generate action potentials. ___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________




Note: Make sure that you hand in your journal for all exercises with this worksheet.


Your Name Lab Partner’s names Learning to Use Labscribe Laboratory Section Date A. RECORDER PROCEDURE Pulse rate and blood flow before exercise State how the pulse rate was determined. Time for 5 recorded pulses = ______________ 300/time for 5 recorded pulses = ______________ = the pulse rate in pulses/minute State the blood flow. Blood flow = ______________ ml/minute Pulse rate and blood flow after exercise State how the pulse rate was determined. Time for 5 recorded pulses = ______________ 300/time for 5 recorded pulses = ______________ = the pulse rate in pulses/minute State the blood flow. Blood flow = ______________ ml/minute



Journal Format for Introduction to iWorx (Guide for producing a complete journal)

Paste the screen showing the pulse rate measurement of the volunteer from

your lab group.

Paste the screen showing the blood flow measurements of the volunteer

from your lab group.

Paste the screen showing the pulse rate measurement of the volunteer from

your lab group.

Paste the screen showing the blood flow measurements of the volunteer

from your lab group.

Every Journal should include every compo-nent listed in this journal format guide. Hand the complete journal in with your worksheet!


B. STIMULUS/RESPONSE DATA Tongue threshold voltage State the threshold voltage . Threshold voltage (Amp) = _________________ volts Frequency of stimuli (should be 1) = _______________ Hz (stimuli/second) Tongue threshold voltage with increased frequency of stimulation State the threshold voltage. Threshold voltage (Amp) = _________________ volts Frequency of stimuli (should be 50) = _______________ Hz (stimuli/second) Skin response State the voltage . Amp = 5V State the frequency of stimulation F(Hz) = ___________________

Make sure all 7 journal pages are in order and turn them in with the worksheet.



Journal Format for Introduction to iWorx (continued) (guide for setting up your journal)

Paste the screen showing the sub thresh-old and threshold voltages and response

of your volunteer.

Paste the screen showing the stimulus and response of your volunteer (only if

there is a response).

Paste the screen showing the threshold voltage with increased frequency of stimulation and the response of your

volunteer.



Laboratory #5 Physiology of Skeletal Muscle

1. To stimulate and observe skeletal muscle contrac-tion.

2. To understand the properties of a twitch, summa-tion of contraction and recruitment.

In the human body, as in the bodies of frogs and other vertebrate animals, skeletal muscle function is directed by the somatic division of the nervous system. Direct nervous stimulation is normally required for a skeletal muscle to contract; in the absence of such stimulation skeletal muscles remain in a relaxed state. In this la-boratory, you will examine how such a muscle re-sponds to stimulation. More specifically, you will focus on how dif-ferent patterns of stimulation result in different types of muscle activity. Remember that the generation of an action potential in an excitable tissue like skeletal muscle first re-quires that the membrane be brought to thresh-old. In skeletal muscle this occurs normally at the neuromuscular junction under the influence of ace-tylcholine re-leased from the alpha motor neuron. In today’s laboratory we will be bypassing the neural stimulation by directly applying a membrane depolarization to the myofibers using the same stimulators we used in the previous

laboratory. If our depolarization is above the threshold value for the myofibers then the muscle will experi-ence an action potential and the result will be a muscle contraction. The gastrocnemius muscle of a frog will be used as your test muscle. Dissection of the muscle from the leg of the animal damages its normal source of stimu-lation — the sciatic nerve. Direct stimulation of the muscle with the stimulator allows us to control various aspects of the stimulation received by the muscle. Remember that you can vary the intensity (strength) of the stimulus delivered to the muscle by adjusting the voltage; you can also vary the frequency of stimu-lation by adjusting the number of stimuli delivered per second. By altering the strength of the stimulus you will be changing the number of muscle fibers involved in the resulting contraction. In the intact animal recruitment

of motor units is utilized to vary the strength of the muscle contraction. We will be using different volt-ages to mimic this process in the isolated muscle. Changes in stimulus frequency can also alter the strength of contraction. However, in the intact ani-mal, contraction is induced by a chain of stimuli which are above the fusion frequency for the mus-cle. The fusion frequency is the frequency of stimu-lation which produces a completely smooth, tetanic contraction of the muscle. It results from the sum-mation of overlapping muscle contractions and the accumulation of calcium within the sarcomeres. We can determine the fusion frequency of the frog gastrocnemius muscle by varying the frequency of stimulation which we apply.

Figure 1 illustrates how the gastrocnemius muscle

will be positioned within the experimental equipment as you assemble it for today’s lab. You will prepare the muscle such that it remains attached to the frog’s femur bone. The femur will be firmly held in place by a “femur clamp”. This will stabilize the origin of the muscle. The insertion of the muscle will be attached

Objectives

Background

Biology 335 Human Physiology: Frog Muscle

Preparation

Figure 1. Gastrocnemius muscle prepared for recording.


to the motion transducer (Figure 2) via a thread tied to the Achilles (calcaneal) tendon. After appro-priate adjustments, contraction of the muscle will result in move-ment of the transducer rod. Move-ment of the rod will produce a permanent record on your comput-er that will provide you with a information about how the muscle has responded to different patterns of stimulation. Please note that in this exercise the mass of the rod is a preload

which acts to pull the cytoskeletal elements within the sarcomeres into alignment prior to contraction. In intact organisms, muscle tone provides a pre-load which serves to pull the cytoskeletal elements in the sarco-meres of our muscles into alignment. This is important because it decreases the time it takes for our muscles to respond to stimuli (latency period). If the force generated by a muscle contraction equals the preload , the muscle will shorten and lift the load (mass actually moved). This type of contraction is referred to as an isoton-ic contraction because the forced needed to move the load is constant during the contraction. If the force generated by a muscle contraction is NOT enough to lift the load then the muscle does not shorten. Since the length of the muscle stays the same it is referred to as an isometric contraction. In this laboratory, we are recording isotonic skeletal muscle contractions. A. Dissection of the muscle The anesthetized frogs will be killed by the instructor in a manner that has been approved by the RIC Institu-tional Animal Care and Use Committee. Each group of students will receive one leg from which the gas-

trocnemius muscle will be dissected. 1. Grasp the cut end of the femur bone with forceps

and pull the skin off the leg. 2. Identify the femur, tibio-fibula (the two bones are

fused in the frog) and the gastrocnemius muscle with its Achilles tendon.

3. Cut the quadriceps and hamstring muscles away from the femur.

4. Insert a glass probe (see Figure 3) between the gastrocnemius muscle and the tibio-fibula and free it from the connective tissue by sliding the glass rod up and down along the bone.

5. Tightly tie a piece of doubled thread approximate-ly 30 cm long to the Achilles tendon.

6. Cut the Achilles tendon from the tibio-fibula dis-tal to the knot. 7. Cut the tibio-fibula just below the knee (see Figure 4). 8. Position the femur in the femur clamp as shown in Figure 1. Place slight tension on the muscle by moving the femur clamp up until you can see that you are pull-ing up on the transducer. 9. Insert the two stimu-lator wires through the muscle by firmly holding the muscle with one hand

while stabbing the wire through the muscle with the other. The wires should be po-sitioned as shown in Figure 1.

It is important to keep the muscle moist during the dissection and throughout the experimental proce-dures. Saline solution is provided in squeeze bottles for this purpose. This fluid must be used instead of water, as its osmolality has been adjusted to prevent osmotic damage to the muscle tissue. You should also note that without an intact circulatory system or mech-anism for replenishing nutrients and removing wastes, the isolated gastrocnemius preparation we are using has a limited period of usefulness.

At this point you should consult your instructor to be sure

that your apparatus is adjusted to provide

optimal recordings of muscle activity.


Gastronemius

Tibia/Fibula Femur

Achilles Tendon

Figure 3. The Frog Leg

Femur

Figure 4.

Figure 2. Motion Transducer


You should be thoroughly versed in all aspects of the following exercises before beginning, as the muscle will probably not last longer than about 30 minutes.B. iWorx Setup After turning on the iWorx unit and starting up the LabScribe software, click on Settings, Load Group and select 5 Muscle Physiology. Click on Settings again then choose 5 Muscle Contraction from the drop down list. This will reveal a window containing two channels. The top channel (Muscle Contraction) will be used to record the muscle contractions while the bottom channel (Stimulus) is reserved for record-ing the stimulator output .

A. Determination of Threshold Remember that when you determine any threshold, you should start with very low inten-sity stimulation and work up to higher intensi-ties. You will be using the iWorx stimulator

panel as illustrated in Figure 5. Remember that if this panel is not visible, click on View and then Stimula-tor panel. Start with the amplitude set at 0.1 Volts, W = 10, F(Hz)=1 and #pulses=5. 1. Whenever you change the settings in the stimula-

tor control panel hit the Apply button before hit-ting Record or adding any marks.

2. Click on Record then Stop to establish a rec-ord so you can begin adding marks or edit your journal. 3. Add a mark to the recording indicating the voltage.

4. One partner should watch the muscle while another partner clicks on Record. As soon as you hit Rec-ord, 5 stimuli will be applied, however you can apply additional stimuli by clicking on the Apply button.

5. After a few stimuli click on Stop. 6. Increase the voltage by changing the Amp= to 0.2,

click on Apply then repeat steps 3 through 5. 7. Repeat steps 3-5, increasing the Amp each time

until you can observe the muscle twitch in response to the stimulus. (See Figure 6).

It is important for someone to watch for the muscle’s response and insure that you are recording properly. 8. When the muscle finally responds, stop recording

then click on the double mountain display time icon until voltages lower than threshold and the threshold voltage data are both visible in one screen. Copy these data to the journal. Remember to put your heading in your journal (as described in the “Introduction to iWorx” laboratory).

B. Summation of Sub-threshold Stimuli In this exercise you will determine whether the threshold voltage is different if the frequency of stimulation is increased. Just as excitatory post synaptic potentials (EPSPs) can summate at a neuron’s axon hillock to bring a neuron to threshold and generate an action potential, below threshold (subthreshold) stimuli (artificially applied or natural) can summate within the motor end plate of a skeletal muscle fiber. Thus, many subthreshold stimuli might summate within our isolated gastrocnemius muscle effectively decreasing the voltage needed to stimulate muscle con-traction and decreasing the threshold voltage. Set the F(Hz) at 50 stimuli per second and #pulses=10, and repeat the determination of threshold (steps A1-A7), starting again from the lowest possible voltage. Record the data just as before and paste them into your journal. C. Effects of Increasing Stimulus Intensity (above threshold) In this exercise you will be stimulating the mus-cle using increasing voltages to recruit more


Figure 5. Stimulator




Exercises


Figure 6. Threshold stimulus

Subthreshold voltage Threshold voltage

Notice that the subthreshold stimuli do NOT result in muscle contraction, while the threshold stimuli clearly produce muscle contractions.

myofibers into the contraction. 1. Return the frequency (F(Hz)) setting on the stimu-

lator to 1 stimulus per second, set the Amp to the threshold voltage (determined in exercise A) and set #pulses=5.

2. Add a mark indicating the starting voltage. 3. Turn the stimulator and recorder on (click Record

in the LabScribe software). 4. After a few contractions, click the Stop button. 5. Increase the voltage in 0.2 volt increments and re-

peat steps 2 through 4 until the magnitude of the contractions no longer increases. Remember to click on Apply each time you change the voltage and before you add your mark or hit Record.

The lowest voltage which can produce the largest mag-nitude contraction is called the maximal stimulus (see Figure 7). Adjust the display time so that you can clear-ly see the effects of increasing voltage and the maximal stimulus and copy this to the journal.

D. Effects of Increasing Stimulus Frequency As the frequency of stimulation increases you will observe incomplete summation, treppe and tetanus (complete summation). The following exercises are designed to illustrate these pro-cesses. 1. Hold the intensity of stimulation constant at the

maximal voltage (if your muscle is already exhibit-ing signs of fatigue you might have to increase the voltage to a level higher than the maximal stimu-lus). Set the frequency back to 1 per second but now set the #pulses to 0 to force the stimulator to apply constant stimuli at the defined amplitude and frequency.

2. Click on Record in the software and you should see the muscle twitch in response to the stimuli. Click on Stop.

3. Increase the frequency to 2 and repeat step 2. 4. Repeat these steps increasing the frequency each

time until you obtain complete summation. Please remember to mark your recording for each fre-quency.

As the frequency was increased to somewhere between 3-6 stimuli per second you should have seen the muscle contract in a pulsatile manner where a new contraction is initiated before the previous contraction has fully re-laxed. This is incomplete summation and should pro-duce a staircase-like pattern of contraction on the record-ing which can be referred to as treppe (Figure 8). As you continued to increase the frequency of stimulation you will eventually reach the fusion frequency. At this point you will observe a smooth, complete contraction. At this point your muscle is experiencing complete sum-mation (tetanus). E. Fatigue Muscle contraction is a very complex cellular event which depends on the ionic and pH envi-ronment of the myofibers as well as a ready supply of ATP to support cross bridge cycling. Fatigue occurs progressively as the ionic and pH envi-ronment is disrupted, and the supply of ATP is depleted. Without a blood supply which can replenish ATP stores and maintain the ion and pH levels in the muscle tissue, the isolated gastrocnemius muscle is more susceptible to fatigue. In this exercise we are going to maximally stimulate the isolated muscle and record the decreasing contraction strength that occurs as the muscle fatigues.


Amp=Part C W(ms)=10 F(Hz)=Varies #pulses=0

Amp=Part C W(ms)=10 F(Hz)=50 #pulses=0

Figure 7. Maximal stimulus


Figure 8. Treppe and Tetanus

Treppe

Tetanus

Maximal stimulus

Notice that the peak height does not in-crease when you increase the voltage above the maximal stimulus.

Since this will effectively “use up” your isolated mus-cle, make sure that you have good data for all other ex-ercises before trying this exercise. 1. Set the stimulator to administer the maximal stimu-

lus at a frequency of 50 stimuli/sec with #pulses set to 0 or more. #pulses=0 provides continuous stimu-lation.

2. Click on Record. 3. Continue stimulating the muscle at a maximum

frequency until you see evidence of fatigue (Figure 9). Once you observe the contraction strength in the Response Window decreasing, click on Stop, adjust the display time appropriately and copy this information to the journal.

F. Properties of a Twitch Although muscle twitches are not exam-ples of normal physiologically meaningful skeletal muscle contraction, they do pro-

vide some insight into the actual workings of skeletal muscles. For each above-threshold stim-ulus, the skeletal muscle will exhibit a brief delay before it exhibits any shortening (contraction). This latency period is the time lag between the onset of the stimulus and the onset of the result-ing muscle contraction. This lag time is due to the “loose” nature of the muscle tissue requiring some initial muscle contraction before any short-ening of the muscle is evident, as well as the time required for the excitatory stimuli to induce mus-cle contraction. Once the muscle begins contract-ing, the contraction occurs fairly rapidly because of the active cross bridge cycling induced by your stimulus. This rapid period of contraction is

called the contraction period and is measured as the time from the onset of contraction to the peak of the contrac-tion. The relaxation of the muscle takes a longer period of time because it is due to the passive recoil of the se-ries elastic elements within the muscle. This relaxation period is measured from the peak of the contraction to the point when the muscle has returned to its baseline length. Using data that you generated early in the laboratory (Part C will work well), bracket a few good twitches with the double cursors in the LabScribe software. Us-ing the cursors measure the latency period, contraction time, relaxation time (as shown in Figure 10) and total twitch time (C+R). Repeat the measurements for 5 dif-ferent twitches and place this data in the table in the worksheet. Calculate the mean latency period, contrac-tion time, relaxation time and total twitch time for your muscle. Copy a representative example of each of these screens into your journal (see Figure 10). Make sure that you save your data file and print or email copies of the journal for each lab member be-fore leaving the laboratory. All calculations and measurements should be clearly indicated in the journal!


Figure 9. Fatigue

Use twitches recorded in Part C.


Figure 10. Properties of a twitch

Latent period

Contraction period

Relaxation period


1) What was the threshold voltage needed to stimulate contraction of the frog’s gastrocnemius muscle when the frequency was set at 1 stimulus per second? ____________

2) What was the threshold voltage necessary when the frequency was set at 50 stimuli per second? ____________

3) Did the threshold voltage change when you increased the stimulus frequency? How and why? ___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

4) Describe the effect of increasing the intensity (voltage) of the stimulus while keeping the frequency constant? ___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

5) Define maximal stimulus.

___________________________________________________________________________________________

___________________________________________________________________________________________

6) What is a motor unit?

___________________________________________________________________________________________

___________________________________________________________________________________________

7) Describe the mechanism by which intact human skeletal muscles exhibit graded contractions. ___________________________________________________________________________________________

___________________________________________________________________________________________

8) Are motor units involved in producing your data with the isolated gastrocnemius muscle? Explain. ___________________________________________________________________________________________

___________________________________________________________________________________________


Laboratory #5 Worksheet Name: Laboratory #5 Worksheet Name: Date: Section:




9) Explain how summation of sub-threshold stimuli works at the membrane/ion level. ___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

10) Define a muscle twitch:

_____________________________________________

_____________________________________________

_____________________________________________

_____________________________________________

11) What is happening inside the myofiber during the la-tency period?

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

12) What is happening inside the sarcomere during the contraction period?

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

13) What is happening inside the sarcomere during the relaxation Period?

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

14) Why does summation of contraction occur? (Do not confuse this with summation of subthreshold stimuli!) ___________________________________________________________________________________________

___________________________________________________________________________________________

Laboratory #5 Worksheet (cont) Name: _______________


Laboratory #5 Worksheet Name: Laboratory #5 Worksheet Name:

(continued)

Data Table 1 Properties of a Twitch

Twitch Ex-ample

Latency Period

1

2

3

4

5

Mean

Contrac-tion Peri-

od

Relaxation Period

Total Twitch Time

15) What was the fusion frequency for your isolated gastrocnemius muscle? What is the ionic basis for the fu-sion frequency? Fusion Frequency: _____________ ___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

16) Why does muscle fatigue occur?

___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

17) Since sarcomeres within skeletal muscles are rigidly aligned with each other what do you think excessive stretch or compression (remember the basic structure of the sarcomere with overlapping thin and thick filaments) will do to the force generation of a muscle contraction? ___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________

19) How does preload in an isolated muscle preparation relate to muscle tone in an intact organism? ___________________________________________________________________________________________

___________________________________________________________________________________________

___________________________________________________________________________________________



Laboratory #5 Worksheet Name: Laboratory #5 Worksheet Name:

(continued)


Your Name Lab Partner’s names Physiology of Skeletal Muscle Laboratory Section Date A. DETERMINATION OF THRESHOLD Threshold voltage at 1 Hz = ______________ B. SUMMATION OF SUB-THRESHOLD STIMULI Threshold voltage at 50 Hz = ______________ C. EFFECTS OF INCREASING VOLTAGE (ABOVE THRESHOLD) Maximal stimulus voltage = ______________ D. EFFECTS OF INCREASING STIMULUS FREQUENCY Fusion frequency = _____________



Journal Format for Physiology of Skeletal Muscle (Guide for producing a complete journal)

Paste the screen showing the threshold voltage and resulting recorded muscle

contraction with F(Hz)=1

Paste the screen showing the threshold voltage and resulting recorded muscle

contraction with F(Hz)=50

Paste the screen showing before and after maximal stimulus was reached.

Paste the screen showing treppe (summation of contraction) and teta-

nus.



E. FATIGUE What voltage and frequency did you use to induce fatigue? ______________ F. PROPERTIES OF A TWITCH Latency period = ______________ Contraction time = ______________ Relaxation time = _____________




Journal Format for Physiology of Skeletal Muscle (continued) (guide for setting up your journal)

Paste the screen showing fatigue

Paste the screen showing the measure-ment of latency period

Paste the screen showing the measure-ment of contraction time.

Paste the screen showing the measure-ment of relaxation time.



Laboratory #6 Skeletal Muscle Physiology: Computer Simulation

The following exercises will explore some basic prop-erties of skeletal muscle contraction using a computer simulation.

Skeletal muscles are composed of hundreds to thou-sands of individual cells, each doing their share of work in the production of force. As their name sug-gests, skeletal muscles move the skeleton. Skeletal muscles are remarkable machines; while allowing us the manual dexterity to create magnificent works of art, they are also capable of generating the brute force needed to lift a 45 kg (~100 lb) sack of concrete. When a skeletal muscle from an experimental animal is electrically stimulated, it behaves in the same way as a stimulated muscle in the intact body, that is, in vivo. Hence, such an experiment gives us valuable insight into muscle behavior. A contracting skeletal muscle will produce force and/or shortening when nervous or electrical stimulation is applied. Unlike single cells or motor units, which follow the all-or-none law of muscle physiology, a whole muscle responds to stimuli with a graded re-sponse. A motor unit consists of a motor neuron and all the muscle cells it innervates. Hence, activation of the neuron innervating a single motor unit will cause all muscle cells in that unit to fire simultaneously in an all-or-none fashion. The graded contractile response of a whole muscle reflects the number of motor units firing at a given time. Strong muscle contraction im-plies that many motor units are activated and each unit has maximally contracted. Weak contraction means that few motor units are active; however, the activated units are maximally contracted. By increasing the number of motor units firing, we can produce a steady increase in muscle force, a process called recruitment or motor unit summation.

Regardless of the number of motor units activated, a single contraction of skeletal muscle is called a muscle

twitch. A tracing of a muscle twitch is divided into three phases: latency, contraction, and relaxation. The latency phase (or latency period) is a short period be-tween the time of stimulation and the beginning of contraction. Although no force is generated during this interval, chemical changes occur intracellularly in preparation for contraction (excitation contraction coupling). During contraction, the myofilaments are sliding past each other and the muscle shortens. Relax-ation takes place when contraction has ended and the muscle returns to its normal resting state and length.

1. Insert the PhysioEx 9.0 CD-ROM into the CD-




4. Once the PhysioEx 9.0 windows opens click on “Exercise 2: Skeletal Muscle Physiology”.

5. Beginning with the Overview, complete the Ac-tivities. At the end of each activity you are given the option of saving your work in a .pdf file. Do so, and submit to your instructor. Save the files with unique file name such as:

Hallsec03pex-02-01 Hallsec03pex-02-02 Hallsec03pex-02-03 Etc.


Objectives

Background

Starting the Program

Biology 335 Human Physiology: Muscle Simulation



Laboratory #7 Reflexes

1. To observe electromyography activity associated with muscle contraction.

2. To observe the Achilles tendon and patellar reflex-es and measure the conduction velocity of the re-flex arc.

3. To observe the effect of conscious motor activity (Jendrassik’s maneuver) on the reflex arc.

Skeletal muscles have specialized receptors which con-vey information about muscle length, tension, and pres-sure to the central nervous system. The sensory recep-tors responsible for providing information about the length, or the rate of change of the length, of a muscle

are called muscle spindles (containing spindle fibers or intrafusal muscle fibers, see figure 1). Arranged in par-allel with the contractile muscle fibers (extrafusal mus-cle fibers), the spindles are stretched when the muscle is stretched by an external force. Therefore, these recep-tors play a significant role in reflexes and maintaining muscle tone. Muscle spindles contain a small bundle of intrafusal muscle fibers which do not contribute to muscle shortening or force production, but regulate the excitability of the sensory afferent spindle nerves by mechanically

stretching the receptors. These fibers are innervated by gamma motor neurons. The majority of a muscle consists of extrafusal muscle fibers, which are inner-vated by alpha motor neurons and are responsible for muscle shortening and production of muscle tension.

When a muscle is stretched, excitation of its spindle fibers causes a reflexive contraction of the muscle (see figure 2). This reflex response is known as a stretch (myotactic) reflex. The minimal delay between the muscle stretching and the reflex contraction is due to its monosynaptic pathway. The sensory afferent nerves from the spindles synapse directly with alpha motor neurons:(there are no interneurons in the path-way). This pathway is the simplest possible reflex arc. As an example of the stretch reflex, consider the reflex response that occurs when a person jumps from a low stool to the floor. The extensor muscles of the legs are stretched on landing, lengthening all their muscle spin-dles. The discharge of the muscle spindles is con-veyed to the central nervous system through the fast-conducting alpha afferent axons. These sensory axons enter the spinal cord through the dorsal root and syn-

apse with the motor neurons of the same extensor muscle. In turn, the motor neurons trigger the contraction of the extensor muscle to oppose the stretch

Objectives

Background

Biology 335 Human Physiology: Reflexes

This laboratory exercise is modified from: iWorx Physiology Laboratory Manual, Exercise HN-2.

Figure 1. Muscle Spindle

Figure 2. Myotactic reflex


produced by landing, completing the reflex arc. This reflex is one of the main reasons you keep your balance and do not fall when changing certain body positions. You will be recording electromyograms (EMGs), the summation of asynchronous electrical activity (muscle action potentials) in the multiple fibers in the muscle, and use them to determine the time between the stretch of the tendon and the arrival of the motor impulse at the muscle.

A. Electrode Placement 1. Use an alcohol prep pad to clean and abrade

three regions on the calf of the left leg for elec-trode attachment. One area is near the ankle, the second is on the skin near the center of the gastrocnemius muscle (calf) and the third is about 6 cm below the back of the knee. Let the areas dry.

2. Place a disposable electrode onto each of the areas.

3. Attach the red (+1) lead to the electrode near the back of the knee.

4. Attach the black (-1) lead to the electrode in the middle of the calf.

5. Attach the green (C ) lead to the electrode near the an-kle.

B. iWorx Setup After turning on the iWorx unit and starting up the Lab-Scribe software, click on Settings, Load Group and select 7 Reflexes. Click on Settings again then choose Reflexes from the drop down list. This will reveal a window containing two channels. The top channel (EMG) will be used to record the EMG activity while the bottom channel (Tendon Tap) is reserved for re-cording the tap on the tendon.

A. Achilles Tendon Reflex A volunteer in each lab group should sit with their leg swinging freely off the ground (you can raise the stool if you need too). The Achilles tendon connects the gas-trocnemius muscle to the tarsal bone of the foot. When you tap on this tendon it will stretch the gastrocnemius muscle and activate the spindle fibers of that muscle. The reflexive response to such stretch produces contrac-tion of the gastrocnemius and a downward movement of the foot (plantar flexion). Click on “Record” and have the volunteer rapidly flex his foot once, pause and repeat. You should now be recording the EMG activity (Figure 5) associated with the contraction


Exercises

Total Path Length ( mm ) Conduction Velocity ( m/sec) = ( M ean Reflex Time ( msec) - 0.5 msec )

Preparation for Achilles Tendon

Figure 4. Achilles Tendon Reflex

Figure 3. Achilles Tendon reflex electrode placement.


Figure 5. EMG recording

of the gastrocnemius muscle. After clicking on “Stop”, copy this recording (using CTRL-C) to your journal. Mark the recording with the subject’s name and “Achilles Tendon Reflex” then hit “Record”. With the volunteer completely relaxed tap the Achilles tendon with the reflex hammer. Record 5 good examples of the Achilles reflex response. Click “Stop” after the 5th trial. Paste one good example of the reflex recording in your journal. Measure the time between the first peak of the tendon tap and the onset of the EMG spike. Then measure the greatest peak to trough magnitude for the EMG spike for each of the five trials and rec-ord these numbers in the worksheet. Calculate the mean Achilles reflex response time and the mean magnitude. Measure the distance between the black electrode and the L5-S1 level of the spinal cord (on the back at the level of the top of the hips). Remember to double this distance to determine the total path length. Calculate the conduction velocity using the formula in the box at the bottom of the pre-vious page. Record the data in the worksheet (Data Table 1).

B. Patellar Tendon Reflex New electrodes should be placed with the black (-1) lead placed about 12 cm from the knee on the thigh overlying the quadriceps muscle group. The red (+1) lead should be placed about 10 cm above the black lead and

the green (C ) lead should be placed on the side of the knee (this is the ground). Click on “Record” and have the volunteer extend their leg rapidly once. Repeat this a couple of times then click on “Stop”. Copy the screen shot of the EMG recording in your journal. Mark the record appropriately, then click “Record” and with the volunteer relaxed tap the patellar ligament with the reflex hammer. Rec-ord 5 patellar reflex responses then click on “Stop”. Measure the reflex time and magnitude of each trial and place a representative screen-shot of each measurement in your journal. Enter the 5 reflex times, the 5 magnitudes and the means in the worksheet. Measure the distance from the black electrode to the L5-S1 vertebrae (top of hips) and calculate the conduction velocity using the same formula as you did for the Achilles reflex. Record the data in the worksheet (Data Table 2). C. Jendrassik’s Maneuver Repeat the patellar tendon reflex recordings on the

same volunteer while the volunteer is pulling their clenched hands away from each other. This is called Jen-drassik’s maneuver. Repeat the calculations of the mean reflex time,



Figure 8. Jendrassik’s maneuver

Figure 7. Electrode placement for knee jerk reflex

Figure 6. Recording Reflex Time and Magnitude

Tap

Response

magnitude and conduction velocities, enter the data in the worksheet (Data Table 3) and copy repre-sentative recordings into the journal.


1) Measure the reflex time (in msec) and magnitude of your 5 trials, calculate the means. then measure the path length in mm and calculate the conduction velocity using the supplied formula.

2) Based on your data and the representative vertebrate conduction velocities shown in the above table, are the nerves involved in your Achilles Tendon Reflex myelinated? Why is this adaptive for the organism? _________________________________________________________________________________________ _________________________________________________________________________________________ _________________________________________________________________________________________ 3) Describe the neural pathway involved in the Achilles tendon reflex (see the diagram on the previous page). ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________



Achilles Tendon Reflex


Trial Time (msec) Magnitude (V)

1

2

3

4

5

Mean:

Path Length (mm):

Data Table 1 Achilles Tendon Reflex

Conduction Velocity (m/sec)

Total Path Length ( mm ) Conduction Velocity ( m/sec) = ( M ean Reflex Time ( msec) - 0.5 msec )

Representative Vertebrate Conduction Velocities

Unmyelinated 1.2 m/sec

Myelinated 45 m/sec


4) Why is this reflex protective and what is it protecting? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________

5) Enter the reflex times, magnitudes, means, path lengths and conduction velocity in the table below :

6) Describe the neural pathway involved in the Patellar tendon reflex. ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 7) The reflex times for the Achilles Tendon Reflex and the Patellar Tendon Reflex should be different? Why? ___________________________________________________________________________________________ ___________________________________________________________________________________________ 8) Compare the conduction velocities of the two reflexes. What factors might explain your observations? ___________________________________________________________________________________________ ___________________________________________________________________________________________

Laboratory #7 Worksheet Name: (Continued)


Patellar Tendon Reflex


1

2

3

4

5

Mean:

Path Length (mm):

Data Table 2 Patellar Tendon Reflex



9) Do you think the patellar reflex would be inhibited or enhanced by actively contracting the quadriceps muscle group? Speculate on the mechanism of inhibition or enhancement? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________

10) Enter the reflex times, magnitudes, means, path length and conduction velocities for the Patellar Tendon Reflex when utilizing Jendrassik’s maneuver:

11) Enter your Mean reflex data in the summary table below:



Jendrassik’s Maneuver


1

2

3

4

5

Mean:

Path Length (mm):

Data Table 3 Patellar Tendon Reflex with Jendrassik’s Maneuver


Patellar Reflex without Jendrassik’s Maneuver

Patellar Reflex with Jen-drassik’s Maneuver

Mean Reflex Time (msec)


Magnitude (V)

Achilles Tendon Reflex


12) Is the patellar reflex altered during Jendrassik’s maneuver? If so, How and why do you think this might hap-pen? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 13) When describing the reflex arcs and the neural pathways involved we often limit ourselves to the nerves car-rying information from the muscle or tendon to the spinal cord and back. However, it should be clear from our ability to feel the tendon tap as well as the results from the Jendrassik’s maneuver test that it isn’t this simple. What other neural connections must be present? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________




Your Name Lab Partner’s names Physiology of Skeletal Muscle Laboratory Section Date A. ACHILLE’S TENDON REFLEX B. PATELLAR TENDON REFLEX C. JENDRASSIK’S MANEUVER



Journal Format for Reflexes (Guide for producing a complete journal)

Paste the screen of a representative myogram when bending the ankle

Paste a screen of a representative myo-gram showing the measurement of

reflex time when tapping the patellar tendon.

Paste a screen showing the measure-ment of reflex time while performing

Jendrassik’s maneuver .


Paste a screen of a representative myo-gram showing the measurement of

reflex time when tapping the Achille’s tendon.

Paste the screen of a representative myogram when bending the knee.

Paste a screen of a representative myo-gram showing measurement of the

magnitude when tapping the Achille’s tendon.

Paste a screen of a representative myo-gram showing measurement of the

magnitude when tapping the patellar tendon.

Paste a screen showing the measure-ment of reflex magnitude while per-

forming Jendrassik’s maneuver .



Biology 335 Human Physiology: Vertebrate Heart

1. To simulate the activity of the autonomic nervous system by applying neurotransmitters directly to an exposed heart.

2. To examine the effects of temperature on cardiac contraction.

3. To examine the refractory period of myocardial contraction.

4. To artificially induce heart block in an exposed frog heart.

The heart of a frog looks very much like that of a small mammal’s. A major internal difference exists, however, in that the frog’s heart has only three cham-bers (Figure 1) while a mammal’s heart possesses four. In mammals, the si-noatrial (SA) node, which is located in the wall of the right atrium, acts as the natural pacemaker. This is the structure that initiates each heart beat. In the frog’s heart, a portion of the sinus venosus plays a similar role. Action potentials that originate within the pacemaker (SA node or sinus venosus) travel via gap junctions through adjacent myocardial cells. The gap junctions are part of the intercalated disks found at the junctions between myocardial cells. Specialized myocardial cells which provide preferential pathways for the propagation of action potentials through these gap junctions form the conduction pathways in the heart. The cells of the conduction pathway are all capable of generating pacemaker potentials and responding to autonomic nervous system stimulation. However, in a normal heart, the SA node (or sinus venosus in the frog) is the pacemaker because the cells in this region have the highest rate of action potential generation. Cardiac function in vertebrates is regulated by the autonomic portion of the nervous system. The heart

Objectives receives input from both sympathetic (cardiac plex-us) and parasympathetic (Vagus nerve) tracts. At any given time, the heart rate and strength of contrac-tion of heart muscle are influenced by the balance that exists between these two sources of excitatory and inhibitory innervation. In this laboratory, after you record the frog’s normal heart action, you will alter the excitatory and inhibito-ry balance by applying various neurotransmitters and receptor blockers directly onto the heart. Depending on the substance that is applied, you will be simulating activation of the sympathetic system or activation of the parasympathetic system.

The neurotransmitters and recep-tor blockers which will be used in this laboratory can alter heart rate by changing the rate at which the pacemaker of the heart (SA node in mammals or sinus venosus in frog) generates action potentials. The normal rate of action poten-tial generation is determined by the rate of spontaneous depolari-zation which occurs between ac-tion potentials within the auto-rhythmic cells found within the

pacemakers. The alternating depolariza-tion and hyperpolarizations are called

pacemaker potentials (Figure 2). The spontaneous depolarization exhibited by autorhythmic cells is caused by a slow calcium current, the rate of which can be modulated by altering the potassium or sodium permeability of the cells. Delaying the closure of po-

Background


Figure 2. Pacemaker Potentials

Figure 1. Amphibian Heart

Laboratory #8 Cardiology with a Vertebrate Heart


tassium channels following an action potential will hy-perpolarize the cell and lead to a slower rate of depolariza-tion between action potentials and a slower heart rate. This is how acetyl-choline and the para-sympathetic nervous system cause a de-crease in heart rate. Conversely, norepi-nephrine and the sympathetic nervous system induce the opening of hyperpolarizing cyclic nucleotide (HCN) channels which increase the sodium current, causing the rate of depolarization to increase, thus increasing heart rate Action potentials (APs) originating within the SA node (or sinus venosus) are propagated throughout the heart via the conduction pathways mentioned previ-ously. At the junction between the atria and the ven-tricle there is a specialized collection of such cells called the atrioventricular (AV) node. Propagation of APs through the AV node is delayed. This time delay is called the AV nodal delay and insures that atrial contraction proceeds prior to ventricular contraction. From the AV node APs are propagated through the Bundle of His, the bundle branches (left and right in human) and finally through Purkinje fibers (cardiac conduction cells) to each of the myocardial cells of the heart.

Instructor A few minutes before the beginning of the laboratory, the instructor will anesthetize the frogs by immersing them in tricaine methane-sulfonate. This anesthetic is readily absorbed through the skin of frogs. After the anes-thetic takes effect, the frog’s brain will be destroyed. From this time forward the animals are “brain dead” and cannot feel pain.

Student When you receive the frog, you will note that the animal’s spinal reflexes may still be in-tact. As part of the central nervous system, the spinal cord alone is capable of integrating sensory (afferent) input and directing “appropriate” muscular responses. The clas-sic example of such a reflex in humans is snatching one’s hand from a hot stove. An-other is contraction of the quadriceps muscles when the patellar ligament is stretched by

tapping it with a rubber mallet. These involun-tary responses are innate and not learned; they

happen very quickly without involvement of the brain. There is awareness of the situation only after the response has occurred, as more time is required for neural input to reach the brain and be integrat-ed by higher brain centers. If the brain is non-functional, as it is in a these frogs, spinal reflexes may continue but the animal cannot be aware of them. Innate reflexes for a frog include retraction of the legs when they are touched. These muscular responses may be evident as you secure the animal to the “frog board”. Place the frog dorsal side down on the board. Make an incision through the abdominal and thoracic walls

Preparation

Figure 3. Frog Heart Preparation

Figure 4. The Force Transducer

Figure 5. Acetylcholine’s effect on cardiac contraction.



with scissors. You must cut through the skin and un-derlying musculature. You also need to cut through the sternum to expose the thoracic cavity. Cut through the ribs on either side of the sternum and remove the sternum. The location of the heart should be readily apparent, since it should be beating within its pericar-dial sac. Carefully cut through the parietal pericardi-um to expose the heart. BE CAREFUL NOT TO CUT ANY MAJOR BLOOD VESSELS! If there is already some blood in the body cavity, use a pipette or paper towels to remove it so that you can see what you are doing. Ringer’s solution may be used to rinse the heart and surrounding structures if necessary. Pin the fully extended forelegs firmly to the board (if neces-sary). Identify the ventricle and the two atria (see Figure 1). Take a bent pin or small fish hook and insert it through the apex of the heart being careful NOT to penetrate the lumen of the ventricle. Tie a 30 cm length of thread to the end of this hook. Tie the other end of the thread to the force transducer (see Figure 4). Adjust the tension on the thread so that it is taut. The heart should be slightly elevated out of the thorac-ic cavity. Tension can be adjusted by moving the transducer up or down on the ring stand.

iWorx Setup Start the LabScribe software and choose 8 Cardiology with a Vertebrate Heart and Vertebrate Heart from the Settings menu. This will reveal a large window labeled Frog Heart. Click Start in the LabScribe software and make sure that you are getting a reasona-bly good record of the heart action on the computer. You will probably have to click on the Autoscale but-ton to get a decent recording. Stop recording when everything is set up properly. Switch to the journal and add your heading. Remember that there is a guide to your journal format at the end of this chapter and please remember to drag the left margin of your jour-nal so that it fills one-half of the computer screen be-fore you copy any screen images into it. Normal Cardiac Contraction Record the action of the heart by clicking on Record. Remember to add a mark to the data (maybe some-thing like “normal contractions”). Stop collecting data after about 10 seconds. Click on Autoscale and deter-


mine the frog’s heart rate the same way you determined the pulse rate in the Introduction to iWorx laboratory (see the box above) and measure the magnitude of a normal contraction (measure the peak to trough height in volts). Place a copy of each of these measurement screens into your journal.

Effects of Cold and Warm on Cardiac Function A dropper bottle of saline solution cooled in an ice bath will be used to apply cold temperature to the heart.

Add an appropriate mark to the recording then begin recording the heart contractions and apply 5-10 drops of cold Saline solution to the heart. Measure the contrac-tion rate and contractility (magnitude) and record each of these screens in the journal. Did cold saline cause any changes in heart rate or contractility? Repeat the above procedure (after rinsing the heart with room temperature saline) using Saline warmed in a wa-ter bath. Measure the contraction rate and contractility (magnitude) and record each of these screens in the jour-nal. Did warm saline cause any changes in heart rate or contractility? Effects of Neurotransmitters and Receptor Blockers Solutions containing epinephrine, acetylcholine and at-ropine will be prepared by the instructor and shared by all members of the class.

Exercises Note: If you have properly marked your recording you can easily move from exercise to exercise to com-pare the data by clicking on the “Marks” icon and choosing the cor-rect location from the list.

Calculating Heart Rate

We are using 5 beats to determine the heart rate. If we divide the time for 5 beats by 5 this will give us the time between each beat. If we then divide 60 by that number we will be calculating the number of beats in 60 seconds. These calcu-lations can be simplified as shown below:

=

=

=Beats/minute

60 Time for 5 beats

5

5 60 X

Time for 5 beats

300 Time for 5 beats

By applying electrical stimuli to the heart muscle while it is contracting you should be able to see when the heart is capable of responding to stimuli and when it is refrac-tory. We will be using the iWorx/114 stimulator for this exer-cise. The electrode should be connected to the red por-tion of the iWorx front panel and held in place by a clamp on your ring stand. Gently position the electrode so that the two electrode wires make contact with some region of the heart without impeding the heart’s contrac-tion. You should see a stimulator toolbar above the Channel 1 screen on the computer. If the stimulator pan-el is not visible click on View and then Stimulator Pan-el. Towards the left end of this toolbar is a button la-beled Apply. Make sure that the frequency setting is at 1. Whenever you click on the Apply button you will stimulate the heart one time. This stimulus will appear as a spike in the window labeled Stimulus.

Record a few normal contractions then stimulate the heart using Apply at different times during the contrac-tions. Your goal is to find a period of time during a contraction when the stimulus has no affect on the heart’s contractions. Copy the screen showing this phe-nomenon into the journal. Heart Block Normal conduction of action potentials through the con-duction pathways of the heart are important for the nor-

mal coordination of cardiac contraction. Especially important is the slight time delay built into the AV node (AV nodal delay). In this exercise we are going to tie a ligature (thread) around the heart between the atria and ventricle (atrio-ventricular sulcus, see Figure 6). By changing how tight this ligature is


1. EPINEPHRINE Before applying epinephrine to the heart, record a few heart beats to establish a baseline immediately prior to drug treatment. Mark this “pre-epinephrine”. Then, apply a few drops of the solution directly onto the heart, aiming as best as you can for the sinus venosus (the site of the pacemaker). This substance requires some time to take effect, so wait about 2 minutes be-fore recording the contractions. After recording the contractions for about 10 seconds, compare this record with the “pre-epinephrine” record to determine whether heart rate or force of contraction (measured in volts using the double cursors) has changed. If no change is initially observed, apply additional epinephrine and allow more time for it to take effect. When you have a good response copy the pre-epinephrine and post-epinephrine screens with the heart rate and contractility measurements to your journal (4 screen shots). 2. ACETYLCHOLINE Rinse the heart thoroughly with Saline solution to eliminate any remaining epinephrine. Allow the heart to normalize for about 10 minutes. Get a “pre-acetylcholine” record and then apply a few drops of acetylcholine (ACh) to the right atrium. The effect is usually rather rapid so you should start looking for a response right away. Record the heart contraction for a

couple of minutes at least. Study the record for changes in heart rate and/or strength of contrac-tion. If no change is observed after two minutes, add additional acetylcholine and allow more time for it to take effect. When you have a good response, copy a tracing to

the journal and enter the heart rate and peak heights which you measured. 3. ATROPINE Rinse the heart again, but you do not need to wait 10 minutes before applying a few drops of atropine solution. Your ACh record will serve as an indication of “pre-atropine” heart action. Record the heart’s response to atropine over the course of a couple of minutes. Study the record for changes in heart rate and/or strength of contraction. Refractory Period


Figure 6. Position of Ligature

Place ligature here

Heart Block: Decreased conduction between the AV node and the Bundle of His can disrupt coordination of atrial and ventricular con-traction. First degree heart block is observed as an increase in the time lag between atrial contraction and ventricular contraction. Second degree heart block is seen as an occasional atrial contraction which is not followed by a properly timed ventricular contraction. Third degree heart block is a complete lack of coordination between atrial and ventricular contraction.

Amp=5 V W(ms)=10 F(Hz)=1 #pulses=1

we should be able to alter the conduction through the AV node and mimic a pathological condition called heart block. Take a piece of thread about 30 cm long. Loop the thread around the atria and tie a loose knot. Record a few normal contractions then slowly tighten the knot (making sure that the thread stays in the proper location). Look at your recording. If there is no change you need to tighten the ligature more until the atria and ventricles are contracting independently of each other. This is called 3rd degree heart block. Copy this recording into your journal. Isolated Heart Remove the heart from the frog by cutting through the major blood vessels and any connective tissue remaining around the base of the heart. Place the heart in a Petri dish containing saline solution. Does it continue to beat now that you have eliminated all nervous input to the heart? Is it contracting at the same rate? Finally, separate the atria from the ventricle by cutting through the AV septum with a scalpel or razor blade. Are any of the individual pieces of tissue still contracting? Record your observations and answer the questions in the Laboratory Worksheet. Please remember to print or email a copy of the journal for each lab member before you leave. Include the journal format form and complete journal with your worksheet.




1) Record the heart rates and contraction strength (vertical displacement or V) during room temperature, cold and warm stimulation in Data Table 1.

2) On the graph below, plot the temperature (Normal, Cold and Warm) on the x axis and heart rate and contrac-

tion strength on the y axes. Use a bar graph and make sure that you label the graph appropriately. (Hint: you can plot heart rate using the left axis and contraction strength using the right axis)

Cold and Warm



Treatment Heart Rate (beats/min)

Contraction Strength (V)

Normal

Cold

Warm

Data Table 1 Effects of Temperature



3) Why does temperature alter heart rate and/or contractility? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________

4) Record the heart rate and contraction strength before and after epinephrine, acetylcholine and atropine expo-sure in Data Table 2.

5) Explain the mechanism by which epinephrine increases heart rate. ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 6) Explain how epinephrine increases cardiac contractility. ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________



Neurotransmitters and Receptor Blockers

Condition or Treatment

Heart Rate (beats/min)

Contraction Strength (V)

Pre-epinephrine

Epinephrine

Pre-Acetylcholine

Data Table 2 Effects of Neurotransmitters

Acetylcholine

Atropine


7) How does acetylcholine induce a decrease in heart rate? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 8) What is the mechanism by which acetylcholine causes a decrease in cardiac contractility? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 9) What is atropine? What is its mechanism of action? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 10) Describe the ionic cause of the prolonged cardiac contractile refractory period. ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 11) How is the refractory period of cardiac muscle different than that of skeletal muscle? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________




12) What role does the AV nodal delay play in normal cardiac function? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 13) Did you notice any change in the ventricular heart rate when conduction between the atria and ventricles was

blocked by ligation? Explain this observation. ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 14) What does myogenic mean? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 15) How did the different portions of cardiac tissue respond after being cut away from each other? Explain your

observations. ____________________________________________________________________________________________ ____________________________________________________________________________________________ ____________________________________________________________________________________________ ____________________________________________________________________________________________ ____________________________________________________________________________________________





16) Should epinephrine or acetylcholine alter the rate of contraction of the separated pieces? Explain. ____________________________________________________________________________________________ ____________________________________________________________________________________________ ____________________________________________________________________________________________ ____________________________________________________________________________________________ ____________________________________________________________________________________________





Your Name Lab Partner’s names Cardiology with a Vertebrate Heart Laboratory Section Date A. NORMAL CARDIAC CONTRACTION Time for 5 beats = ______________ seconds Heart rate (300/time for 5 beats) = _______________ beats/minute Contraction Strength = _________ volts B. EFFECTS OF COLD AND WARM ON CARDIAC CONTRACTION Time for 5 beats = ________ seconds Cold heart rate = __________ beats/min Contraction Strength = _________ volts Time for 5 beats = ________ seconds Warm heart rate = __________ beats/min


Biology 335 Human Physiology: Frog Heart

Journal Format for Cardiology with a Vertebrate Heart (Guide for producing a complete journal)

Paste a screen of normal cardiac con-tractions with measurement of heart

rate.

Paste the screen showing the measure-ment of heart rate after cold saline.

Paste the screen showing the measure-ment of contraction strength after cold

saline.

Paste a screen showing the measure-ment of contraction strength.

Paste the screen showing the measure-ment of heart rate after warm saline.

Paste the screen showing the measure-ment of contraction strength after

warm saline.



C. EFFECTS OF NEUROTRANSMITTERS AND RECEPTOR BLOCKERS Epinephrine Time for 5 beats = ______________ seconds Heart rate (300/time for 5 beats) = _______________ beats/minute Contraction Strength = _________ volts Time for 5 beats = ______________ seconds Heart rate (300/time for 5 beats) = _______________ beats/minute Contraction Strength = _________ volts Acetylcholine Time for 5 beats = ________ seconds Heart rate = __________ beats/min Contraction Strength = _________ volts



Journal Format for Cardiology with a Vertebrate Heart (continued) (guide for setting up your journal)

Paste a screen of the measurement of heart rate pre-epinephrine.

Paste the screen showing the measure-ment of heart rate pre-acetylcholine.

Paste the screen showing the measure-ment of contraction strength pre-

acetylcholine.

Paste a screen showing the measure-ment contraction strength pre-

epinephrine

Paste a screen of the measurement of heart rate post-epinephrine.

Paste a screen showing the measure-ment contraction strength post-

epinephrine


PaRevised Fall 2014

Time for 5 beats = ______________ seconds Heart rate (300/time for 5 beats) = _______________ beats/minute Contraction Strength = _________ volts Atropine Time for 5 beats = ______________ seconds Heart rate (300/time for 5 beats) = _______________ beats/minute Contraction Strength = _________ volts D. REFRACTORY PERIOD E. HEART BLOCK




Journal Format for Cardiology with a Vertebrate Heart (continued) (guide for setting up your journal)

Paste the screen showing the measure-ment of heart rate post-atropine.

Paste the screen showing the measure-ment of contraction strength post-

atropine.

Paste a screen of the measurement of heart rate post-acetylcholine.

Paste a screen showing the measure-ment contraction strength post-

acetylcholine

Paste a screen clearly indicating at least one stimulus applied during the

refractory period.

Paste a screen showing disruption of the cardiac cycle due to ligation and

damage to the AV node.

Laboratory #9 Electrical Properties of the Heart

1. To understand the origins of normal and abnormal heart sounds.

2. To take blood pressures and pulse rates. 3. To record electrocardiograms. 4. To examine the effects of exercise on these meas-

urements.

Background In mammals (including humans), the normal cardiac cycle occurs with the rhythmic opening and closure of the four heart valves as a consequence of the heart’s rhythmic contraction. These valves include the right atrioventricular valve (tricuspid), the left atrioventricu-lar valve (bicuspid), the pulmonary semilunar and the aortic semilunar valves. The atrio-ventricular valves are located at the entrances to the ventricles while the semilunar valves are located at the exits. The opening and clo-sure of these valves occurs because of hydro-static pressure differentials which occur within the heart and the great vessels (vena cavae, aorta, pulmonary arteries, pulmonary veins). During cardiac contraction (systole) the blood pres-sure within the left and right ven-tricles will exceed the blood pres-sure within the vena cavae and

Objectives pulmonary veins (and thus within the atria) and the atrioventricular valves will be forced closed. The clo-sure of these valves causes turbulence and vibrations within the blood and the great vessels. These vibra-tions can be heard at the chest wall as the first heart sound (“lubb” or S1). As the contraction ends, the heart will enter a resting period called diastole. As the heart relaxes, the pressure within the ventricles will drop below the arterial pressure in the pulmonary arteries and aorta. When this occurs the semilunar valves will close producing the vibrations which we hear as the second heart sound (“dubb” or S2). We can hear these sounds using a stethoscope. [The following link: http://depts.washington.edu/physdx/heart/demo.html will bring you to a website which provides examples of both normal and abnormal (murmurs) heart sounds.] Procedure We will use the iWorx 214 equip-ment and an electronic stetho-scope to record heart sounds as we listen to them at the base and apex of the heart. Insert the mini DIN plug from CH4 of the iWorx 214 unit into the jack on the side of the ES100 stethoscope (Figure 3). Turn on the iWorx 214 and start the Lab-scribe 2 software. Under Set-tings, Load Group click on 9 Electrical Properties of the Heart then under the Settings menu click on Electrical Properties of the Heart. You should now see one window labeled “Heart Sounds”.

Cardiac Auscultation

Biology 335 Human Physiology: EKG

Korotkoff, Nikolai Sergie-yevich (b 1874), Russian physi-cian. Korotkoff introduced the auscultation method of deter-mining blood pressure in 1905.

According to the American Heart Association, Korotkoff's sounds occur in five phases. Phase 1: faint, clear, tapping sounds. This is the systolic pressure. Phase 2: murmurs or swishing sounds Phase 3: crisper, more intense sounds Phase 4: distinct, abrupt muffle of sound. In children, this is the diastolic pressure; in adults, it reveals hyperkinetic state (increased movement in blood vessels from disease or stren-uous exercise) if it remains throughout deflation. Phase 5: no longer any sound. This is diastolic pressure in adults

Figure 1. Stethoscope and BP Cuff

Figure 2. Stethoscope placement


http://depts.washington.edu/physdx/heart/demo.html�




1. Use the ES100 electronic stethoscope (instructions for the use of the stethoscope are provided) to listen to a partner’s heart sounds at the apex (bottom) and at the base (top) of the heart (see Figure 2) while recording the sounds with the iWorx. Please note that you must hold the stethoscope very steady to avoid noise. Copy screen shots of the recordings at the base and apex into your journal (see Figure 4). Label S1 and S2.

2. As you listen to the heart sounds attempt to dis-criminate between the first and second heart sounds and attempt to determine the difference between systole and diastole.

3. Answer the questions on the worksheet.

Background As the heart contracts it produces pressure waves which travel through your blood vessels. During car-diac contraction (systole) this hydrostatic pressure reaches a high point and during cardiac relaxation (diastole) it reaches a low point. These pressures are referred to as the systolic and diastolic blood pres-sures. They are easily measured using a blood pres-sure cuff and sphygmomanometer. The blood pres-sure cuff is used to occlude the blood vessels in a per-son’s arm while the sphygmomanometer (“sphygmo” = artery) measures the pressure exerted by the cuff. Once the blood flow is completely occluded by the blood pressure cuff, the pressure exerted by the cuff is gradually decreased. When the pressure exerted by the cuff is less than the systolic blood pressure, blood will be forced past the cuff in a pulsatile fashion set-ting up vibrations which can be heard using a stetho-scope placed just downstream from the occluding cuff. This pressure represents the pressure of blood flowing through the arteries of the arm while the heart contracts. As the pressure is allowed to decrease even more, you will eventually hear a “whoosh” and then silence as blood flow is no longer hindered. This oc-curs as blood is allowed to flow freely through the blood vessels. These sounds are called Korotkoff sounds. In this part of the laboratory we will record pulse waves in the finger and correlate them with the korotkoff sounds as recorded using the ES100 stetho-scope. In the second part of this exercise you will determine the effects of exercise upon blood pressure and cardiac output of a volunteer. Recording Pulse and Korotkoff Sounds Procedure In the Labscribe software click on the Edit menu then click on Preferences. This will bring up a screen which determines what you will be recording and the inputs that are being utilized. Make sure that both Ch 2 Pulse and Ch 4 Heart Sounds are checked and that EKG is NOT checked, then click on OK. You should now see two windows. The top window is labeled “Pulse” and the lower window is labeled “Heart Sounds”. 1. Attach a blood pressure cuff to the arm of a vol-

unteer in your lab group, place the pulse oximeter on the thumb of the same hand, and place the bell of the ES100 electronic stethoscope in the ante-


Blood Pressure

Figure 3. iWorx 214 with ECG leads and ES100 electronic stethoscope.

Figure 4. Recording of heart sounds with ES100.


cubital region of that same arm. 2. Make sure the stethoscope is turned on and

plugged into the iWorx 214 unit. 3. Click on Record then increase the pressure in the

cuff until blood flow in the finger is halted (you can see this in the “Pulse” window (see figure 5).

Slowly release the pressure from the blood pressure cuff while listening for the Korotkoff sounds. Your recording should look like Figure 5. Copy a screen shot of your recording in the journal. Label the events that you recorded. Heart Rate and Blood Pressure Before and After Exercise. Procedure 1. Establish a baseline “resting” blood pressure and

heart rate (use the pulse rate taken from the sub-ject’s radial artery) for your volunteer. Record these data in the worksheet.

2. The volunteer should exercise vigorously for 2 minutes (jumping jacks work fine).

3. Immediately following exercise (0 minutes), the volunteer’s blood pressure and pulse should be recorded again.

4. Repeat the measurements one, three, and five minutes later. (Alternate arms between measure-ments)

5. For each measurement, calculate the pulse pres-sure, stroke volume and cardiac output.

Pulse pressure (mm Hg) = systolic BP - diastolic BP Stroke volume (mL) = pulse pressure x

1.7 Cardiac Output (mL/min) = stroke

volume x pulse rate Record these data in your worksheet. Stroke volume and Cardiac output are very important parameters of cardiovas-cular function. Stroke volume refers to the amount of blood pumped by each ventricle with each contraction. Cardiac output represents the total amount of blood pumped by each ventricle in a minute.

Background An electrocardiograph is an instrument that allows an investigator or clinician to obtain a record of electri-cal events that occur dur-ing the cardiac cycle. Several electrodes, placed at different locations on the surface of the body, are used to detect electrical activity that originates within the heart. The recording obtained, an electrocardio-gram (ECG or EKG), represents a plot of the voltage difference measured between any two of these elec-trodes (Y axis) against time (X axis). The specific pair of electrodes being used to produce a recording is re-ferred to as a “Lead”. Lead I records the voltage be-tween the electrodes located on the left arm (LA) and right arm (RA); Lead II records the voltage between the left leg (LL) and right arm (RA); Lead III records the voltage between the left leg (LL) and left arm (LA). These three standard leads (I, II and III) use only two electrodes at a time. Polarity between the two contact points is specified in such a way that the investigator knows which way the pen on the recorder will move relative to a zero potential. For example, with Lead I, when the LA is positive relative to the RA, the pen will


Page 67 Revised Fall 2014 This laboratory procedure was adapted from De-Coursey, R.M. and F. Dolyak, Laboratory Manual of Human Anatomy and Physiology, 3rd ed., New York, McGraw-Hill, 1974.

Figure 5. Pulse and Korotkoff sounds recording during blood pressure measurements.

Electrocardiogram

Figure 6. Standard Limb Leads

deflect above zero. Lead II is defined as LL positive relative to RA, and Lead III is defined as LL positive relative to LA. Stand-ardization among all electrocardi-ographs is the objective in defin-ing polarity. In addition to these standard leads, augmented leads can be used to produce electro-cardiograms. These leads are designated AVR, AVL and AVF. With these leads, three electrodes (LA, RA and LL) are used, and the last letter in each designation defines polarity, For AVR, the right arm (RA) is posi-tive relative to the other two elec-trodes. For AVL the left arm (LA) is positive; for AVF the left leg is positive. These can easily be remembered if you keep in mind that “R” and “L” stand for right and left sides of the body, while "F" can be related to a “foot” which is associated with the left leg. Although an electrode is customarily attached to the right leg, it is not part of any of the standard or aug-mented leads described above. Chest leads are com-monly used in clinical settings and provide a great deal more information about the electrical activity of the heart. In today’s laboratory we will just be using Limb Lead I with the iWorx 214 apparatus. Procedure We will use our iWorx 214 equipment to obtain elec-trocardiograms before and after exercise in order to examine the electrical activity of the heart under differ-ent conditions. Make sure the EKG leads are connected to the iWorx 214 apparatus as shown in Figure 3. Click on Edit then on Preferences and in then check the “EKG” and “Pulse” channels. Make sure that you uncheck the “Heart Sounds” channel. Click on the mode/function of the EKG channel and set it to 0.03-150 Hz. Adjust the windows so that you have a large “EKG” recording area and a smaller screen beneath labeled “Pulse” as shown in Figure 7. This need not be the same subject as used for the blood pressure exercise, but it must be a person able to do the

2 minutes of exercise. The subject should remove his/her watch and any other jewelry that might contact and interfere with the electrode. Attach the electrodes to both wrists and the left ankle of your volunteer lab partner as follows: Red = right wrist Black = left wrist Green = left ankle During recording, the subject must be quietly seated and sit away from the lab bench without moving. This will minimize electrical interference. When the subject is sitting quietly, click the Record button (after placing a mark onto the record) and record for approximately 15 seconds. Copy this recording into the journal (please remember to enter the heading). From these data you can easily measure heart rate, P-R interval and ventricular systolic and diastolic times.

Heart rate: Measure the time it takes for 5 cardiac cycles (see Figure 7). Divide 300 by this number to give you the heart rate in beats/minute.

P-R interval: Using the double cursors, measure the time from the onset of the P wave to the onset of the next Q (or R)


Page 68 Revised Fall 2014 This laboratory procedure was adapted from De-Coursey, R.M. and F. Dolyak, Laboratory Manual of Human Anatomy and Physiology, 3rd ed., New York, McGraw-Hill, 1974.

Figure 7. Measuring Heart Rate

1 2 3 4 5

T2-T1

wave (see figure 8). V-systole: Measure the time from the

peak of one R wave to the peak of the next T wave (see figure 9).

V-diastole: Measure the time from the peak of the T wave to the peak of the next R wave (see figure 10).

Now ask your subject to do jumping jacks or other exercise vigorously for two minutes. You should leave the electrodes in place on the wrists and ankle, but DETACH THE WIRES FROM THE ELEC-TRODES. This allows the subject freedom of move-ment, prevents damage to the apparatus, and allows you to hook it all back up quickly when the exercise is completed. As soon as possible, obtain another rec-ord . Repeat the measurements you performed on the pre-exercise data and copy all of this into the journal. Pulse Delay Place the pulse oximeter on a finger of the volunteer then with your volunteer sitting quietly record the EKG and pulse simultaneously for about 15 seconds. Copy this record into your journal and measure the time de-lay between peak of the QRS complex and the onset of the pressure pulse in the finger (see figure 11). See the Journal Format Form Provided at the End of the Chapter!



This laboratory procedure was adapted from De-Coursey, R.M. and F. Dolyak, Laboratory Manual of Human Anatomy and Physiology, 3rd ed., New York, McGraw-Hill, 1974.

Figure 8. P-R interval

Figure 9. V-systolic time

Figure 10. Diastolic time

Figure 11. Pulse delay


1) Describe the differences you heard between the heart sounds when you listen at the base compared to the apex of the heart.

___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 2) Could you detect any abnormalities in your volunteer’s heart sounds? If so, describe the sounds. ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________

Remember: Pulse pressure = systolic-diastolic BP Stroke volume = pulse pressure X 1.7 Cardiac Output = heart rate X Stroke volume 3) Calculate the cardiac outputs and place the data in the following table:

4) Using the graph on the next page, plot the cardiac output on the y axis and time on the x axis.

Cardiac Auscultation

Blood Pressure

Time Heart Rate (b/min)

Systolic BP (mm Hg)

Diastolic BP (mm Hg)

Resting

0 minutes post-exercise

1 minute post-exercise



Pulse Pressure (mm Hg)

Stroke Volume (ml)

Cardiac Output

(ml/min)






5) What do your data tell you about the effects of exercise on heart rate, systolic BP, diastolic BP, and cardiac output? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 6) What happened to the cardiac output just after exercise and during recovery from exercise? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________

7) Label the P, QRS and T waves on a normal cardiac cycle in your journal. 8) Enter the pre– and post-exercise heart rate and

EKG information in the table. Calculate the dif-ference between the pre- and post-exercise data and the percent difference. Please make sure that your measurement screens are included in the journal as well.

9) How does the subject’s pre-exercise P-R interval

compare to a normal interval of 120-200 msec? __________________________________________ __________________________________________ __________________________________________ ___________________________________________________________________________________________



Electrocardiogram

Parameter Pre-Exercise

Post-Exercise

Difference (Pre-Post Exercise

Heart rate (beats/min)

P-R Interval (msec)

V-systole (msec)

V-diastole (msec)

% Difference (Difference/

Pre-Exercise)*100


10) What does a P-R interval greater than 200 msec mean for a patient? ___________________________________________________________________________________________ ___________________________________________________________________________________________ 11) Why would you expect diastole to be longer than systole for someone with a resting heart rate? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 12) Assuming that your subject’s heart rate increased, this means that each cardiac cycle must be completed in a

shorter period of time. This could be accomplished by shortening systole, diastole, or both. Which phase of the cardiac cycle shortened the most?

________________________________

13) Why do you think that shortening of this part of the cycle does not seriously hinder ventricular filling? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 14) What was the time delay between the QRS complex and the onset of the pressure pulse in your subject’s

finger? ________ msec 15) Do you think this delay would change if you measured the pulse using a toe instead of a finger? Explain. ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________




Your Name Lab Partner’s names Electrical Properties of the Heart Laboratory Section Date CARDIAC AUSCULTATION BLOOD PRESSURE


Biology 335 Human Physiology: ECG

Journal Format for Electrical Properties of the Heart (Guide for producing a complete journal)


Paste a screen showing a recording of heart sounds from the base of the heart.

Label S1 and S2.

Paste a screen showing a recording of heart sounds from the apex of the

heart. Label S1 and S2.

Paste a screen showing a recording of the pulse waves and Korotkoff sounds when measuring blood pressure. Label

the events on the recording.


ELECTROCARDIOGRAM Pre-Exercise EKG Time for 5 beats = ______________ seconds Heart rate (300/time for 5 beats) = _______________ beats per minute Pre-exercise P-R interval = __________ seconds Pre-exercise ventricular systolic time = _________ seconds Pre-exercise ventricular diastolic time = __________ seconds



Journal Format for Electrical Properties of the Heart (Guide for producing a complete journal)

Paste the screen showing the measure-ment of pre-exercise P-R interval.

Paste the screen showing the measure-ment of pre-exercise ventricular systol-

ic time.

Paste the screen showing the measure-ment of pre-exercise ventricular dias-

tolic time.

Paste a screen showing the measure-ment and calculation of heart rate. Clearly mark the P, Q, R, S and T

waves on one of the cycles.

P

Q

R

S

T


Post-exercise EKG Time for 5 beats = ______________ seconds Heart rate (300/time for 5 beats) = _______________ beats per minute Post-exercise P-R interval = __________ seconds Post-exercise ventricular systolic time = _________ seconds Post-exercise ventricular diastolic time = __________ seconds Pulse Delay Pulse delay = _____________ seconds



Journal Format for Electrical Properties of the Heart (guide for setting up your journal)

Paste the screen showing the measure-ment of post-exercise ventricular sys-

tolic time.

Paste the screen showing the measure-ment of post-exercise ventricular dias-

tolic time.

Paste the screen showing the measure-ment of pulse delay.


Paste a screen showing the measure-ment and calculation of post-exercise

heart rate.

Paste the screen showing the measure-ment of post-exercise P-R interval



Laboratory #10 Circulatory Physiology

1. Observe living capillary beds and understand how blood flow through a capillary bed is regulated.

2. Explore the physiological control of blood flow to human skin.

Background The purpose of the first part of this laboratory is to ob-serve microcirculation. Microcirculation refers to blood flow in the smallest blood vessels within the body -- arterioles, capillar-ies, and venules. It is usual-ly very difficult to observe blood flowing in these ves-sels, because most of these networks, centered around capillary beds, are located deep within organs. How-ever, there are a few ana-tomical situations that are particularly conducive to such study -- specifically wing mem-branes of bats and the webs between the toes of frogs. Both of these structures consist of a “sandwich” of two epithelia with a layer of vascular connective tissue between. Because each of these “sandwiches” is so thin, blood flow in the connective tissue can be observed simply by shining light through the wing or web and observ-ing with a microscope. Today you will work with frogs. Following the in-structions below, obtain the best possible view of blood flowing through the small blood vessels in the web of the foot. You should be able to distinguish arterioles, capillaries, and venules. Arterioles have muscular walls and are larger in diameter than capil-laries; blood flow through arterioles often appears

Objectives pulsatile. As blood enters capillaries, it will flow more slowly and in a non-pulsatile manner. The di-ameter of a capillary is just slightly greater than that of a red blood cell, so blood cells pass through capil-laries in single file. The walls of capillaries are very thin. Venules, which receive blood that leaves capil-lary beds, are the most difficult to identify with cer-tainty, but these are larger in diameter than capillaries and blood flow within them is non-pulsatile. Blood normally flows from arterioles to capillaries to ven-ules (two exceptions include the hepatic portal and

hypothalamo-hypophyseal portal systems). Capillary function depends upon blood flow. If more blood is flowing through a capillary bed, more oxygen will be delivered to that tis-sue. Within capillary beds, there is at least one through fare channel (Figure 1) which provides a preferen-tial avenue for blood flow through the tissue. There are also a number of other capillaries penetrating into the tissue whose function is dependent upon the quantity of blood flowing into the capillary bed. Vasocon-

striction or vasodilation of the arterioles or the precapillary sphincters can dramatically alter the quantity of blood flowing through a capillary bed. These alterations in blood flow will alter the number of capillaries which are actually carrying blood at a given time. Changes in blood flow into a capillary bed are thus most easily detected as changes in the number of functioning capillaries. Vasoconstriction is physiologically regulated by neu-ral, endocrine and local metabolic mechanisms. The neural regulation of vasoconstriction involves the sympathetic nervous system stimulation of the vascu-lar smooth muscle of arterioles by norepinephrine released by autonomic motor neurons. The endocrine regulation involves the release of epinephrine from the adrenal medulla in response to activation of the

Microcirculation

Biology 335 Human Physiology: Circulation

Figure 1. Capillary Bed


sympathetic nervous system originating in the medulla oblongata of the brain. Local controls include tempera-ture and the levels carbon dioxide, oxygen, pH, hista-mine and other chemicals within the tissues. In this laboratory, local effects will be examined by bathing the frog foot in warm or cold saline solution. Autonomic nervous system and endocrine effects will be examined by applying epinephrine (parasympathetic inputs do not normally alter vasoconstriction) to the tissue while the effects of local modulators will be examined by applying histamine to the frog foot. The arterioles in the skin express alpha-1 adrener-gic receptors while arterioles in skeletal muscle express beta-2 receptors. The alpha-1 receptors are excitatory while beta-2 receptors are inhibitory. When norepinephrine (or epinephrine) bind to these receptors the simultaneously cause vasoconstriction in the peripheral arterioles and vasodilation of skel-etal muscle arterioles. This means that activation of the sympathetic nervous system initially will cause peripheral vasoconstriction and skeletal muscle vasodilation increasing blood flow to skeletal mus-cle and decreasing blood flow to the skin (and vis-cera).

Exercise causes an immediate increase in sympathetic nervous system activity which is a consequence of sympathetic stimulatory inputs from the primary mo-tor cortex of the brain to the medulla oblongata auto-nomic nervous system control centers. The medulla oblongata also receives input from hypothalamic ther-moregulatory centers. Increases in core body temper-ature are detected by thermoreceptive cells in the hy-pothalamus and can alter the autonomic control cen-ters in the medulla and cause decreased sympathetic output to the peripheral vasculature thus causing vaso-dilation and increased blood flow to the skin. This increased flow is instrumental in increasing radiated heat loss (in conjunction with sweating) and thus cooling of the core body temperature. Procedure The instructor will immobilize the frogs by placing them in a container containing tricaine methane sul-fonate. This substance is an anesthetic that is ab-


Figure 4. Frog Foot

Figure 2. Stereomicroscope

Stage

Lighting controls

Magnification dial

Focusing dial

Power

Figure 3. Wrapped frog on stage


sorbed across the skin. When the frog no longer re-sponds to touch (this requires at least 15 minutes) the instructor will remove it from the anesthetic and wrap it in a paper towel that has been soaked in tap water, leaving its hind feet exposed. This will prevent the animal’s skin from drying out while you are observing the circulation. The circulation within the webbing of the foot will be observed using a Stereomicroscope (Zoom or Dissect-ing microscope) as shown in Figure 2. The frog wrapped in a moistened paper towel should be placed ventral surface down on the stage of the microscope with one foot over the “window” in the base (Figure 3). Spread the toes in such a way that the toes are widely separated, with the webs between the toes be-ing as flat as possible (Figure 4). Using the highest magnification of the dissecting mi-croscope (350X), identify arterioles, capillaries, and venules. Observe blood flow through these vessels and get a “feel” for how differently it flows through each type of vessel. Remember that blood flows from arterioles into capillaries and then into ven-ules. Temperature Effects Drip some warm saline onto the web. Is blood flow faster or slower? Do the arterioles in your field of view dilate or constrict? Do you think the blood flow (volume) is higher or lower? Describe what you see in the space provided in the worksheet. Apply some ice water to the web. What is happening now? Describe what you see in the space provided in the worksheet. Epinephrine and Histamine Allow the circulation to return to normal by allowing the foot to warm up to room temperature (but don’t let it dry out). Then, test the effects of epinephrine and histamine on the microcirculation. First, drip some of the epinephrine solution (available on the front desk) onto the web and record in the space below any chang-es you observe. Be sure to allow enough time for the solution to “soak into” the tissue. After each treat-ment record your observations in the space provid-

ed in the worksheet. Remember that generally, epi-nephrine is considered a vasoconstrictor while hista-mine is a vasodilator.

Background The purpose of this experiment is to determine how exercise affects blood flow to the skin in human sub-jects. Chose as your subject a member of your group who can safely do at least 5 continuous minutes of moderate exercise.

The skin is the largest organ of the human body and contains the largest reservoir of blood in the body. It is also extremely important in regulating body temper-ature (thermoregulation). As a blood reservoir, in-creased metabolism within other tissues will cause a decrease in blood flow to the skin as it is diverted to supply those tissues with oxygen, etc. As a ther-moregulatory organ, increased core body temperature will cause an increase in blood flow to the skin to in-crease radiated and evaporative heat loss. These two functions (blood reservoir and thermoregulation) can be easily observed during exercise. In this exercise we will be measuring the blood flow through a finger/thumb using a pulse oximeter. These devices detect pressure changes in the finger and using the LabScribe software we can automatically display


Exercise and the Skin


the integral of the pressure change which is a relative measurement of the blood flow through the finger (peripheral blood flow). Procedure Place the pulse oximeter on the index finger or thumb of your volunteer. Use the same digit each time a measurement is obtained. In the Settings menu Load 10 Circulation, then under settings choose 10 Circu-lation. One channel is titled Pulse, the second is la-beled % O2 and the other is titled Blood flow. Record the normal resting blood flow of your volunteer (be sure to mark the recording appropriately) and save this to your journal (don’t forget to start with your head-ing). Repeat the measurement for 5 separate pulses, record the data in the data table in the worksheet. Cal-culate the mean blood flow. Send the subject off to exercise for 1 minute. When he or she returns, record the pulse and blood flow as above (5 measurements) and record the data in the data table and copy a screen shot in your journal. Finally, as soon as possible, have the subject do 3 more minutes of moderate vigorous exercise and repeat the measurements. If there are no differences you may have to repeat the measurements after an additional 2 minutes of exercise. Blood flow can then be determined by measuring the trough to peak difference [see below] (V2-V1=ml/min) for each of the time points (0, 1, 3 and possibly 5 minutes of exercise). As always these data need to be copied to the journal and saved. Before you leave you should perform all of the measurements and print a copy of your journal for


1) Describe how blood flows through the microcirculation of a typical tissue. Include brief descriptions of the different blood vessel types which are involved.

_________________________________________________________________________________________ _________________________________________________________________________________________ _________________________________________________________________________________________ _________________________________________________________________________________________ _________________________________________________________________________________________ 2) What observations can you make concerning blood flow through the different types of vessels in the frog foot? _________________________________________________________________________________________ _________________________________________________________________________________________ _________________________________________________________________________________________ 3) What did you observe when you applied warm water to a capillary bed? _________________________________________________________________________________________ _________________________________________________________________________________________ 4) What happened when you applied cold water? ___________________________________________________________________________________________ ___________________________________________________________________________________________ 5) How can you detect vasoconstriction or vasodilation in the frog’s foot using low magnification stereo micro-scopes? _________________________________________________________________________________________ _________________________________________________________________________________________ 6) Which treatments caused vasoconstriction of blood vessels in the frog’s skin? ____________________________________ _______________________________________ 7) Which treatments caused vasodilation of blood vessels in the frog’s skin? ____________________________________ _______________________________________

Microcirculation






8) According to Poiseuille’s Law, vasodilation is associated with an increase in blood flow through a capillary bed. How do your observations support this concept? ________________________________________________________________________________________ ________________________________________________________________________________________ ________________________________________________________________________________________

9) Record your blood flow data in the table provided. 10) On the following page, graph the mean blood flow on the y axis and exercise time on the x axis. 11) Describe the effects that exercise had on the peripheral circulation of your subject at each time point. _____________________________________________ _____________________________________________ _____________________________________________ 12) What is the physiological significance of vasocon-

striction and/or vasodilation of the peripheral circula-tion during exercise? When during exercise might they occur?

________________________________________________________________________________________ ________________________________________________________________________________________ _________________________________________________________________________________________ 13) How does the brain control peripheral vasoconstriction and vasodilation? ________________________________________________________________________________________ ________________________________________________________________________________________ ________________________________________________________________________________________ 14) During exercise the same neurotransmitter can cause vasodilation in one tissue and vasoconstriction in a differ-ent tissue. How does this work? ________________________________________________________________________________________ ________________________________________________________________________________________ _________________________________________________________________________________________

Blood Flow and Exercise

Blood Flow (ml/min)

Before Exercise

1 min of exercise

3 min of exercise

5 min of exercise

1

2

3

4

5

Mean







15) Most immediate physiological adjustments to exercise occur prior to any change in tissue metabolic demand for oxygen or nutrients. This occurs because exercise increases sympathetic nervous system activity. With your knowledge of exercise and the control of alpha motor neurons by the primary motor cortex of the brain, how does increased exercise influence the sympathetic nervous system? ________________________________________________________________________________________ ________________________________________________________________________________________ ________________________________________________________________________________________ ________________________________________________________________________________________ ________________________________________________________________________________________ ________________________________________________________________________________________ ________________________________________________________________________________________




Your Name Lab Partner’s names Circulation Laboratory Section Date Mean blood flow before exercise = _______________________ Mean blood flow after 1 minute of exercise = _______________________ Mean blood flow after 3 minutes of exercise = _______________________ Mean blood flow after 5 minutes of exercise = _______________________

Make sure all 4 journal pages are in order and turn them in with the worksheet



Journal Format for Circulation (Your journal should contain the following components)

Paste a screen showing the measure-ment of pre-exercise blood flow.

Paste a screen showing the measure-ment of blood flow after 1 minute of

exercise.

Paste a screen showing the measure-ment of blood flow after 3 minutes of

exercise.

Paste a screen showing the measure-ment of blood flow after 5 minutes of

exercise.




Laboratory #11 Mechanisms of Breathing

1. Examine the mechanics of ventilation. 2. Measure lung volumes of a human volunteer. 3. Examine the effects of exercise on lung volumes

and respiratory rhythmicity. 4. Explore the mechanisms underlying the regulation

of respiration.

Background Respiration is commonly considered under two head-ings — internal respiration and external respiration. Internal or cellular respiration is concerned with the physical and chemical factors involved in the utiliza-tion of oxygen and formation of carbon dioxide by tissue cells. Mitochondria contain the enzymes that catalyze the chemical reactions of cellular respiration. External respiration includes: 1) Breathing (or ventilation of the lungs): the gase-

ous exchange between an organism and its envi-ronment, which provides for maintenance of an adequate oxygen supply in alveolar air and elimi-nation of carbon dioxide. Breathing requires the action of the diaphragm and other thoracic and abdominal muscles. The coordination of these muscles is regulated and controlled by the nerv-ous system.

2) Exchange of oxygen and carbon dioxide between alveolar air and the blood within lung capillaries.

3) Transport of oxygen and carbon dioxide by the blood between the lungs and metabolizing tissues.

Breathing is characterized by the bulk flow of air into and out of the lungs. This flow is driven by air pres-sure changes within the thoracic cavity which occur because of changes in the volume of the thoracic cavi-ty. Such volume changes are accomplished via the abdominal and thoracic musculature, primarily the diaphragm. You should quickly review an anatomy

Objectives text to re-familiarize yourself with the muscles in-volved in breathing. The changes in thoracic volume (which will alter lung volume) that occur during the ventilation can be esti-mated by measuring the changes in thoracic and ab-dominal diameters and widths during inhalation and exhalation. These changes in thoracic diameter should reflect actual changes in thoracic cavity volume which will cause changes in intra-alveolar pressure and changes in lung volume. Changes in lung volume can be measured using a spi-rometer. A spirometer is a device which measures the volume of air being inhaled or exhaled while a subject breathes. The most important of these volumes are: Tidal volume (TV, see volume a in figure 1)—the

volume of a normal resting breath (normally around 500 mL).

Inspiratory Reserve Volume (IRV, see volume c in figure 1) — the volume which can be inhaled in addition to the normal TV.

Expiratory Reserve Volume (ERV, see volume b in figure 1) — the volume which can be exhaled in addition to the normal TV.

Residual Volume (RV, see volume e in figure 1) — the volume of air left in the lungs after a maxi-mal exhalation.

Vital Capacity (VC, see volume d in figure 1) — the maximum volume of air which can be ex-

Respiration

Figure 1. Lung Volumes

Biology 335 Human Physiology: Breathing


changed in a single breath. Total Lung Capacity (TLC, d+e) = VC + RV The rate at which air can be expelled from the lungs is a useful clinical tool in diagnosis of obstructive pul-monary diseases. Normally, a person should be able to expel between 75% - 85% of his/her vital capacity in the first second of a forced exhalation. This meas-urement is called the FEV1. If airways are obstructed, however, as in asthma, exhalation is hindered and the FEV1 will be lower. A person’s normal respiratory rate is determined by the medullary rhythmicity center in the medulla oblon-gata of the brain. This area of the brain in turn re-ceives input from chemoreceptors in the body which are sensitive to carbon dioxide levels (actually sensi-tive to H+ levels) in the blood. There are chemore-ceptors in the aortic arch and carotid bodies (the pe-ripheral chemoreceptors) as well as chemoreceptors in the medulla (central chemoreceptors). The peripheral chemoreceptors can be influenced by any source of H+

in the blood whether from changes in CO2 level (because CO2 is converted to H+ and bicarbonate by carbonic anhydrase) or changes in acid production (i.e. lactic acid). The central chemoreceptors are only sen-sitive to changes in respiratory CO2 levels because H+ can’t cross the blood brain barrier (CO2 does cross and is converted into H+ and bicarbonate by carbonic an-hydrase). At rest, alterations in ventilation pattern will cause alterations in blood carbon dioxide levels. Thus, increased respiratory rate (hyperventilation) will de-crease blood CO2 while decreased respiratory rate (hypoventilation) will increase blood CO2. Such alter-ations in blood CO2 will, in turn, induce changes in the respiratory rate and depth in order to bring blood CO2 levels back to normal. Although it is true that changing blood carbon dioxide level will alter the respiratory rhythm, it should be noted that exercise induced changes in the respiratory rhythm are NOT due to changes in blood carbon diox-ide level. During exercise the autonomic nervous sys-tem immediately enters a state of sympathetic tone (sympathetic dominance) and the increased activity of the noradrenergic neurons and blood levels of epi-nephrine serve to increase the respiratory rhythm prior to any actual change in blood carbon dioxide level. In today’s laboratory we will be examining each of these physiological concepts: 1) How thoracic and abdominal dimensions change with breathing, 2) Lung volumes before and after exercise and 3) How chang-

ing blood carbon dioxide level can alter the breathing rhythm.

Procedure Using a centimeter measuring tape, determine the fol-lowing dimensions and record them in the table on the worksheet. Circumference of the chest at the level of the 3rd

rib (just under the armpit) during resting and forced inhalation and exhalation (4 measure-ments).

Circumference of the abdomen at the level of the umbilicus during resting and forced inhalation and exhalation (4 measurements).

Using the calipers, determine the following dimen-sions and record them in the table in the worksheet. Anterior-posterior thickness of the chest at the

level of the 3rd rib during resting and forced inha-lation and exhalation (4 measurements). Position the calipers over the shoulder to make these meas-urements.

Side-to-side thickness of the chest at the level of the 3rd rib during resting and forced inhalation and exhalation (4 measurements).

Procedure See figure 2 for spirometer setup, turn on the IWorx unit, then start up the LabScribe software and from the settings menu choose load and 11 Mechanisms of Breathing then Settings again and choose Mecha-nisms of Breathing. The resulting screen should have 2 recording screens labeled Air Flow and Volume. When you breathe through the spirometer the air flow is detected by the iWorx air flow transducer and the

Thoracic and Abdominal Dimensions

Measuring Lung Volumes


Figure 2. iWorx Spirometer


software converts this into air flow in liters. The bottom tracing automatically converts the rate of air flow into a vol-ume. You must al-low the equipment to warm up for at least 10 minutes before beginning the exer-

cises and you should NOT breathe through the ma-chine after commencing recording for about 5 seconds before placing your mouth on the mouthpiece. In addition, always breathe through the spirometer head with the tubing pointing up to avoid the formation of condensation in the tubing. While inhaling you should see the tracing on the computer screen rise. If it de-creases you should breath through the other end of the spirometer. Tidal Volume (TV) Have your subject put on a nose clip (or hold the nose closed with their fingers). About 5 seconds after clicking the Record button have the subject breathe as normally as possible for about 10 cycles (see Figure 3). The subject should NOT be watching the recorder. Copy the resulting screen into your journal after you enter the heading into the journal. Using the lower Volume screen, determine the average tidal vol-

ume (in liters) of 5 respiratory cycles by positioning the left cursor on the trough and the right cursor on the peak of each of 5 different cycles. Record the average of these 5 tidal volumes in the worksheet and your journal. Measure the peak to peak distance (time) for 5 respiratory cycles. Divide 300 by this number to get the res-piratory rate in breaths per minute (the same way you measured heart rate). Rec-ord these data in your journal. From the respiratory rate and the tidal volume you can determine how much air your subject breathes every minute. This is called the respiratory minute volume and is calcu-lated by multiplying the respiratory rate

by the tidal volume. Record these values in the journal and in your worksheet. Expiratory Reserve Volume (ERV) The subject’s nose should be closed off, as previously described. To determine the expiratory reserve vol-ume, which is the additional amount of air that can be expelled beyond normal exhalation, the subject should breathe normally for a few breaths, then exhale maxi-mally, without looking at the computer screen. The expiratory reserve volume (ERV) is measured as the increase in exhaled air volume over and above a nor-mal exhalation (see Figure 4). Repeat this measure-ment 3 times and calculate the average. Record these screens and calculations in your journal and work-sheet. Vital Capacity (VC) The nose should be closed off as before. To determine the vital capacity (the maximum volume of air the subject can exchange), the subject should, after a few normal breaths, inhale as hard as possible, then exhale into the spirometer as deeply as possible while record-ing, without looking at the computer screen (see Fig-


TV

ERV

TV

ERV

Figure 4. Measuring ERV

Figure 3. Main spirometry screen with tidal volumes shown.

Take all measurements from this screen

Do not measure from this screen!

Take all measurements from this screen

Do not measure from this screen!

time

volume

VERY IMPORTANT Do not breathe through the flow head during the first 5 seconds after click-ing on the Record button. The iWorx uses this time to zero the volume re-cording. If your baseline is not level you probably did not allow enough time before breathing into the flow head.


ure 5). The greatest volume reached is a measure of the vital capacity. Repeat this 3 times and calculate the average. Record these data in the journal and work-sheet. Inspiratory Reserve Volume (IRV) From Figure 1, you can see that: VC = TV + ERV + IRV With measurements for VC, TV and ERV, you can solve for IRV and calculate the inspiratory reserve vol-ume of your subject. Record this value in the journal. Total Lung Capacity (TLC) From Figure 1 you can also see that: TLC = VC + RV Assuming that the average human has a residual volume of 1.2 liters, solve this equation for TLC. Record this value on the worksheet.

Have the same subject exercise moderately for 2 minutes. Immediately have the subject close off his/her nose, and then record his/her breathing in order to com-pare rate and depth with the resting pattern previously recorded. Copy this tracing into the journal. Determine the post exercise TV, respiratory rate and respiratory

minute volume. Enter these data in the worksheet and your journal. In the next two exercises you will examine how blood carbon dioxide level influences the respiratory control centers in the brain. Since we can’t directly alter carbon diox-ide levels we are going to indirect-ly change the levels and examine

the consequences.

Decreasing Blood CO2 (Hyperventilation) Hyperventilation will decrease carbon dioxide levels in the blood. Since carbon dioxide level is a proximal stimulator of breathing, if we hyperventilate for a peri-od of time we will release more CO2 from the blood and we should be able to hold our breath longer than if we didn’t hyperventilate. You can change subjects for this exercise if you wish and you do not need to use the spirometer. First, the subject should hold his/her breath for as long as possible with a lab partner timing him/her. This time should be recorded on the worksheet. Then, the same subject should now hyperventilate by breathing deeply at the rate of 2 breaths/second for about 30 seconds; then the subject should take a deep breath and hold it as long as possible. Record this time on the worksheet. Increasing Blood CO2 (Hypoventilation For this exercise you will again use the spirometer and


Changing Carbon Dioxide Levels

Effects of Exercise

VCVC

Figure 5. Measuring Vital Capacity

The determination of residual volume in humans requires specialized equip-ment and the use of inert radioactive tracers. This determination is beyond the scope of an undergraduate physiol-ogy laboratory. Thus we assume a residual volume of 1.2 liters.


recorder. The same volunteer you used for the spirome-try previously should record at least 5 normal breathing cycles (mark the recording “pre-CO2”). They should then breathe into a paper bag for 3 minutes (or as long as possible). The volunteer should then record their breathing in the same fashion as before (mark the re-cording “post-CO2”). Measure the tidal volume and respiratory rate before and after the volunteer breathed into the paper bag. Copy and record these data into the journal and worksheet.




1) Enter the chest and abdominal measurements in the following table. 2) Do these measurements correspond with your understanding of how changes in thoracic and abdominal cavity dimensions should change during breathing? Please explain. __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ 3) Describe how the diaphragm moves/works during a normal breathing cycle. __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ 4) What happens to intra-alveolar pressure during inhalation and exhalation? __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________

Thoracic and Abdominal Dimensions

Resting

Inhalation Exhalation Inhalation Exhalation

Chest Circumference (cm)

Abdominal Circumference (cm)

Ant. - Post. Chest Dimension (cm)

Side-to-Side Chest Dimension (cm)

Forced





5) How do abdominal muscles contribute to inhalation and exhalation? Explain. __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ 6) During forced breathing, you might expect the abdominal circumference to change dramatically. Why? __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________

Pre-exercise Tidal Volume (TV), Respiratory Rate and Respiratory Minute Volume Respiratory Rate: Time for 5 cycles = ____________ sec Respiratory rate (300/time for 5 cycles) = __________ breaths/min Average tidal volume = __________ Liters Respiratory minute volume (TV X Respiratory rate) = __________ Liters/min

7) Record these data in the summary table on the next page. Pre-exercise Expiratory Reserve Volume (ERV)

8) Enter the ERV value in the summary table on the next page.



Lung Volumes and Exercise

Tidal volume (liters)

Average:

ERV (liters)

Average:


Vital Capacity (VC) 9) Enter the average VC into the summary table. Inspiratory Reserve Volume (IRV) IRV = VC - (TV + ERV) IRV = __________ liters 10) Enter the IRV into the summary table. Total Lung Capacity (TLC) TLC = VC + RV (RV = residual volume = 1.2 liters) TLC = _________ liters 11) Enter the TLC into the summary table. Effects of Exercise on Tidal Volume, Respiratory Rate and Respiratory Minute Volume

Respiratory Rate: Time for 5 respiratory cycles = _________ sec Respiratory rate (300/time for 5 cycles) = __________ breaths/min Average post-exercise TV = __________ Liters Post-exercise Respiratory Minute Volume (TV X Respiratory rate) = __________ Liters/min



Summary Data

Parameter Pre-Exercise Post-Exercise Difference

Tidal volume (L)

Respiratory rate (breaths/min)

Respiratory minute volume (L/min)

Expiratory re-serve volume (L)

Inspiratory reserve volume (L)

Vital Capacity (L)

Total Lung Capacity (L)

VC (liters)

Average:

Post-Exercise Tidal Volumes (liters)

Average:





12) From the summary table it should be evident that respiratory minute volume changes most dramatically with exer-cise. What is the physiological significance of this change? ________________________________________________________________________________________________ ________________________________________________________________________________________________ ________________________________________________________________________________________________ ________________________________________________________________________________________________

Effects of Hyperventilation (decreased blood CO2) 13) Before hyperventilating subject held breath for _________ 14) After hyperventilating subject held breath for _________ 15) Can you think of a sport in which this might be an important (and legal) part of increasing performance? __________________________________________________________________________ Pre Hypoventilation Tidal volume Respiratory rate: Time for 5 respiratory cycles = __________ sec Respiratory rate (300/time for 5 cycles) = __________ breaths/min Average tidal volume = _________ liters Respiratory minute volume = __________ liters/min Post Hypoventilation (increased blood CO2) Tidal Volume Respiratory rate: Time for 5 respiratory cycles = __________ sec Respiratory rate (300/time for 5 cycles) = __________ breaths/min Average tidal volume = _________ liters Respiratory minute volume = __________ liters/min

Changing Carbon Dioxide Levels

Pre Hypoventilation Tidal Volume

(liters)

Average:

Post Hypoventila-tion Tidal Volume

(liters)

Average:





16) What differences in the tidal volume and/or respiratory rate can you see from the beginning to the end of the 3 mi-nute hypoventilation period? __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ 17) What effect did hypoventilation have on respiratory minute volume? Why? __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ 18) How is blood PCO2 detected in the human body? __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ 19) Where are the most important receptors and how do they work? __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________ __________________________________________________________________________________________






Your Name Lab Partner’s names Mechanisms of Breathing Laboratory Section Date A. PRE-EXERCISE TIDAL VOLUME AND RESPIRATORY RATE Pre-exercise average (from the summary table) tidal volume = _________ Liters Pre-exercise respiratory rate = __________ breaths/min B. EXPIRATORY RESERVE VOLUME Average expiratory reserve volume (from summary table) = __________ Liters C. VITAL CAPACITY Average vital capacity (from summary table) = __________ Liters E. EFFECTS OF EXERCISE Average post-exercise tidal volume (from summary table) = __________ Liters Post-exercise respiratory rate = __________ breaths/min



Journal Format for Mechanisms of Breathing (Guide to producing a complete journal)

Paste a screen showing the measure-ment of pre-exercise respiratory rate.

Paste a representative screen showing the measurement of tidal volume.

Paste a representative screen showing the measurement of expiratory reserve

volume. < (exhale)

Paste a representative screen showing the measurement of vital capacity.

(deep inhale) >

< (deep exhale)


Paste a representative screen showing the measurement of post-exercise tidal

volume.

Paste a screen showing the measure-ment of post-exercise respiratory rate.


Make sure all 10 journal pages are in order and turn them in with the worksheet

F. EFFECTS OF HYPOVENTILATION Average pre-hypoventilaion tidal volume = __________ Liters Pre-hypoventilation respiratory rate = __________ breaths/min

Average post-hypoventilaion tidal volume = __________ Liters Post-hypoventilation respiratory rate = __________ breaths/min


Journal Format for Mechanisms of Breathing (continued)

Paste a representative screen showing the measurement of pre-

hypoventilation tidal volume.

Paste a screen showing the measure-ment of pre-hypoventilation respiratory

rate.

Paste a representative screen showing the measurement of post-

hypoventilation tidal volume.

Paste a screen showing the measure-ment of post-hypoventilation respirato-

ry rate.


Laboratory #12 Restrictive and Obstructive Lung Disease

1. Gain an understanding of the importance of air flow during inhalation and exhalation

2. Simulate the effects of restrictive lung disease on air flow during respiration.

3. Simulate the effects of obstructive lung disease on air flow during respiration.

In the last laboratory (Chapter 11), spirometry was used to measure the major lung volumes and capaci-ties during human respiration. In addition, the regula-tion of the respiratory rhythm was explored by manip-ulating blood CO2 levels through hyperventilation or hypoventilation. Human pulmonary diseases are often diagnosed by measuring lung volumes and the rate of air flow through the pulmonary airways. Pulmonary diseases are usually classified as either restrictive or obstruc-tive. Restrictive diseases are those characterized by a decreased ability of the lungs to change volume result-ing in a decrease in the vital capacity and an increase in residual volume. Any disease which decreases lung compliance (emphysema) or decreases the ability of the lung to be inflated (myasthenia gravis) is consid-ered a restrictive disease. Obstructive diseases are characterized by reduced air flow through the pulmo-nary airways (emphysema or asthma). The determination of lung volumes can be carried out using spirometry as in La-boratory #11. These measurements can be used to detect changes in lung infla-tion in the diagnosis of restrictive lung diseases. The determination of changes in airflow, however; requires a different measurement. The rate of air flowing through the pulmonary airways is meas-ured by recording a patient’s FEV1/FVC. FEV1 is the Forced Expiratory Volume in the first second of the exhalation. FVC is the Forced Vital Capacity or the total volume exhaled during the same exhala-tion. Figure1 illustrates how FEV1/FVC is measured

Objectives using the iWorx spirometry system. Forced exhalation is utilized for this measurement because it is more sensitive to airway changes than inhalation. During exhalation the lungs and airways are subjected to a positive pressure which tends to force the airways to partially close. If the airways are already narrower than normal this will result in a measurable decrease in airflow. On the other hand, during inhalation, the lungs and airways are subjected to a negative pressure which forces airways open thus masking abnormal airway issues. The FEV1/FVC is usually expressed as a percentage (simply multiply FEV1/FVC by 100) and normal values are 80% or higher.

Procedure Set up the spirometer as describe in laboratory #11. From the settings menu choose load and 12 Lung Disease then Settings again and choose Lung Disease. The resulting screen should have 2 recording screens labeled Air Flow and Volume. With the computer recorder on, the volunteer should breathe normally for a couple of cycles then inhale maximally, hold their breath momentarily, then exhale maximally AS LONG, FAST AND HARD AS POS-SIBLE (keep that nose closed!). Repeat this for a total

Air Flow During Respiration

Biology 335 Human Physiology: Lung Disease

1 second

FEV1FVC

1 second

FEV1FVC

Figure 1. FEV1/FVC

Normal FEV1/FVC


This laboratory exercise was adapted from a protocol devel-oped by Dr. Debra Mulliken-Kilpatrick of Boston College.

of 3 trials, allowing some recovery time between tri-als. This is very similar to the measurement of VC performed in Laboratory #11, however, if the subject pauses momentarily prior to exhaling it will make the measurement easier (see Figure 1). Using the double cursors (use the arrow keys on the keyboard for precision), measure the TV, IRV and ERV as shown in Figure 2. Then from the beginning of the exhalation, mark off 1 second (T2-T1) and determine the volume exhaled during that 1 second (V2-V1; Forced Expiratory Volume in the first sec-ond[FEV1]). Then move the right hand cursor over to the lowest point of exhalation (Forced Vital Capacity; FVC) and determine the volume (V2-V1). Divide the FEV1 by the FVC then multiply by 100 to express the volume as a percent of the maximal exhalation. This is called the FEV1/FVC. Repeat this measurement 2 more times and enter the data in the table provided in the worksheet. Copy your best tracing into the journal and record the average FEV1/FVC in the worksheet and the journal.

Restrictive pulmonary diseases are characterized by a decrease in lung compliance or ability to expand the thoracic cavity. This being true, we can induce a re-strictive condition by having a subject wear a medical corset which decreases the maximum expansion of the thoracic cavity. A volunteer in your lab group should put the medical corset on tight enough to restrict expansion without being unduly un-comfortable. Now have the volunteer breath through the spirometer as de-scribed in the previ-ous exercise. Rec-ord three normal tidal volumes fol-lowed by a maxi-mum inhalation (pause) then a maxi-mal exhalation (long, hard and forceful). See Fig-ure 2 for an exam-ple. If your output doesn’t appear simi-lar to the example you will need to ask

your instructor for assistance. Repeat the measurements 2 more times with the same volunteer resting briefly between each measurement (3 times total) Record the data in the table in the work-sheet and provide a sample recording in the journal. If you don’t see a change in the FVC you may need to make the corset a little tighter. Note: The corset should be tight but not uncomforta-ble. However, it should be tight enough to restrict the expansion of the chest to some degree.

Obstructive pulmonary diseases are characterized by decreases in air flow through the respiratory tree. We can examine the impact of changes in the pulmonary airways in the laboratory by decreasing the diameter

Restrictive Disease


Obstructive Disease—Decreased Airway Diameter

Lung Volume Volume (ml)

Tidal volume (TV) 500

Inspiratory Reserve Volume (IRV) 3100

Expiratory Reserve Volume (ERV) 1200

Forced Vital Capacity (FVC) 4800

Residual Volume (RV) 1200

Average Adult Lung Volumes

Figure 2. Pulmonary Function Testing


of the tube leading to the spirometer head. Use the narrowed airway attachment on the spirometer head to test this. Measure your volunteers pulmonary function volumes as you did in the previous exercises. Repeat 2 more times for a total of 3 sets of measurements. Enter your data in the table provided in the worksheet and include a representative recording in your journal.



1. Enter the normal FEV1, FVC and FEV1/FVC data in the table provided below. When complete, enter the aver-age TV, IRV, ERV, FVC and FEV1/FVC in the summary table.

2.Why is the measurement of FEV1/FVC used instead of the rate of inhalation in pulmonary function testing? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 3. What affect will a poor seal around the mouthpiece have on the results of an FEV1/FVC measurement? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________

4.Enter the data collected from the subject wearing the medical corset into the table provided here. When com-plete, enter the average TV, IRV, ERV, FVC and FEV1/FVC in the summary table.


Normal FEV1/FVC


FEV1 (Liters) FVC (Liters) FEV1/FVC (%)

ERV (ml)

IRV (ml)

TV (ml)

Averages:

Restricted and Obstructive Pulmonary Diseases


ERV (ml)

IRV (ml)

TV (ml)

Averages:



5. Enter the data collected from the volunteer breathing through the obstructed spirometer. When complete, enter the TV, IRV, ERV, FVC and FEV1/FVC in the summary table. 6. Why is the rate of exhalation a better measurement than the rate of inhalation for the diagnosis of obstructive diseases? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 7. Do your results agree with the expected physiological changes for restrictive and obstructive pulmonary dis-eases? Why or why not? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________



ERV (ml)

IRV (ml)

TV (ml)

Averages:

Summary Table

Measurement Restrictive Obstructive

TV (ml)

IRV (ml)

ERV (ml)

FVC (L)

FEV1/FVC (%)


8. What changes in the FEV1/FVC would be expected with restricted pulmonary disease? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 9. What changes in FEV1/FVC should you see with obstructive disease? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 10. What pulmonary volume should be most altered by restricted lung disease? Does your data support this ex-pectation? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 11. What effect should restrictive disease have on the residual volume? Why? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 12. Why is asthma considered an obstructive disease? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________ 13. Why is epinephrine an effective emergency treatment for an acute asthmatic attack? ___________________________________________________________________________________________ ___________________________________________________________________________________________ ___________________________________________________________________________________________



D. FEV1AND FORCED VITAL CAPACITY Average FEV1/FVC (from summary table) = __________ %

Your Name Lab Partner’s names Mechanisms of Breathing Laboratory Section Date A. NORMAL PULMONARY FUNCTION VOLUMES B. RESTRICTIVE DISEASE C. OBSTRUCTIVE DISEASE — DECREASED AIRWAY DIAMETER


Biology 335 Human Physiology: Lung Diseases

Journal Format for Lung Diseases (Guide to producing a complete journal)

Paste a representative screen showing the measurement of normal pulmonary

function. See Figure 2 for example

Paste a representative screen showing the measurement of pulmonary func-tion while wearing the medical corset.

Paste a representative screen showing the measurement of pulmonary func-tion while breathing narrowed device.




1. To determine the mean metabolic rate of student

volunteers. 2. Compare human metabolic rate to the metabolic

rate of a small mammal.

Since the time of Lavoisier (1743-1794), it has been realized that the body behaves somewhat like a furnace, requiring fuel to “burn” as a source of energy with a consequent elimination of heat. The chemical reactions that occur in the body may be classified as energy trapping (endergonic) or energy releasing (exergonic). The exergonic reactions make available the largest increments of energy for four es-sential processes: (1) energy for muscle movement; (2) energy for digestion and associated processes; (3) ener-gy for adjustment of body temperature; and (4) energy for basal maintenance of all the body cells. This last is basal metabolism, and in order to measure it the first three energy needs must be held to a minimum. Basal metabolic rate (BMR) as determined clinically is the metabolic level of the individual under the following three conditions corresponding to minimal levels for the first three of the above considerations:

1. Mentally relaxed, rested in the recumbent posi-

tion for 30 minutes before the test, and in a re-cumbent position during the test.

2. In a post-absorptive state; that is, the subject has not taken any food for 12 hours prior to the test.

3. At a comfortable environmental temperature — between 20o to 25o C (68o to 77o F) and at a normal body temperature. This is known as the thermal neutral temperature.

There are a number of methods for measuring metabolic rate including heat production which is a waste product of cellular metabolism (see formula below). If you think about this a few minutes you will realize that de-termining total body heat production over time under

Objective

the conditions listed above might be a problem. In-stead, a very handy way to estimate metabolic rate is to determine oxygen consumption (since anaero-bic metabolism contributes very little to total me-tabolism in mammals).

Most body heat is produced by the physiological oxidation of fats, proteins and carbohydrates. These foodstuffs are digested in the gastrointesti-nal tract, and are absorbed into the vascular system (blood and lymph) as fatty acids and glycerol, amino acids, and simple sugars. Each type of

foodstuff, when oxidized, will liberate a given amount of heat for each liter of oxygen consumed. The amount of heat liberated is called the calorific value of a liter of ox-

ygen for that material. The established values are: kcal per L O2 Fat 4.69 Protein 4.48 Carbohydrates 5.00 Typical Mixed Diet 4.83

Although there are specific calorific values for each foodstuff, the average person is on a mixed diet which has been found to have a calorific value very close to 4.83 kcal (large calories) per liter. Thus, for each liter of oxygen consumed, 4.83 large calo-ries of heat are produced.

Once oxygen consumption has been determined, this can be related to the total average number of kilocalories (kcal) consumed for a given amount of oxygen consumed (4.83 kcal/L oxygen). This will give you an equivalent to the rate of heat loss per given period of time. This can be compared to standard population averages for a given body size and expressed as a percentage of that value.

1 kcal (large calorie) is the amount of heat re-quired to warm 1 kilogram of water by 1 degree Cel-sius.

Laboratory #13 Basal Metabolic Rate

Background

Biology 335 Human Physiology: Metabolic Rate


The next figure shows that as animals become larger, the mass-specific BMR actually decreases. The mass-specific BMR is an animal’s basal metabolic rate ex-pressed on a per gram body weight basis. In other words, it is the metabolic rate for each gram of mass of an ani-mal. We are going to determine and compare the average BMR of a group of students and the average BMR of a group of laboratory mice.

Procedure Materials: MedGem Indirect Calorimeter Nose clip Calculator Subject: It will be impossible to measure oxygen consumption under truly basal conditions in our laboratory. There is simply too much going on in the room. However, at least 6 students in each class should come to class without eating or drinking caffeinated beverages. The reason for this is that they will be in a post-absorptive state, satisfy-ing at least one of the conditions under which BMR should be measured. Oxygen consumption of the subjects will be determined using a MedGem Indirect Calorimeter. Each subject will take turns determining his or her oxygen consumption according to the directions supplied by your instructor.


There are several standard ways which this infor-mation can be expressed. The one we shall use is the Mayo Foundation Normal Standard. The standard is expressed as large calories (kcal) of heat produced per square meter of body surface area per hour. In com-puting the metabolic rate of the given individual (which is compared with that of the standard), it is necessary to know his/her body surface area. The body surface area is read off the DuBois Surface Area Chart when height and weight of the subject are known.

The basal metabolic rate (BMR) is expressed as the percentage amount by which the metabolic rate of the individual lies above or below the standard. Thus, if any individual had a metabolic rate of 40 calories per hour per square meter of body surface area and his standard was 30 calories per hour per square meter of body surface area, his metabolic rate would be 40/30 of the standard, or 33.3 percent above the standard. Thus, his BMR would be expressed as +33.3. Why use surface area? It turns out that an animal’s (or a person’s) metabolic rate is affected by body sur-face area. What will lose heat faster, a 10 cm3 steel block or a 1 cm3 steel block? The smaller object will lose heat much faster than the large block. This is because the smaller object actually has a larger sur-face area for its given mass. Generally, smaller ani-mals exhibit higher metabolic rates on a per gram basis. The total heat production for a human is much greater than that for a mouse, but based on a mouse’s body weight they exhibit higher metabolic rates than humans. The relationship between basal metabolic rate and body size for many different animals is illustrated in the next two figures. The first graph shows that as body size increases, heat production per day increas-es.


Log [mass] (kg)

Log[kcal/day]

Mass-specificBMR

mass (g)

Human Basal Metabolic Rate

The MedGem The MedGem is a state-of-the-art, handheld, indirect calorimeter that accurately measures oxygen consumption (VO2). Indirect calorimetry is a process whereby the rate of energy expenditure is estimated based upon the rate of oxygen consumption or car-bon dioxide production.


Calculations: The numbers below refer to specific lines on the la-boratory worksheet. You can enter your numbers on that sheet as you progress through the calculations. 1. Observed volume of oxy-

gen consumed per minute (mL/min). This value will be taken from the indirect calorimeter as directed by the instructor.

2. Volume in liters per mi-nute. Convert the oxygen consumption value on line (1) to liters per minute by dividing by 1000.

3. Heat production per mi-nute in kcal. Multiply the number on line (2) by 4.83. The reason for using this figure is discussed in the “Introduction”. Briefly, you are taking your subject’s oxygen consumption (liters) and multiplying it by 4.83 kcal/liter of oxygen, which is an estimate of how much heat is produced for each liter of oxygen a person consumes. This converts units of oxygen con-sumption into units of energy (kcal).

4. Heat production per hour. Multiply the num-ber on line (3) by 60. This is a very meaningful number because it reflects the rate at which the subject burns “calories” (which are actually kilo-calories) while sitting quietly.

5. Subject’s height. Express this in cm (1 in = 2.54 cm).

6. Subject’s mass. Convert pounds to kilograms by dividing by 2.2.

7. Subject’s mass. Convert kilograms to grams by multiplying by 1000.

8. Subject’s body surface area. Use the following formula to determine the body surface area:

9. Heat production per hour per square meter of body surface. Divide line (4) by line (8). This is one way to express the metabolic rate of your subject. It must now be compared with published standard values.

10. Standard. Refer to the table of Mayo Founda-tion Normal Standards in this handout and find

the standard BMR for a person of the same sex and age as your subject.

11. Basal metabolic rate. BMR can be expressed as a comparison to this published standard. The number on line (9) may be above or below the standard. Ex-press the metabolic rate as a percentage of the stand-ard. Divide line (9) by line (10), then multiply by

100. A percentage greater than 100 means your value was above the standard; a per-centage less than 100 means your value was below the standard. 12. Mass-specific metabolic rate. Another way to express BMR is in units of heat pro-duced per hour per gram body mass. To calculate this, di-vide the number on line (4) by the subject’s mass in grams on line (7). Report this value in the worksheet.

Record your group data in the table on the blackboard, overhead or instructor’s computer as directed by your in-structor. BMR data, surface area and body weights should be recorded for subsequent calculation of class averages. Calculate the class average human surface area to mass ratio by dividing the class average surface area (m2) by the class average mass (g). Record this value in the worksheet.

In this exercise we will determine the average BMR for a group of mice. Procedure Materials: Metabolism chambers Soap solution Balance Barometer Thermometer

THE MAYO FOUNDATION NORMAL STANDARDS Mean Basal Metabolic Rate (Kcal per square meter per hour) Age Males Females 17 44.80 41.45 17.5 44.03 40.74 18 43.25 40.10 18.5 42.70 39.40 19 42.32 38.85 19.5 42.00 38.30 20-21 41.13 37.82 22-23 40.82 37.40 24-27 40.24 36.74 28-29 39.81 36.18 30-34 39.34 35.70 35-39 38.68 34.94 40-44 38.00 33.96


S.A. (m2)

height (cm) X mass (kg)

3600

Mouse Resting Metabolic Rate

Subjects: Adult laboratory mice. As we did for human subjects earlier, we will calculate meta-bolic rates for mice using oxygen consumption data. Each group of students will obtain a mouse, determine its mass, and place it in a metabolism chamber. The metabolism chamber is a cylinder with a large one-hole rubber stopper blocking one end. Soda lime has been place in the bottom of the chamber to absorb carbon di-oxide. To isolate the mouse from the soda lime and to limit its movement during the metabolic rate determination it will be placed in a small mesh cylinder which is sealed at either end with a cap. When a mouse is placed in the mesh cylinder and the metabolism chamber is closed, the carbon dioxide which the mouse generates via cellular respiration will be ab-sorbed by the soda lime. The resulting decrease in volume represents oxygen consumption. The rubber stopper has a graduated tube penetrating it such that if you place a soap bubble at the end of the tube, when the mouse breathes and the carbon dioxide is absorbed, the volume of air in the cylin-der decreases and the soap bubble will be “sucked” in to-wards the mouse. The actual oxygen consumption which you measure needs to be multiplied by a correction factor which adjusts for barometric pressure and altitude such that all measurements are standardized to these conditions. You didn’t have to do this for the human data because the MedGem automatically introduces this correction. 1. Weigh the mouse using the supplied balance. 2. Place the mouse in the metabolic chamber. 3. Allow the mouse to acclimate (get used to) its surround-

ings for at least 5 minutes. 4. Insuring that the chamber is closed, place a soap bubble

over the end of the pipette using a moistened finger tip. 5. Record the oxygen consumption over a given period of

time. Try to record the time it takes for a full 5 mL of oxygen to be consumed. If this proves very difficult rec-ord both the volume (at least 2 mL) and the time and de-termine the oxygen consumption in mL/min based upon these data.

6. Repeat this 5 additional times. Between trials, briefly “flush” the air in the chamber by removing the rubber stopper and allowing room air to enter.

7. Record your data on the laboratory worksheet. 8. Determine the average oxygen consumption of your

mouse based on the six measurements you obtained. 9. Use the oxygen consumption value for your mouse to

calculate the various BMR measurements (see calcula-tions below).


Calculations: 1. Oxygen consumption per minute (mL/min). This val-

ue is the average oxygen consumption of your mouse. 2. Chamber temperature (oC). The temperature in the

metabolic chambers as provided by the instructor. 3. Barometric pressure (mm Hg). Current barometric

pressure as supplied by the instructor. 4. Correction factor (from table or calculated). This fac-

tor standardizes your oxygen consumption for variations in barometric pressure and temperature.

5. Corrected oxygen consumption for the mice. Multiply line 1 by line 4.

6. Oxygen consumption in L/min. Convert to L/min by dividing line 5 by 1000.

7. Heat production (kcal/min). Multiply line 6 by 4.83. 8. Heat production (kcal/hour). Multiply line 7 by 60. 9. Mouse mass (g). 10. Surface area (m2). Use the formula in the box below to

calculate your class average mouse surface area. 11. Heat production (kcal/h/m2). Divide line 8 by line 10. 12. Mass specific BMR (kcal/h/g). Divide line 8 by line 9.

The Mouse Metabolic Chambers The mouse is placed in a tube containing soda lime which absorbs any carbon dioxide the mouse produc-es. By placing a soap bubble at the end of the pi-pette, the chamber becomes a closed space. As the mouse consumes oxygen and exhales carbon diox-ide, the carbon dioxide is absorbed and the volume of gas in the chamber decreases in an amount equiva-lent to the amount of oxygen the mouse consumed. The decreased volume pulls the soap bubble through the pipette. This change in volume represents the oxygen consumption of the mouse.


Formula for estimating rodent surface area: S.A. (cm2) = 0.437+(2.143 X mass(g)) Divide this by 10,000 to give m2

Correction Factor

Baro. P. (mm Hg)

760 X

273

Temp. (oC) + 273


Cautions:

1. Mice may bite. If you are not used to handling mice, allow the instructor to assist you in transferring the mice from their cages to the chamber and back again.

3. Return each mouse to the same cage

from which it was taken. This will minimize disruption of their social order and prevent unnecessary fighting among the animals.


Record your group data in the table on the black-board, overhead or instructor’s computer as di-rected by your instructor. BMR data, surface area and body weights should be recorded for subse-quent calculation of class averages.

Calculate the surface area to mass ratio by divid-ing the class average surface area (m2) by the body weight (g). Record this value in the work-sheet. The surface to mass ratio of a species or organism represents the proportion of that organism’s mass which is exposed to the environment. An animal with a larger surface area to mass ratio will lose heat from their bodies more rapidly. This heat must be replaced by metabolic activity. Smaller animals will exhibit a larger surface area to mass ratio and will thus typically exhibit higher per gram metabolic rates.

Surface/Mass Ratios



1) Enter your group volunteer’s data in the spaces provided below: Subject’s name: ________________________ Subject’s sex: _______ Subject’s age: _______

1- Oxygen consumption (mL/min) ______

2- O2 volume in L/min ______

3- Heat production (kcal/min) ______

4- Heat production per hour (kcal/hr) ______

5- Subject’s height (cm) ______

6- Subject’s mass (kg) ______

7- Subject’s mass (g) ______

8- Surface area (m2) ______

9- Heat production per hour per m2 ______

10- Standard BMR ______

11- BMR (% of standard) ______

12- Mass specific BMR (kcal/h/g) ______

2) Enter the human volunteer data for each group in the table below and calculate the means:


Lab. Group Heat Production (kcal/h)

Mass Specific BMR (kcal/h/g)

1

2

3

4

5

6

Class Average

Class Data — Human Mass (g)

Heat Produc-tion (kcal/h/m2)

Surface Area (m2)

Human Basal Metabolic Rate

THE MAYO FOUNDATION NORMAL STANDARDS Mean Basal Metabolic Rate (kcal per square meter per hour) Age Males Females 17 44.80 41.45 17.5 44.03 40.74 18 43.25 40.10 18.5 42.70 39.40 19 42.32 38.85 19.5 42.00 38.30 20-21 41.13 37.82 22-23 40.82 37.40 24-27 40.24 36.74 28-29 39.81 36.18 30-34 39.34 35.70 35-39 38.68 34.94 40-44 38.00 33.96




3) Calculate the average human surface area to mass ratio using your class data: 4) Under what circumstances might the BMR data you obtained be inaccurate or a poor representation of a basal meta-

bolic rate? ________________________________________________________________________________________________ ________________________________________________________________________________________________ ________________________________________________________________________________________________ ________________________________________________________________________________________________ 5) Explain the connection between oxygen consumption and a person’s metabolic rate. ________________________________________________________________________________________________ ________________________________________________________________________________________________ ________________________________________________________________________________________________ ________________________________________________________________________________________________

6) What hormone is primarily responsible for the regulation of resting metabolic rate? How does it work? ______________________________________________________________________________________________ ______________________________________________________________________________________________ ______________________________________________________________________________________________ ______________________________________________________________________________________________ 7) Exercise involves the generation of action potentials by the alpha motor neurons that control the skeletal muscle mo-tor units in your body. The alpha motor neurons are ultimately controlled by the primary motor cortex of the brain which also has input into autonomic nervous system (ANS) control centers of the brain. Increased activity of the pri-mary motor cortex (exercise) also stimulates the ANS. What hormone/neurotransmitter is released in response to this activity and how does it relate to metabolic rate? ______________________________________________________________________________________________ ______________________________________________________________________________________________ ______________________________________________________________________________________________


Class Average Human Surface Area to Mass Ratio: ________________________________________

Do the math! Surface area di-vided by mass.



8) Enter your group mouse oxygen consumption data in the table below, calculate the average oxygen consumption and com-plete the series of calculations indicated (explained in the manual). Cage Number:______________ Mass of mouse: ______________ grams

9) Complete the class data mouse BMR table below:


Lab. Group Heat Produc-tion (kcal/hr)

Mass of Mouse (g)

Heat Production (kcal/hr/m2)

1

2

3

4

5

6

Class Average

Class Data — Mouse Surface Area

(m2)

Mass Specific BMR (kcal/hr/g)

Time (min):

Starting Volume

(usually 0 mL)

Ending Vol-ume (mL) when the

bubble burst

Change in Volume

(mL)

O2 Consumption (mL/min)

Average:

Mouse BMR Group Data Table

Mouse Basal Metabolic Rate

1- Average O2 consumption (mL/min) ______

2- Chamber Temperature (oC) ______

3- Barometric pressure (mm Hg) ______

4- Correction factor (from table or calculated) ______

5- Corrected O2 consumption (mL/min) ______

6- O2 volume in L/min ______

7- Heat production (kcal/min) ______

8- Heat production per hour (kcal/h) ______

9- Mouse mass (g) ______

10- Surface area (m2) ______

11- Heat production (kcal/h/m2) ______

12- Mass specific BMR (kcal/h/g) ______




10) Using your data, calculate the Surface to Mass ratios for the class average mouse:

11) Rewrite the class average metabolic rate data for the human and mouse in the table below to make it easier to

compare the data: 12) Why is human heat production (kcal/h) greater than mouse heat production (kcal/h)? ______________________________________________________________________________________________ ______________________________________________________________________________________________ ______________________________________________________________________________________________ 13) Why is human mass-specific heat production (kcal/hr/g) less than mouse mass-specific heat production (kcal/h/g)? ______________________________________________________________________________________________ ______________________________________________________________________________________________ ______________________________________________________________________________________________

Mouse Surface Area to Mass Ratio: ________________________________

BMR Measurement Human Mouse

Heat production (kcal/h)

Surface Area-Specific Heat Production (kcal/h/m2)

Mass Specific Heat Produc-tion (kcal/h/g)

Surface area to Mass Ratio

Comparing Mouse and Human BMR Measurements

Do the math! Surface area di-vided by mass.




14) If surface area is the largest contributor explaining the difference between the human and mouse mass-specific

metabolic rates, explain your surface area-specific metabolic rate data. ______________________________________________________________________________________________ ______________________________________________________________________________________________ ______________________________________________________________________________________________ ______________________________________________________________________________________________



Laboratory #14 Renal Physiology— The Function of the Nephron

To understand the three nephron processes responsible for urine formation: 1. Glomerular filtration 2. Reabsorption 3. Secretion

Waste products resulting from the metabolic pro-cessing of nutrients are either recycled by the liver, excreted by the liver into the bile, exhaled, or filtered out of the blood and ex-creted in the urine. The filtration of the blood and production of urine is ac-complished by the kid-neys. Each kidney con-tains a million or so neph-rons that are composed of two major parts: a glomer-ulus and a renal tubule. The glomerulus is a tan-gled capillary knot that filters fluid from the blood into the lumen of the renal tubule. The function of the renal tubule is to pro-cess that fluid, also called the filtrate. The beginning of the renal tubule is an en-larged end called the glomer-ular capsule (Bowman’s capsule), which surrounds the glomerulus and serves to funnel the filtrate into the rest of the renal tubule. Collectively, the glomerulus and the glomerular capsule are called the renal corpus-cle. As the rest of the renal tubule extends from the glo-merular capsule, it becomes twisted and convoluted, then dips sharply down to form a hairpin loop, and then coils again before entering a collecting duct. Starting at the glomerular capsule, the anatomical parts of the renal tubule are as follows: the proximal convoluted tubule, the loop of Henle (including the

Objectives descending and ascending portions), and the distal convoluted tubule. Two arterioles are associated each glomerulus: an afferent arteriole feeds the glo-merular capillary bed and an efferent arteriole drains it. These arterioles are responsible for blood flow through the glomerulus. Constricting the afferent arte-riole lowers the downstream pressure in the glomeru-lus, whereas constricting the efferent arteriole will increase the pressure in the glomerulus. In addition, the diameter of the efferent arteriole is smaller than the diameter of the afferent arteriole, restricting blood flow out of the glomerulus. Consequently, the pressure in the glomerulus forces fluid through the endothelium of the glomerulus into the lumen of the surrounding glomerular capsule. In essence, everything in the

blood except the cells and proteins are filtered through the glomerular wall. From the capsule, the filtrate moves into the rest of the renal tubule for processing. The job of the tubule is to concentrate the urine and reabsorb all the beneficial substances from its lumen while al-lowing the wastes to travel down the tubule for elimi-nation from the body. The nephron performs three important functions to process the filtrate into

urine: glomerular filtration, tubular reabsorption, and

tubular secretion. Glomerular filtration is a passive process in which fluid passes from the lumen of the glomerular capillary into the glomerular capsule of the renal tubule. Tubular reabsorption moves most of the filtrate back into the blood, leaving principally salt water plus the wastes in the lumen of the tubule. Some of the desirable or needed solutes are actively reabsorbed, and others move passively from the lu-men of the tubule into the interstitial spaces. Tubular secretion is essentially the reverse of tubular reab-sorption and is a process by which the kidneys can rid the blood of additional unwanted substances such as creatine and ammonia.

Background

Figure 1. The Nephron and Tubule Cell Types

Biology 335 Human Physiology: Renal


The reabsorbed solutes and water that move into the interstitial space between the nephrons need to be re-turned to the blood, or the kidneys will rapidly swell like balloons. The peritubular capillaries (vasa rec-ta) surrounding the renal tubule reclaim the reab-sorbed substances and return them to general circula-tion. Peritubular capillaries arise from the efferent arteriole exiting the glomerulus and empty into the veins leaving the kidney. Start the PhysioEx program as described in previous laboratories. Choose the Renal System Physiology module. Overview

1. Insert the PhysioEx 9.0 CD-ROM into the CD-




4. Once the PhysioEx 9.0 windows opens click on “Exercise 9: Renal System Physiology”.

5. Beginning with the Overview, complete the Ac-tivities. At the end of each activity you are given the option of saving your work in a .pdf file. Do so, and submit to your instructor. Save the files with unique file name such as:

Hallsec03pex-09-01 Hallsec03pex-09-02 Hallsec03pex-09-03 Etc.


Getting Started

Biology 335 Human Physiology: Renal


Documents

Laboratory Manual Fall 2014