Hematology 7.1
BIOLOGY 3420
ANIMAL PHYSIOLOGY
FALL 2011
LAB #7
HEMATOLOGY OF SHEEP AND CATTLE
Objectives:
As a result of attending this lab, students should be able to:
1) Identify the procedures for determining the hematocrit, the red blood cell count and the
hemoglobin concentration of whole blood.
2) Calculate MCH, MCHC and MCV given the measurements in 1).
3) Comment on how environmental adaptations are reflected in animal hematology.
TERMINOLOGY: erythrocytopenia, polycythemia, anemia, hemoglobin, deoxyhemoglobin,
oxyhemoglobin, carbaminohemoglobin, carboxyhemoglobin, methemoglobin, cyanmethemoglobin,
hemocytometer, mean corpuscular volume, mean corpuscular hemoglobin concentration, mean
corpuscular hemoglobin, oxygen-carrying capacity, hematocrit (corrected and apparent)
INTRODUCTION
Advance reading: Moyes and Schulte pp. 444-446 (2
nd
ed.).
Blood, like any other connective tissue, consists of specialized cells (erythrocytes, leucocytes, and
platelets) suspended within a relatively large amount of intracellular fluid (blood plasma). Blood has
several important functions concerned with maintenance of constant internal environment (homeostasis),
including the transport of respiratory gases, nutrients, metabolic wastes, and hormones; as well as the
regulation of acid-base status, the electrolyte balance, and body temperature. It also participates in
defense of the organism against potentially toxic foreign materials or organisms.
The erythrocytes (red blood cells, RBCs) of humans and other vertebrates contain a respiratory pigment,
hemoglobin (Hb), which imparts the characteristic red color to the cells. Hemoglobin readily associates
and dissociates with oxygen and carbon dioxide and is responsible for the red blood cell's ability to
transport these gases. In most animals, the mature erythrocytes are flexible biconcave discs that lack
nuclei; there are no organelles, ribosomes, or mitochondria. In all other vertebrates erythrocytes are
ovaloid and nucleated. The characteristic shape of erythrocytes provides (1) a high surface area to
volume ratio, (2) minimal diffusion distance, and (3) allows for a greater osmotic water intake without
threatening the integrity of the cell membrane. The shape is maintained by molecular structure of
hemoglobin and by special contractile proteins of the cell membrane.
There are some variations in shape and more in size of erythrocytes among mammalian species. The
dog‟s red blood cells have a typical biconcave shape, whereas in the goat they are more spherical, elliptic
in the camel and somewhat sickle-shaped in the deer. The size of red blood cells in mammals of the
same zoological family is related to a body size but such correlation is easily overruled by environmental
factors. Erythrocytes in species living at high altitudes are small, whereas in diving mammals they are
relatively thick, serving as a slow-release oxygen store. The normal packed cell volume (PCV, hematocrit)
is relatively constant in mammals and birds, since animals with small red cells have high cell counts and
vice versa. Normally red cells size is quite uniform in most healthy individuals of the same species aside
from minor age-related differences.
The total circulating blood volume accounts for about 8 % of the lean body weight. In a 70-kg (154-lb)
adult human about 30 trillion RBCs circulate in 5.6 L of blood, approximately 5 million per µl. Each human
RBC contains about 280 million Hb molecules, has a diameter of 7.4 µm, a volume of 87 µm 3, and a
thickness of 2 µm at the edges and 1µm in the center. The oxygen-carrying capacity of normal human
blood is approximately 20 ml/dL (20 milliliters of oxygen per deciliter of whole blood). In humans, a
reduction in the oxygen-carrying capacity of the blood below 13.5 ml/dL is a cardinal sign of anemia.
Hematology 7.2
A condition in which the oxygen supply of the tissue is inadequate to the body‟s needs is called anemia.
Anemia results when red blood cells are diminished in number, are deficient in Hb, or both. General
causes of anemia are:
Loss of blood due to hemorrhage or internal bleeding.
Decreased production of hemoglobin or red blood cells (as a result of iron deficiency, vitamin
B12, or folic acid, deficiency, bone marrow disease, infection).
Increased rate of RBC destruction (as a result of hemolytic disease, e.g. sickle cell anemia).
In the veal industry, anemia is deliberately produced to create pale muscles. By the age of 8-10 weeks,
commercial veal calves fed wholly on an anemia-inducing milk diet show reduced hematological values
that fall into the pathological range (Table 1).
Hematology is the study of the physiology of blood. It uses laboratory procedures to evaluate blood and
blood-forming organs. The following blood tests will be carried out in this lab: RBC (red blood cell) count,
hematocrit (HCr, or packed cell volume), hemoglobin concentration, as well as MCV (mean corpuscular
volume), MCH (mean corpuscular hemoglobin) and MCHC mean corpuscular hemoglobin concentration)
determination.
PROCEDURE
1) Students will observe prepared slides of the blood smears of different animals.
2) Students will work in pairs with blood sample from one (of two available) species.
3) Students will use Microsoft Excel ™ for simple statistical analysis of the data.
- OBSERVE STANDARD HYGIENIC PRACTICES AND AVOID DIRECT CONTACT WITH BLOOD
- DISPOSE OF SOLUTIONS AND CONTAINERS WITH BLOOD IN MARKED DISPOSAL UNITS
A. RED BLOOD CELL COUNT
A decreased number of circulating erythrocytes is called erythrocytopenia, a condition which is
commonly found in association with many types of anemia. On the other hand, an abnormal increase in
the number of RBCs is called polycythemia.
Because the number of RBCs is very high, blood is diluted (typically 1:200) with an isotonic solution
(Hayem's solution) before the number of cells is counted, and this number is then corrected by the dilution
factor to give the RBC count.
A hemocytometer (blood cell counting chamber) is a finely-ruled glass slide which is used to determine
the numbers of red and white blood cells or platelets in a specific volume of a dilute blood sample. The
grid-covered platforms on the hemocytometer are depressed 0.1 mm below the slide surface so that
when a special coverslip is in place, a chamber 0.1 mm deep is formed. The central portion of the grid
consists of 25 squares (total area=1 mm 2) each of which is further subdivided into 16 smaller squares
(Fig. 1). Five of these 25 squares (R1-R5) are used for RBC counts. To avoid counting a cell twice, only
RBCs which touch a grid line at the left (but not the right) and at the top (but not the bottom) of a square
are considered. Each of the smallest squares occupies an area of 1/20 X 1/20 mm (= 1/400 mm 2).
Because the depth of the chamber is exactly 1/10 mm, the volume of one of these squares is 1/4000
mm3.
Hematology 7.3
To calculate the number of RBCs in a mm 3 of blood, this formula is used:
RBCs/mm3 of blood = Total number of RBCs counted in R1-R5 X dilution X 4000
Number of small squares counted
If the blood dilution is 1:200 and if all of the RBCs in the 16 small squares within each of the 5 R squares
are counted, the formula is simplified to:
RBCs/mm3 = Number of RBCs counted x 200 x 4000 = Number of RBCs counted X 10,000
5 x 16
To convert the RBC count per mm 3 to SI units (viz. cells per liter), multiply RBC count per mm 3 by 106 (1
liter = 106 mm3). Thus if the total number of RBCs counted in the 5 x 16 (=80) small squares is 500, then
the RBCs/mm3 = 500 x 10,000 = 5 million = 5 x 106 and, therefore, RBCs/L = 5 x 106 x 106 = 5 x 1012/L.
Familiarize yourself with the hemocytometer grid by placing a hemocytometer under a microscope set on
the lowest objective (4X on Olympus). Locate the grid and find the 5R cells you will use for the RBC
count. Note that the outer edges of 1-4R cells have double or triple rules; the inner rule identifies the
edge of the cell for counting.
HEMOCYTOMETERS ARE EXPENSIVE. HANDLE VERY CAREFULLY.
RBC Count Procedure.
One method for collecting a dilute blood sample for the purpose of counting RBCs involves the use of a
Becton & Dickinson 5851 Unopette. This device consists of a reservoir which contains a precise volume
of Hayem's isotonic diluent, and a pipette which is used to collect a specific quantity of the blood sample.
Blood drawn into the pipette is expelled into the diluent-containing reservoir, such that a dilution factor of
1:200 is obtained.
Hematology 7.4
1) Label a Unopette reservoir with species name and your bench number. Puncture the top of the
reservoir with the pipette shield.
2) Place a drop of animal blood on a microscope slide.
3) Remove the shield from the Unopette pipette, and touch the tip of the pipette to the drop of blood.
Allow the pipette to fill by capillary action (volume = 10 µl). There must be no air bubbles.
4) Wipe excess sample from outside the capillary pipette, but do not soak up any sample from the
capillary bore. Place your index finger over the opening of the overflow chamber on the capillary
pipette.
5) Squeeze the reservoir and, while squeezing, insert the pipette with its content of blood. When
you release pressure on the reservoir, the blood will be expelled into the premeasured diluent in
the reservoir.
6) Gently squeeze reservoir two or three times to rinse capillary bore. Do not squeeze fluid through
the overflow chamber.
7) Mix the blood and the diluent for about one minute by covering the overflow chamber opening
and inverting several times.
8) Place a cover slip on the hemocytometer so that it covers the grid.
9) Convert the Unopette to a dropper. To do this, remove the pipette from the reservoir, turn it
around, and reinsert into the reservoir.
10) Discard the first 3 drops of blood from the Unopette. Place the next drop of diluted blood at the
edge of the cover slip. The diluted blood will be drawn underneath the cover slip by capillary
action. Wait 3 minutes for the RBC to settle. Do not move the cover slip once the chamber has
been charged.
11) Cover the Unopette with the capillary shield to store the sample until you are ready to discard it.
12) Locate the grid on the hemocytometer. Change to 40X, and count the total number of red blood
cells in each of the 16 small squares within each of the five R squares.
13) Thoroughly and carefully wash and dry hemocytometer and cover slip.
Calculate the number of RBCs/L of blood by using the formula presented above.
B. HEMATOCRIT (PACKED CELL VOLUME, PCV)
Upon the centrifugation of whole blood, its heavier elements (like red and white blood cells) settle, while
the lighter elements form the supernatant fraction. Red blood cells are the most abundant of the blood
cells and account for the majority of the packed cell volume. White blood cells and blood platelets form a
thin pale-colored buffy coat at the interface between the packed red blood cells and the plasma. The
supernatant portion of the centrifuged blood is the plasma, a straw-coloured liquid consisting of water
and dissolved solutes such as sodium ions, hormones (e.g. testosterone, insulin), enzymes (e.g. creatine
phosphokinase), organic molecules (e.g. glucose), and plasma proteins (e.g. albumins). The percentage
of the total blood volume occupied by the packed blood cells is called the hematocrit (HCt). Because
approximately 4% of the plasma remains trapped between the packed cells, to calculate a corrected
hematocrit, the value of the apparent hematocrit must be multiplied by 0.96.
Hematology 7.5
Hematocrit Procedure.
1) Obtain two heparinized capillary tubes (heparin is an anticoagulant). Notice that one end of the
tube is marked with a red band. Touch this end of the capillary tube to the drop of blood, allowing
blood to enter the tube by capillary action until it is at least two-thirds full. Air bubbles are not
important.
2) Place a finger over the other end of the capillary tube and withdraw tube. Using clay capillary
sealant, seal the blood-filled end of the tube by gently pushing it into the Critoseal, twisting the tube,
and then removing it. Repeat steps 1 and 2 for the second tube.
3) Place the sealed capillary tubes in numbered slots on the microcapillary centrifuge, with the
plugged end of the tubes facing outward and against the rubber gasket. Note the number of the
slots containing your samples. Screw the top plate onto the centrifuge head and centrifuge for 5
minutes.
4) Determine the hematocrit with the hematocrit reader. Do this by placing the centrifuged capillary
tube on the HCt scale so the bottom of the red cell layer rests on the zero line and the top of the
plasma layer is even with the 100% line. Find the line which intersects the top of the RBC layer.
The value that this line corresponds to is the apparent (uncorrected) HCt value. Calculate the
corrected HCt (= 0.96 x apparent HCt). Report an average of the two readings for both uncorrected
and corrected HCt.
5) Discard the capillary tubes in the appropriate container.
C. HEMOGLOBIN CONCENTRATION
Because hemoglobin absorbs light, its concentration can be measured in a hemolysed sample by
measuring the intensity of its color. Red blood cells contain different types of hemoglobin, each of which
absorbs light in a different region of the spectrum. Hb which has not bound oxygen is called
deoxyhemoglobin. Upon oxygenation, it forms oxyhemoglobin. A less common form of hemoglobin is
called carboxyhemoglobin; these molecules of hemoglobin have carbon monoxide bound to them.
Hemoglobin may also be found in its oxidized form with the Fe ion in the Fe 3+ rather than the Fe2+ state.
This type of Hb is called methemoglobin.
Hemoglobin concentration is determined by converting all of the various forms of Hb to
cyanmethemoglobin and determining Hb concentration photometrically with reference to a standard
curve. When a sample of blood is added to an alkaline potassium ferricyanide solution, hemoglobin and
its derivatives (except sulfhemoglobin which normally occurs in minute concentrations and can be
ignored) are oxidized to methemoglobin. The latter then reacts with potassium cyanide to form
cyanmethemoglobin, a stable Hb derivative which has a maximum absorption at 540 nm. The
concentration of Hb in unknown blood samples can be determined spectrophotometrically by interpolation
from a standard curve.
Hematology 7.6
Hemoglobin Concentration Procedure.
Turn on the Spectronic 20 and allow it to warm up for at least 15 minutes.
Set the spectrophotometer at 540 nm.
Zero the spectrophotometer so that the needle on the transmittance scale is set to 0%. There should be
nothing in the sample compartment and the lid should be closed.
1) In this portion of the lab you will be working with Drabkin's cyanomethemoglobin reagent, a
buffered potassium ferricyanide-potassium cyanide solution. AVOID CONTACT OF REAGENT
WITH SKIN OR CLOTHES. IF CONTACT OCCURS, FLUSH COPIOUSLY WITH WATER. DO
NOT SWALLOW OR INHALE REAGENT OR FUMES. Label a test tube with your name.
Transfer 5.0 ml of the Drabkin's reagent to the tube.
2) Touch the tip of a 20 µl capillary tube to the drop of blood. Allow the tube to fill by capillary action
to the 20 µl mark indicated on the tube. There must be no air bubbles.
3) Wipe excess sample from outside the capillary pipette, but do not soak up any sample from the
capillary bore. Place your index finger over the opposite end of the tube.
4) With the pipette bulb, flush the blood sample from the tube into your pre-labeled test tube of
Drabkin's reagent.
5) Squeeze and release the bulb two or three times to rinse the capillary bore. Be careful not to
squeeze fluid into the bulb.
6) Mix the blood and reagent by covering the test tube with parafilm and inverting several times.
Allow it to stand at room temperature for 10 minutes.
7) To calibrate the spectrophotometer prepare the blank, by dispensing 5.0 ml of Drabkin's reagent
into a cuvette (one standard per bench). Ensure that the cuvette is clean and dry before placing it
®
in the sample chamber (use a Kimwipe ).
8) Open the sample chamber on the left side of the machine and insert the blank. Close the cover
and adjust the needle to 0 Absorbance using the dial on the lower right of the machine.
9) Transfer your blood sample to a cuvette and read absorbance.
10) Determine the hemoglobin concentration of the blood (g/dL) from the standard curve provided by
your instructor.
The mean corpuscular volume (MCV), mean corpuscular hemoglobin concentration (MCHC), and
the mean corpuscular hemoglobin (MCH) are three parameters derived from the values for corrected
HCt, Hb concentration, and RBC count.
MCHC = hemoglobin (in g/dL) X 100 = # grams per deciliter (of RBCs)
hematocrit (in %)
MCH =
hemoglobin (in g/dL) X 10 = # grams (per cell)
RBC count (in cells/L)
MCH is usually expressed in pg where 1 picogram = 10-12g.
MCV =
Hematocrit (as decimal fraction) = # liters
RBC count (in cells/L)
MCV is usually expressed in fl where 1 femtoliter = 10 -15 L.
Hematology 7.7
Table 1. Normal Hematological Values for humans, sheep, and cattle.
Value
RBC
12
(x 10 /L)
PCV or HCt
Hb (g/dL)
MCV (fL)
MCH (pg)
MCHC (g/dL)
Human
Sheep
Cattle
4.3-5.5
9.0-15.0
5.0-9.0
8.4
6.7
0.37-0.48
12-16
0.26-0.42
8-16
0.24-0.40
8-14
40-60
11-17
30-36
0.33
10.9
39.4
13.0
33.0
0.24
7.4
35.4
11.1
31.3
Male
Female
4.5-6.0
0.45-0.52
14-18
80-94
27-32
32-36
28-40
8-12
31-38
Calves (3 months)
Normal
Veal
Sources:
Doxey, D.L. 1977. Haematology of the ox. pp. 215-269. In Comparative clinical haematology. (Eds.
R.K. Archer and L..B. Jeffcott). Blackwell, Oxford.
Greenwood, B. 1977. Haematology of the sheep and goat. pp. 305-344. In Comparative clinical
haematology. (Eds. R.K. Archer and L..B. Jeffcott). Blackwell, Oxford.
Potter, P.A. 1986. Pocket Nurse Guide to Physical Assessment. Mosby.
LAB REPORT – HEMATOLOGY OF ANIMAL BLOOD
Abstract
Write a short lab report following the style indicated in the Scientific Report Writing section of this manual.
In your lab report, include:
A title page with an appropriate title for your paper plus the usual identifying information.
An abstract (250-300 words) that summarizes the experiment, the key findings, and the important
points of your discussion. In proper format make reference to the results of your statistical
(Student's t) tests. Make sure to include both measured and calculated hematological
parameters.
References
Archer, R.K. and L.B. Jeffcott (eds.). 1977. Comparative clinical haematology. Blackwell, Oxford.
Harmening, D. M. (ed.). 1991. Clinical Hematology and Fundamentals of Hemostasis, 2
Company, Philadelphia.
nd
ed. F. A. Davis
Renal Function 8.1
BIOLOGY 3420
ANIMAL PHYSIOLOGY
FALL 2011 LAB #8
RENAL FUNCTION
Objectives:
As a result of attending this lab, students should be able to:
1. Describe the roles of ADH and aldosterone in the regulation of fluid and electrolyte balance.
2. Explain how the kidneys respond to water and salt loading.
3. Explain how homeostasis of blood plasma osmolarity is achieved and why homeostasis is
important (think about the osmotic fragility lab).
TERMINOLOGY: ADH, aldosterone, renin, glomerular filtration rate, filter, reabsorb, secrete, osmolarity,
glomerulus, Bowman‟s capsule, proximal convoluted tubule, Loop of Henle, distal convoluted tubule,
collecting ducts, concentration, dilution, specific gravity, glycosurea.
INTRODUCTION
Advance reading: Moyes and Schulte pp. 497-514 (2
nd
ed.).
The urinary system functions to keep the body in homeostasis. Drinking excess water or eating salty food
results in a rising blood volume. To maintain homeostasis of the blood, the kidneys adjust the urine
volume, solute concentration, and electrolyte content by removing and restoring appropriate amounts of
water and various solutes. The kidneys also excrete various waste products, regulate the concentration of
+
+
electrolytes (Na , K , HCO3 and other ions) in the plasma, help control blood pH, help regulate blood
pressure by secreting rennin (which activates the renin-angiotensin pathway), take part in
erythropoiesis by secreting erythropoietin, participate in the synthesis of vitamin D, and perform
gluconeogenesis during periods of fasting.
The heart pumps 7000 litres of blood per day, and almost 1/4 goes through kidneys. The kidneys of adult
humans filter about 180 litres of plasma per day. These 180 filtered litres are concentrated into 1-2 litres
of urine daily. How is this volume reduced? Most of the water in the filtered plasma is reabsorbed.
Additionally, substances such as glucose are reabsorbed so that normally none of the filtered glucose
remains in the final urine. However, in people with diabetes mellitus, urinary glucose exceeds 0.03 mg/dL
of urine. Excretion of abnormal amounts of glucose in urine is termed glycosurea. The renal threshold
for glucose is about 8.9 mmol/L (160 mg/dL) of blood; glycosurea indicates that the blood levels of
glucose exceed this amount and the kidneys are unable to reabsorb 100% of the glucose that is filtered.
About 99% of filtered sodium and 50% of filtered urea are reabsorbed. The kidney tubules are also able
to secrete substances such as bicarbonate ions, hepatic metabolic end-products (e.g. hippuric acid), and
foreign substances (e.g. penicillin). Thus the amount of a given substance in the final urine depends not
only on how much was filtered by the glomerulus, but also how much was subsequently removed by
tubular reabsorption and how much was added by tubular secretion. The reabsorption of fluid and
electrolytes (ions) into the blood from the renal tubular filtrate is regulated by hormones. The major
hormones involved in this process are antidiuretic hormone (ADH), released by the posterior pituitary
gland, and aldosterone, secreted by the cortex of the adrenal gland. ADH secretion depends on blood
pressure, blood volume and plasma osmolarity.
+
Control of plasma Na is important in the regulation of blood volume and pressure and also related to
+
renal control of acid-base balance of the blood. When Na is reabsorbed by the renal tubules, Cl follows
passively by electrostatic attraction, and water follows sodium by osmosis. Thus, changes in
Renal Function 8.2
+
blood Na cause secondary changes in blood volume and are critical to health. The final composition of
the urine reflects the integrity of kidney function and changes in blood composition. An analysis of urine
can yield valuable information about the general health of the body. The volume of urine produced and its
specific gravity give information on the state of hydration (or dehydration). The pH of plasma is normally
kept within a narrow range: values of less then 7 or greater then 7.8 are incompatible with continued life.
Acidosis results when the blood pH drops below 7.4: alkalosis is the condition resulting from an increase
in pH above this value. The pH of the urine slightly acidic; however, the pH can be lowered by a diet rich
in proteins or citrus fruits. Thus, urine pH by itself is not very informative. An abnormally low pH coupled
with high glucose and ketones are characteristics of diabetes mellitus. Blood and proteins in the urine
may indicate nephritis, a disease characterized by damaged glomeruli that allow plasma proteins and
erythrocytes to leak into the urine. Several diseases are characterized by abnormal metabolism and the
abnormal byproducts can be found in the urine. Recent advances in urinalysis techniques have made it
possible to perform, in a few seconds, tests which formerly took hours. The Multistix test is a combined
test of pH, proteins, glucose, ketones and occult blood**.
**Occult blood – blood that is present in amounts too small to be seen and can be detected only by
chemical analysis or microscopic examination.
The specific gravity of urine is the weight of a volume of urine divided by the weight of the same volume
of distilled water. Urine weight per volume is higher than distilled water because of the presence of
solutes in urine. Pure water should have a specific gravity of 1.000. The normal Sp. Gravity of urine varies
from 1.001 to 1.025, depending on the state of hydration and the time of the day. It may occasionally rise
to even 1.035.
PROCEDURE
In this lab, we will again make use of the most convenient experimental animals to examine renal
responses to an oral intake of various solutions. Students will work in groups of four, each acting as
subject for one of the experimental treatments.
Pre-Lab preparation
All students should refrain from drinking alcoholic beverages or excessive quantities of
caffeinated beverages during the 24 h preceding the lab.
1) Do not drink anything but water (as much as you wish) during the 4 hour period before lab.
DO NOT DRINK ANY FLUIDS DURING THE TWO HOURS BEFORE THE LAB.
2) Urinate one hour before coming to lab. Note the exact time. Do not urinate again until the first
collection at the beginning of the lab.
Lab Procedures
Each student will be assigned to drink one of the following solutions:
Subject 1
- Distilled water
- 16 ml per kg of body weight
Subject 2
- Distilled water
- 7.5 ml per kg of body weight
Subject 3
- Cola
- 7.5 ml per kg of body weight
Subject 4
- Sprite
- 7.5 ml per kg of body weights
Renal Function 8.3
If there are ungrouped (and brave) students in the lab they may be assigned to drink another solution,
such as 0.5% NaHCO3 (7.5 ml per kg of body weight), or 20% sucrose (7.5 ml per kg of body weight), or
0.9% NaCl (7.5 ml per kg of body weight), or 2% NaCl (3 ml per kg of body weight).
The sodium chloride solutions are rather difficult to drink (especially 2% one); if any signs of nausea or
cramping begin to occur, discontinue drinking the solution. Be sure to determine the exact volume of
solution drunk by each subject.
Subjects with normal kidney function will be required for the experiment. If you have circulatory problems,
poor kidney function or have any medical problems, do not volunteer as a subject for this experiment.
For each urine sample, you will collect data allowing you to determine the rate of urine formation,
the concentration of urinary solids, and the pH of the urine.
Analyze only your own urine!
Wear gloves.
1) Obtain beakers/cups for drinking and for urine collection. Prepare the assigned test solution.
Then drink it, as fast as possible.
2) Collect the first urine sample right after the drinking of fluids (i.e. 0 min - control). Note the exact
time.
3) Measure the volume of urine sample with a measuring cylinder.
4) Save a small amount (50 ml?) of urine sample in a labeled beaker/cup and discard the rest.
5) After 60 (or 90) minutes period each subject should again empty his or her bladder, collecting the
full sample. Note the exact time and volume of urine produced. Record your results in the
appropriate table in the Lab Book.
Note: If you need to urinate more frequently, keep a record of the total volume of urine
collected during 60 minutes interval.
6) Determine urine specific gravity (see below).
7) Calculate urine flow rate (V) in ml urine per minute.
8) Test both urine samples for pH and some of the clinically important constituents (see below).
SPECIFIC GRAVITY
Specific gravity is usually a good index of urine osmolarity, with a specific gravity of 1.008 approximating
the normal plasma osmolarity of 290 mosm/kg.
The specific gravity is greatly influenced by the temperature. Hence, it is best to make all examinations at
current room temperature.
Determine the specific gravity of the urine samples by floating a urinometer in a cylinder nearly
filled (less than an inch from the top) with the specimen.
Gently spin the device so that it rotates freely for at least two turns.
Read the specific gravity at the meniscus on the urinometer scale, making sure that the
urinometer float is not touching the bottom or the sides of the cylinder.
Renal Function 8.4
URINALYSIS
The urine normally has a pH between 5.0 and 7.5.
If the urine is allowed to stand for a few hours, the pH will change to a basic value as a result of the
decomposition of urea to ammonia.
Obtain a Multistix from the side bench.
Dip the test area of the strip in the urine and remove it immediately.
Wipe off the excess urine by dragging the test strip across the lip of the beaker.
The specified number of seconds read the strip by comparing it to the Multistix color chart
provided.
References
Vander, A. J. 1985. Renal physiology. McGraw-Hill. New York.
LAB ASSIGNMENT – RENAL FUNCTION
Assignment will be handed out at the beginning of the lab period.
Although you will be working in groups to obtain results, assignments must be completed independently.
Due Date: November 28-29
Metabolism 9.1
BIOLOGY 3420
ANIMAL PHYSIOLOGY
FALL 2011
LAB #9
METABOLISM
Objectives:
As a result of attending this lab, students should be able to:
1) Explain the concepts of metabolism and metabolic rate.
2) Learn and practice a method for indirectly measuring metabolic rate.
3) Comment on various factors affecting metabolic rate.
4) Make the calculations necessary to determine RER and RQ.
5) Understand the difference between basal and standard metabolic rates.
TERMINOLOGY: metabolic rate, indirect calorimetry, basal metabolic rate, standard metabolic rate,
respiratory exchange ratio, respiratory quotient, specific oxygen consumption, minute ventilation
INTRODUCTION
Advance reading: Moyes and Schulte pp. 60-64; 630-637 (2
nd
ed.).
Metabolic rate
Metabolism is the sum of all chemical reactions that occur within a cell or organism. Metabolism is
generally reported as either work (how much energy did an animal use) or more commonly as power
(how fast did an animal use energy or what was the rate of its chemical reactions). Metabolic rate refers
to energy expended by the body per unit time. The metabolic rate of an animal is affected by the ambient
temperature, the time of day and year, its diet, age, level of activity, and many other factors. The ingestion
of mix-diet meal may increase metabolism by10-20 percent and the ingestion of protein alone - up to 30
percent. The greatest changes in metabolic rate occur as a result of muscular activity. In this experiment
you will determine the metabolic rates of different animals using the respirometry system. Metabolic rate
can be calculated from measurements of an animal‟s oxygen consumption under a given set of conditions
(indirect calorimetry). It is valid to determine an animal‟s basic metabolic rate by indirect calorimetry if:
The animal‟s diet is known.
There is no anaerobic metabolism and ATP and creatine phosphate stores are maintained.
The change in body oxygen stores over the measurement period is minimal.
The animal is at rest and under no thermal stress.
In mammals and birds, the stable, fasting, minimum rate of metabolism is called the basal metabolic
rate (BMR). BMR is the rate at which the organism releases heat as a result of breaking down fuel
molecules; the organism‟s basic cost of living. The metabolic rate of animals other than mammals and
birds is largely dependent on the environment so there is no metabolic rate that can be called „basal‟ for
these animals. Instead, the minimum metabolism of fasting animals at a given external temperature is
called the standard metabolic rate for that temperature.
The amount of heat produced for each liter of oxygen consumed in metabolism is relatively constant,
whether carbohydrate, fat or protein is being oxidized. The values are 4.5 KCal/L O 2, 4.7 KCal/L O2, and
5 KCal/L O2 for protein, fat, and carbohydrate respectively. The average value of 4.8 KCal/L O 2 is
generally used in calculating metabolic rate.
Metabolism 9.2
Respiratory quotient
The respiratory exchange ratio (RER) is the number of CO2 molecules produced relative to the number
of oxygen molecules consumed by intermediary metabolism. In steady state at rest, this is equivalent to
the respiratory quotient (RQ).
(%CO2 expired - %CO2 inspired ) / (%O2 inspired - %O2 expired)
The RQ value is directly related to the animal‟s diet. In metabolizing carbohydrate alone, one CO 2
molecule is produced for every O2 molecule consumed, for an RQ of 1.0. In metabolizing fat alone, 0.7 of
a CO2 molecule is produced for every O2 molecule consumed, for an RQ of 0.7. In metabolizing protein
alone, approximately 0.81 of a CO2 molecule is produced for every O2 molecule consumed, for an RQ of
0.81. At rest after a period of fasting the RQ will be closer to 0.7 because the animal will be metabolizing
stored fat. During exercise, the RER will increase with the level of exercise intensity. This is due to a
progressive shift from fat to glucose metabolism.
The effectof size on the metabolic rate of an organism.
Organisms vary in size by many orders of magnitude (an elephant and a mouse). The influence of size on
an organism's metabolism is relatively easy to envision: bigger organisms should have greater total
metabolisms since they have more tissue. However, the relationship is not linear. Given equivalent
conditions of diet and activity, if metabolic rate is considered on a „per unit mass‟ basis, smaller animals
always have higher metabolic rates than larger animals. In general, for each doubling in body weight,
standard metabolic rate increases by about 75%. For a wide variety of animals, it has been found that
0.75
metabolic rate is proportional to (body weight)
. This is known as the “Brody-Kleiber” relationship. In
mammals and birds, this relationship may be partly due to smaller animals having larger surface areas
relative to their body weights. Since the rate of heat loss is proportional to surface area, small mammals
and birds must produce a greater amount of heat per unit of body weight in order to maintain a constant,
warm body temperature. For other types of animals, the association between metabolic rate and body
weight may be due to the relationship between the rate of oxygen consumption per unit body mass
(specific oxygen consumption) and body weight. For diverse types of animals, it has been shown that
the log of the specific oxygen consumption is inversely proportional to the log of body mass.
The effect of temperature on the metabolic rate of endotherms and ectotherms
The body temperatures of ectotherms, such as the frog and other amphibians, are largely determined by
the temperature of their surroundings. Ectotherms rely almost entirely on environmental sources of heat.
The metabolic rate of an ectotherm is therefore linked closely to the external temperature. In fact, the
relationship between metabolic rate and external temperature can be almost linear within a wide range of
temperatures. Within these limits, the metabolic range of an ectotherm generally doubles for every 10ºC
increase in external temperature. This exponential increase in metabolic rate can be attributed to
increased enzyme activity. The reactions of both anaerobic and aerobic cellular respiration are catalyzed
(facilitated) by enzymes and are extremely sensitive to small changes in temperature.
Endotherms, such as birds and mammals, regulate their own body temperatures. In most mammals, the
normal physiological range for core body temperature is from 37ºC -38ºC. For birds, core body
temperature is closer to 40ºC. Endotherms regulate temperature through a high rate of endogenous heat
production and by controlling the rate of heat exchange. Because of the endogenous heat production, an
endotherm usually has a metabolic rate at least five times that of an ectotherm of equal size and body
temperature.
An endotherm can maintain a constant body temperature within a particular range of external
temperatures called the thermal neutral zone. In this zone, external temperature has very little effect on
Metabolism 9.3
an endotherm‟s metabolic rate because it is able to compensate for changes in ambient temperatures by
varying its thermal conductance. It can do this by changing the supply of blood to superficial areas,
increasing or decreasing the degree of insulation afforded by fur or feathers, or making changes in body
orientation
In this lab, you will determine the effect of size on the metabolic rate of an organism and examine the
effect of external temperature on the metabolic rate of a different type of animals. The calculated
metabolic rates at the three test temperatures should show the classic endotherm and ectotherm
responses.
Procedures
Students will work in group of four with one kind of animal per bench. Class data will be combined.
Gas exchange system set up
Arrange the apparatus in the order shown in the picture below.
Assemble the animal chamber. Insert the temperature probe into the port below the fan.
Assemble a condensing ice bath.
Connect the outflow tubing from the chamber to the ice bath inflow port.
Make sure the gas analyzer is warmed-up, and that a filter and Naflon sampling tube have
been attached to the inlet port.
The GA-200 gas analyzer is calibrated at the factory and is ready to take
measurements.
Connect the source of the gas sample (the ice bath outflow port) to the gas analyzer with the
proper connections and press the RUN key.
Ensure that the temperature probe and the gas analyzer CO2 and O2 ports are connected to the
Lab Pro interface.
A. MEASUREMENT OF METABOLIC RATE
1. Start data collection in Logger Pro. (Click on „Collect‟.) The graphs displayed by Logger Pro on
the computer screen should show the temperature signal in degrees Celsius (upper graph), CO 2
concentration in % (middle graph) and O2 concentration in % (lower graph).
Metabolism 9.4
2. Monitor the temperature, CO2 and O2 values until they stabilize. (This should take at least two
minutes.) Record the temperature in Table 1. Record the O2 and CO2 values in Table 2.
Note: Once the recordings reach a steady level, record for another ten seconds.
3. Place the animal in the chamber and ensure that the pump is plugged in. Warning: Do not leave
any animal in the chamber without plugging in the pump! Choose the chamber according to
the animal size, otherwise it may take a very long time for the CO 2 and O2 levels to stabilize.
4. Monitor the temperature, CO2 and O2 values until they stabilize once more. (This should take
from five to twenty minutes.) Record the stable temperature attained in Table 1. Record the O 2
and CO2 values in Table 2.
5. Record another sample of reference air (bypassing animal chamber).
6. Stop data collection by clicking on the „Stop‟ icon at the top of the screen. DO NOT use the quick
save icon. You will write over the set-up file! Instead, save your data by clicking on „File‟ and
„Save As‟ in the main menu.
7. Turn off the pump, remove the animal from the chamber and weigh it.
B. THE EFFECT OF TEMPERATURE ON METABOLIC RATE OF ENDOTERMS AND ECTOTHERMS
1. Fill the large, shallow tray with crushed ice. Place the tray under the chamber and pack the ice
around it to a height of three centimeters. Fill a large zip-lock bag with crushed ice. Drape the
bag over the top of the chamber.
Repeat the experiment described in part A.
Monitor the temperature, CO2 and O2 values. The temperature should decline gradually with
time and you should make some notes about the effect of the colder temperature on the
observed O2 and CO2 readings. How long did it take for the O2 and CO2 values to stabilize?
(Response time.) (If you are testing a mouse, be prepared to wait several minutes for it to
react to the colder temperature.) Is there any change in the animal‟s appearance or
behavior? It is important to record any increase in activity level as this would result in a
higher metabolic rate. If the animal is ectothermic, this might minimize or mask the expected
reduction in the animal‟s metabolic rate which should result from the lower ambient
temperature. If the animal is endothermic, any rise in metabolic rate would have to be
attributed partly to the animal‟s increased level of activity.
Record the stable low temperature attained in Table 1. Record the O2 and CO2 values in Table
2. Remove the tray of ice and the ice bag.
2. Turn on the chamber-heating block. Heat the chamber until an internal temperature of no greater
than 38°C is achieved. If the temperature rises above 38°C, turn the heating block off until the
temperature starts to fall.
CAUTION: The material that the chamber is made from retains heat well. Do not allow the internal
temperature of the chamber to rise above 40°C!
Monitor the temperature, CO2 and O2 values. Make some notes about the effect of the hotter
temperature on the observed O2 and CO2 readings. How long does it take for the O2
Metabolism 9.5
and CO2 values to stabilize? (Response time.) Is there any change in the animal‟s
appearance or behaviour? It is important to record any increase in activity level as this would
result in a higher metabolic rate.
Record the highest stable temperature attained in Table 1. Record the O 2 and CO2 values at
this temperature in Table 2.
Note: Before storing the GA-200 connect an external filter to the inlet port of the unit to prevent
contaminations of the sensors.
Data Analysis
1. Refer to Table 2. Determine the change in CO2 and O2 concentrations (Δ %CO2 and Δ %O2) the
air in the bag (inflow) to the outflow from the chamber.
Record your results in Table 3.
CO2 = outflow CO2 - inflow CO2
O2 = inflow O2 - outflow O2
2. Convert the ΔCO2 and ΔO2 values from units of % to units of µL O2 or CO2 per mL of gas.
Remember that 1% O2 or CO2 is equivalent to 10µL/mL.
3. You can use these Δ values together with the measured flow rate through the system to calculate
the oxygen uptake and carbon dioxide output for each animal.
CO2 output (µL/min) = flow rate (mL/min) x ΔCO2 (µL/mL)
O2 uptake (µL/min) = flow rate (mL/min) x ΔO2 (µL/mL)
4. Divide each animal‟s oxygen uptake in µL/min by its weight in grams to obtain its oxygen uptake in
µL per gram per minute. Record your results in Table 4.
5. Calculate the animals‟ Respiratory exchange ratios: RER
= CO2 output
O2 uptake
Record your results in Table 5.
6. The average amount of heat produced for each liter of oxygen consumed in metabolism is about
4.8 KCal or 20.1 KJoules. You can therefore obtain the metabolic rate of each animal in calories
per gram per minute by multiplying its oxygen uptake in µL per gram per minute by 0.0048. To
obtain each animal‟s metabolic rate in Joules per gram per minute, multiply its oxygen uptake by
0.0201 instead. Record your results in Table 6.
References
th
Schmidt-Nielsen, K. 1997. Animal physiology: adaptation and environment. 5 ed. Oxford University
Press, N.Y.
Animal Use 1
BIOLOGY 3420
ANIMAL PHYSIOLOGY
FALL 2011 APPENDIX A
USE OF EXPERIMENTAL ANIMALS
For the acquisition of any knowledge, one must turn to the best source of information. In animal
physiology, that source is the animal itself. A thorough and complete understanding of physiological
principles and their application requires laboratory exercises that use living tissue (nerves and muscles),
body fluids (urine and blood), and whole living animals (species of different phylogenetic levels, including
humans). Such hands-on laboratory exercises provide a valuable learning experience that cannot be
achieved by other means. Of course, this brings up ethical and legal obligations related to the use of
animals. Rules and regulations governing the use of animals in research have been established to ensure
humane approach.
All institutions in Canada that maintain animals for teaching, research, or testing are required to have a
Local Animal Care Committee that approves protocols involving the use of animals. The membership of
such committees includes animal users, non-users from within the institution, a DVM (vet), and an
outsider representing community interests. At regular intervals (at least once every 3 years), each
institution is visited by an external assessment panel which inspects all facilities and animals to ensure
that the institution is adhering to the guidelines for the care and use of experimental animals formulated
by the Canadian Council on Animal Care (CCAC). Institutions found in non-compliance can have their
animal facilities closed and their funding withdrawn.
The CCAC, established in 1968, is funded by the Natural Sciences and Engineering Research Council
(NSERC) and the Medical Research Council (MRC). The Council has about 20 members representing
government agencies that use animals (such as the Department of National Defence), professional
societies (such as the Association of Canadian Medical Colleges), granting agencies (such as the
Canadian Heart Foundation), and the Canadian Federation of Humane Societies. More information about
the CCAC can be found online at http://www.ccac.ca.
The CCAC publishes the "Guide to the Care and Use of Experimental Animals" (vols. 1 & 2) which
explains the philosophy of the CCAC with regard to the use of animals, describes the caging and housing
requirements for captive animals, appropriate methods of anesthesia and euthanasia, and appropriate
surgical techniques. All users of animals should be familiar with these guidelines.
The CCAC states that the use of animals in research, teaching and testing is acceptable only if it
promises to contribute to the understanding of environmental principles or issues; fundamental biological
principles; or development of knowledge that can reasonably be expected to benefit humans, animals or
the environment.
Animals should be used only if the researcher‟s best efforts to find an alternative have failed. A continuing
sharing of knowledge, review of the literature and adherence to the Russell-Burch „Three R‟ tenet of
“Replacement, Reduction and Refinement‟ are also requisites.
Replacement
Use non-animal models wherever possible such as Tissue Culture, Computer Simulations and
Phylogenetic Replacement (use of animal as low on the phylogenetic tree as possible).
Reduction
Animals and animal tissues should be shared, and a statistical design of the experiment be attempted in
order to determine the least number of animals needed to prove the hypothesis.
Refinement
Use of the least invasive technique possible and the most advanced instruments available to collect data
and protect animals is key.
Animal Use 2
To reduce the number of animals used in laboratory part of this course you will perform exercises in small
groups and share your results; where possible, other methods will be used, such as demonstrations,
models, computer simulations, video etc. The university library has excellent learning resources available
to you.
To name a few new arrivals:
Interactive physiology. Cardiovascular system. CD-ROM. QP 101 I57 2000.
Anatomy and physiology. CD-ROM. QP 34.5 A52 2002 (Separate quiz and tutorial modes allow
students to test their mastery of key content).
Whole volumes have been written on the ethics of animal experimentation. If you want to learn more
about the arguments both for and against animal research visit some of the following “links”:
The Principles of Humane Experimental Technique by W.M.S. Russell and R.L. Burch, published
in 1959, this was the first formulation of the 3 R-s for humane animal research.
o http://altweb.jhsph.edu/publications/humane_exp/het-toc.htm
The Northwest Association for Biomedical Research.
o http://www.wabr.org/about/fgg.html
Altweb, the Alternatives to Animal Testing Web Site.
o http://altweb.jhsph.edu/
Visit University Library and check out Rain without thunder: the ideology of the animal rights movement
by Gary L. Francoine for additional information on the discussion animal rights and animal welfare. HV
4764 F73 1996.
Over the course of the semester we will use mostly noninvasive techniques in our experiments; and you
will be required to handle the animals involved. Animals are very sensitive to the mood and actions of
their handler. Be confident and smooth as you pick up the animal. Avoid darting movements and failed
grabs; the animal will become nervous and harder to handle. If you have upset your animal, leave it
alone for a few minutes until it - and you - become calm again.
Qubit Systems 1
BIOLOGY 3420
ANIMAL PHYSIOLOGY
FALL 2011
APPENDIX B
Qubit Systems’ laboratory package
Qubit Systems‟ laboratory package enables students to conduct experiments on the physiology of
animals using computer-based instrumentation that allows acquiring and displaying data in real time. The
components of this package can be used in different combinations to provide specific experimental setups to study human electrophysiology, human respiratory and cardiovascular physiology, as well as to
measure and compare the metabolic rates of different organisms.
The components of the package are described below.
LabPro Interface
The LabPro interface works in conjunction with the Logger Pro software.
LabPro stores 12,000 data points internally and has sampling rates from one reading per day to 50,000
readings per second. This interface allows the computer to record and save the data from sensors and
gas analyzers used in lab experiments.
There are three buttons on top of the LabPro. Do not use the buttons if LabPro is connected to a
computer running Logger Pro. When LabPro is connected to a computer, its operation is controlled via
the Logger Pro software. There are three LEDs on top of the LabPro. The red LED indicates an error
condition. The yellow LED indicates that LabPro is ready to collect data samples and the green LED
indicates that LabPro is collecting data.
To conduct the experiments you should connect LabPro to a computer via an unused modem, printer or
USB port. The serial and USB connections are on the right side of the interface. LabPro can be powered
by AC power adapter by plugging the round plug on the 6V-power supply into the left side of the interface.
If you hear six beeps and observe the LEDs blink, the unit has been powered up successfully.
Connect the sensors to the appropriate channels as specified in the experiment that you are running. In
Qubit‟s experiments, you will be using the analog channels on the left side of the interface.
♦ Use the serial cable to connect LabPro to the modem or printer port of your computer or use the USB
cable to connect LabPro to the USB port of your computer.
♦ Connect the power supply to LabPro via the AC adapter port. Plug in the power supply.
♦ Locate the Logger Pro 2.2 or Logger Pro 3.4.5 file alias on your computer desktop. Double click on it.
If Logger Pro finds LabPro, the main Logger Pro screen will appear. If Logger Pro cannot find an
interface, the “Setup Interface” dialog box will appear. If the interface is connected to the printer port,
select that port on the Setup Interface dialog box and then click the “Scan” button. Once the interface
has been identified, "LabPro" will appear in the interface field of the Setup Interface dialog box. Click
“OK” to proceed to the main screen. If the interface has still not been identified, secure all cable
connections and ensure that the interface has power.
Qubit Systems 2
EMG Sensor
Qubit Systems‟ EMG sensor is a simple device for measuring the electrical activity of muscles. Three
cables terminating in alligator clips are attached to electrodes placed at strategic points on the subject‟s
skin. Two of the electrodes are placed on the skin overlying the muscle to be studied, and the third
electrode (ground) is placed on the skin a short distance from the other two outside the region where the
muscle under study is active (e.g. on the ankle for calf muscle activity). The signals from the EMG sensor
are measured as voltages in Data Logger or Logger Pro software and shown as graphical images plotted
against time. If desired, they may also be heard as audible signals.
The configuration of the front and back panels of the EMG Sensor are shown below:
Front Panel
Back Panel
The black, green and white electrode leads that are attached to the front of the instrument are for the
ground, negative and positive electrodes, respectively. These terminate in alligator clips that are colorcoded in the same way. A potentiometer is provided for controlling the volume of the audible signal, and
a second potentiometer is provided for adjusting the voltage output for a given muscular effort. This is
used during set-up to ensure that maximum effort does not produce a signal greater than 5 V which
cannot be read by the software.
Power for the instrument is provided by a 6 VDC 500 mA positive center power supply, attached to the
back panel, that should be used only with a 120 VAC mains supply.
There are two analog out ports on the back of the instrument. One, labeled “Raw Analog Out” provides
the complete electrical signal from the muscle. When the muscle is excited, positive and negative
potentials occur above and below the muscle‟s background resting potential. The raw analog out signal
sets the background potential to a nominal value close to 2.5 V, so that it is in the middle of the
measurement range of the computer interface. The positive and negative potentials are measured above
and below this value.
The “Rectified Analog Out” port provides a rectified signal in which the background resting potential of the
muscle is set close to zero volts. The negative potentials that occur during muscle activation are rectified
so that all electrical events are shown as positive values. This makes identification of motor unit spikes
easier, and is the preferred signal for educational use.
A switch on the back panel of the instrument allows the speaker to be turned on or off as required. The
cable carrying the analog out signal to the interface terminates in a BT connector for use with the Lab Pro
Interface. If the EMG sensor is used with Qubit Systems customized set up files it must be plugged into
Channel 2 of the Lab Pro Interface.
Qubit Systems 3
Setting EMG Gain with Data Logger Software.
It is important to set the gain of the EMG sensor so that its maximum output for a given muscular
contraction does not exceed the maximum 5 V that the Data Logger software can read. To achieve this,
do the following:
Open an appropriate EMG file.
Attach the electrodes to the skin in the optimum position for the muscle to be studied.
Start data collection and contract the muscle with the greatest possible force while observing the
voltage output on the computer screen.
Adjust the gain potentiometer on the control box to read a voltage below 5 V, but above 4 V so
that there is good resolution of the muscles electrical activity.
The output of the EMG sensor is now set for your experiment. Be careful not to adjust the gain
control again before the experiment is complete.
Use and Positioning of Electrodes
A pack of 100 electrodes are supplied with the sensors. These are attached to cards by conductive gel,
and are ready to use on the skin without further application of electrode gel. The electrodes operate
optimally when the skin is clean and free of grease. For best results, use a piece of fine sandpaper to
abrade the skin lightly at the point where the electrode is to be placed. This is to remove dead skin and
abrasion should not cause reddening or breaking of the skin surface. Use rubbing alcohol on a piece of
cotton wool to clean the area after abrasion, and allow it to dry before applying the electrode.
Press the electrode into place firmly with the tab pointing downwards for most easy attachment of the
alligator clip on the electrode cable. Having attached the alligator clips, ensure that they are unlikely to
fall off or drag the electrode off the skin. This may involve using a piece of surgical tape to hold the
electrode and lead in place.
Breathing Monitor
Wireless Exercise Heart Rate Monitor
The Wireless Exercise Heart Rate Monitor uses a
Polar electrode and transmitter/receiver system to
transfer heart rate data to the Lab Pro Interface.
Taking the length of the receiver cable into
consideration, the subject can be up to five feet from
the interface. This should allow for heart rate
monitoring during most types of exercise.
The Breathing Monitor (BM) belt is worn around
the lower chest and upper abdomen. The belt
contains a rubber inflatable bladder which is
connected to an absolute pressure sensor. The
sensor monitors the change air pressure within
the bladder as the subject breathes in and out.
During inhalation, the diaphragm and intercostal
muscles cause expansion of the rib cage and
plural cavity. This compresses the bladder
within the BM belt, increasing the signal from the
pressure sensor. During exhalation, the rib cage
relaxes, the air bladder decompresses and the
pressure signal declines. The strength of the
signal is proportional to the degree of inflation of
the lungs. The voltage signal is transmitted to
the computer via the interface and is displayed
as “voltage” by Logger Pro software.
BM peak amplitude can be used as a relative
measure of breathing depth between conditions
in the same subject. The relationship between
BM peak amplitude and actual lung volume will
be different for each subject because of body
morphology, initial degree of inflation of the air
bladder, etc.
Qubit Systems 4
Spirometer
The Spirometer is designed to make human respiratory
measurements at rest and during moderate activity. It can
be used to perform a variety of tests related to air flow and
lung volume. The sensor includes a removable flow head for
easy cleaning and sterilization, and a differential pressure
transducer. The Spirometer package includes 1 sensor
handle, 1 flow head, 5 disposable mouthpieces, 1
disposable bacterial filter, and 1 nose clip.
Patellar Reflex Hammer
Electronic percussion hammer for recording
impact stimulation of the patellar tendon.
Gas Analyzer
The GA-200 (iWorx) is an analyzer that integrates a gas sampling system with sensors to measure and
display the concentrations of oxygen and carbon dioxide in a sample as the percentage of a gas in the
sample by volume. This method of expressing the concentration of a gas in a sample is also known as
the percent volume fraction. In addition to the sampling and sensing systems in the GA-200, the unit has
a fluorescent display used for programming the unit and observing measurements, a keypad for
programming the unit‟s operation and calibration, and analog outputs that allow the unit to be connected
to data recording units like LabPro.
Qubit Systems 5
The front and rear panel of the GA-200 gas analyzer with filter.
When measuring the concentrations of oxygen and carbon dioxide in a gas, a sample is pumped from the
input port on the front panel of the GA-200, through the sample cell, and out an exhaust port on the back
panel of the unit. Gas samples are drawn into the unit through an external filter on the input port that
protects the sensors from contamination and through Nafion tubing on the filter that removes moisture
from the samples that would affect the measurements and the unit calibration.
The oxygen concentration of the gas in the sample cell is measured using laser diode absorption
technology. The laser diode in the oxygen sensor produces light at a wavelength (760 nanometers) that is
absorbed by oxygen. The light passes through the gas pumped into the sample cell and onto the surface
of a detector. The output of the sensor is inversely proportional to the concentration of oxygen in the
sample because the amount of light reaching the detector decreases as the concentration of oxygen in
the sample increases. The GA-200 has a very fast response time that enables breath to breath analysis
of gas concentrations because the unit analyzes the gas sample every 10 milliseconds, or 100 times per
second. During each measurement interval, the analyzer is zeroed automatically by electronic tuning of
the laser to a wavelength not absorbed by oxygen.
The GA-200 gas analyzer is calibrated and ready to take measurements. Just connect the source
of the gas sample to the gas analyzer with the proper connections and press the RUN key.
Gas Bags
The gas bags are made from a gas-impermeable nylon-polyethylene laminate and are heat-sealed.
Tygon tubing is attached to each bag by a luer-lok fitting. Once inflated, the bags should be sealed by
capping the luer-lok fitting on the end of the Tygon tubing with the clamp. Bags should not be overinflated as this can cause weakening of the seams and eventual leakage.
Temperature Sensor
The temperature sensor is a semi-conductor device mounted on
the end of a stainless steel tube. It is inherently linear with an
operational range from -40ºC to +125ºC. The calibration for the
temperature sensor is loaded automatically when the software is
activated. The sensor is mounted on a stainless steel support
which fits through the temperature sensor ports in an animal
chamber. The temperature sensor port must be plugged with a
solid rubber stopper when the sensor is not in use
Data Collection Instructions
Graph Display
The system software uses graph(s) on the screen showing output from the sensor(s). You can customize
most of the options of the graph(s) by double clicking on the graph and selecting the options in the dialog
box that is displayed. For example, you can plot any available sensor on any graph. Click on the y-axis of
the graph to show the options available and select the ones that you want to plot. A tick mark will appear
against each parameter that will be plotted. Click against the parameters to add or remove ticks. The
graph title will change depending on the parameters that you select.
Qubit Systems 6
Graph Ranges, Run Time and Data Collection Rate
To modify the ranges of the x- and y-axes, click on the maximum and/or minimum values and type in the
new values. If you wish to alter the maximum time allotted to your experiment or the rate of data
sampling, select “Experiment”, then “Sampling”. Type in the run time of your experiment in the box
labeled “Experiment Length”.
Note that the maximum value you select for the Time axis limits the time over which you can
collect data for a particular run.
You may also alter the rate at which you collect data. The default rate is one point per second. For long
experiments this may be too fast, resulting in very large data files. For shorter experiments with important
transient conditions, you may wish to sample at a faster rate. If your experiment exceeds the allotted time,
select “Data”, then “Store Latest Run”. You may then restart data collection. Data from the first part of
your experiment will remain on the screen as a faint trace and new data will be plotted in bold. You can
collect numerous runs in this way. Each run will be collected to a separate data table. When you save
your data, all of the runs will be saved under the same file name.
Data Collection and File Saving.
To start data collection, click on the “Collect” icon above the graphs. This will change to a “Stop” icon.
Click on the “Stop” icon to stop data collection.
If you wish to save your data and any changes to your Logger Pro set-up, select “File” from the main
menu, then “Save As”. You should save your data files under a new name and store them in an
appropriate location on your hard drive, or on a 3.5” disk. You may open your stored file by selecting
“File” then “Open” from the main menu and navigating to the file via the “Look In” dialog box.